Chapter 1 Cell Culture

Current Protocols in Cell Biology 　 Chapter 1 Cell Culture Introduction PDF Unit 1.1 Basic Techniques for Mammalian Cell Tissue Culture PDF Unit 1.2 Media for Culture of Mammalian Cells PDF Unit 1.3 Aseptic Technique for Cell Culture PDF Unit 1.4 Sterilization and Filtration PDF Unit 1.5 Assessing and Controlling Microbial Contamination in Cell Cultures PDF Unit 1.6 Media and Culture of Yeast PDF Unit 1.7 BY-2 Cells: Culture and Transformation for Live Cell Imaging PDF

Chapter 2 Preparation and Isolation of Cells Introduction PDF Unit 2.1 Establishment of Fibroblast Cultures PDF Unit 2.2 Preparation and Culture of Human Lymphocytes PDF Unit 2.3 Preparation of Endothelial Cells PDF Unit 2.4 Generation of Continuously Growing B Cell Lines by Epstein-Barr Virus Transformation PDF Unit 2.5 Laser Capture Microdissection PDF Unit 2.6 Preparation of Human Epidermal Keratinocyte Cultures PDF

Chapter 3 Subcellular Fractionation and Isolation of Organelles Introduction PDF Unit 3.1 Overview of Cell Fractionation PDF Unit 3.2 Isolation of Rat Hepatocyte Plasma Membrane Sheets and Plasma Membrane Domains PDF Unit 3.3 Isolation of Mitochondria from Tissues and Cells by Differential Centrifugation PDF Unit 3.4 Purification of a Crude Mitochondrial Fraction by Density-Gradient Centrifugation PDF Unit 3.5 Isolation of Peroxisomes from Tissues and Cells by Differential and Density Gradient Centrifugation PDF Unit 3.6 Isolation of Lysosomes from Tissues and Cells by Differential and Density Gradient Centrifugation PDF Unit 3.7 Overview of Subcellular Fractionation Procedures for the Yeast Saccharomyces cerevisiae PDF Unit 3.8 Isolation of Subcellular Fractions from the Yeast Saccharomyces cerevisiae PDF Unit 3.9 Isolation of Golgi Membranes from Tissues and Cells by Differential and Density Gradient Centrifugation PDF Unit 3.10 Isolation of Nuclei and Nuclear Membranes From Animal Tissues PDF Unit 3.11 Free-Flow Electrophoretic Analysis of Endosome Subpopulations of Rat Hepatocytes PDF

Chapter 4 Microscopy Introduction PDF Unit 4.1 Proper Alignment and Adjustment of the Light Microscope PDF Unit 4.2 Fluorescence Microscopy PDF

Unit 4.3 Immunofluorescence Staining PDF Unit 4.4 Fluorescent Staining of Subcellular Organelles: ER, Golgi Complex, and Mitochondria PDF Unit 4.5 Basic Confocal Microscopy PDF Unit 4.6 Immunoperoxidase Methods for Localization of Antigens in Cultured Cells and Tissues PDF Unit 4.7 Cryo-Immunogold Electron Microscopy PDF Unit 4.8 Correlative Video Light/Electron Microscopy PDF Unit 4.9 Polarization Microscopy PDF Unit 4.10 Fluorescent Speckle Microscopy (FSM) of Microtubules and Actin in Living Cells PDF Unit 4.11 Two-Photon Excitation Microscopy for the Study of Living Cells and Tissues PDF Unit 4.12 Total Internal Reflection Fluorescence Microscopy for High-Resolution Imaging of Cell-Surface Events PDF Unit 4.13 Fluorescent Labeling of Yeast PDF Unit 4A Organelle Atlas: Appendix to Chapter 4

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Chapter 5 Characterization of Cellular Proteins Introduction PDF Unit 5.1 Overview of the Physical State of Proteins Within Cells PDF Unit 5.2 Determining the Topology of an Integral Membrane Protein PDF Unit 5.3 Determination of Molecular ) Unit 5.4 Analysis of the Association of Proteins with Membranes PDF Unit 5.5 Determination of Molecular ) Unit 5.6 Identification of Proteins in Complex Mixtures Using Liquid Chromatography and Mass Spectrometry PDF

Chapter 6 Electrophoresis and Immunoblotting Introduction PDF Unit 6.1 One-Dimensional SDS Gel Electrophoresis of Proteins PDF Unit 6.2 Immunoblotting and Immunodetection PDF Unit 6.3 Detection and Quantitation of Radiolabeled Proteins in Gels and Blots PDF Unit 6.4 Two-Dimensional Gel Electrophoresis PDF Unit 6.5 One-Dimensional Electrophoresis Using Nondenaturing Conditions PDF Unit 6.6 Staining Proteins in Gels PDF Unit 6.7 Agarose Gel Electrophoresis of Proteins PDF Unit 6.8 Fluorescence Detection of Glycoproteins in Gels and on Electroblots PDF Unit 6.9 Digital Electrophoresis Analysis PDF

Chapter 7 Protein Labeling and Immunoprecipitation Introduction PDF Unit 7.1 Metabolic Labeling with Amino Acids PDF Unit 7.2 Immunoprecipitation PDF Unit 7.3 Metabolic Labeling with Sulfate PDF Unit 7.4 Metabolic Labeling with Fatty Acids PDF Unit 7.5 Metabolic Labeling of Prenyl and Carboxyl-Methyl Groups PDF

Unit 7.6 Metabolic Labeling and Immunoprecipitation of Yeast Proteins PDF Unit 7.7 Metabolic Labeling and Immunoprecipitation of Drosophila Proteins PDF Unit 7.8 Metabolic Labeling of Glycoproteins with Radioactive Sugars PDF Unit 7.9 Analysis of Oxidative Modification of Proteins PDF Unit 7.10 Radioiodination of Cellular Proteins PDF

Chapter 8 Cell Cycle Analysis Introduction PDF Unit 8.1 Overview of the Cell Cycle PDF Unit 8.2 Assays for CDK Activity and DNA Replication in the Cell Cycle PDF Unit 8.3 Methods for Synchronizing Cells at Specific Stages of the Cell Cycle PDF Unit 8.4 Determining Cell Cycle Stages by Flow Cytometry PDF Unit 8.5 Centrifugal Elutriation to Obtain Synchronous Populations of Cells PDF

Chapter 9 Cell Adhesion Introduction PDF Unit 9.1 Cell-Substrate Adhesion Assays PDF Unit 9.2 Quantitative Measurement of Cell Adhesion Using Centrifugal Force PDF Unit 9.3 Cadherin-Dependent Cell-Cell Adhesion PDF Unit 9.4 Analyzing Integrin-Dependent Adhesion PDF Unit 9.5 Analysis of Cell-Cell Contact Mediated by Ig Superfamily Cell Adhesion Molecules PDF Unit 9.6 Measurement of Adhesion Under Flow Conditions PDF

Chapter 10 Extracellular Matrix Introduction PDF Unit 10.1 Overview of Extracellular Matrix PDF Unit 10.2 Preparation of Basement Membrane Components from EHS Tumors PDF Unit 10.3 Preparation of Gelled Substrates PDF Unit 10.4 Preparation of Extracellular Matrices Produced by Cultured Corneal Endothelial and PF-HR9 Endodermal Cells PDF Unit 10.5 Purification of Fibronectin PDF Unit 10.6 Purification of Vitronectin PDF Unit 10.7 Proteoglycan Isolation and Analysis PDF Unit 10.8 Matrix Metalloproteinases PDF Unit 10.9 Preparation of Extracellular Matrices Produced by Cultured Fibroblasts PDF Unit 10.10 Purification and Analysis of Thrombospondin-1 PDF Unit 10.11 Purification of SPARC/Osteonectin PDF

Chapter 11 In Vitro Reconstitution Introduction PDF Unit 11.1 Overview of Eukaryotic In Vitro Translation and Expression Systems PDF Unit 11.2 In Vitro Translation PDF Unit 11.3 In Vitro Analysis of Endoplasmic-Reticulum-to-Golgi Transport in Mammalian Cells PDF Unit 11.4 Cotranslational Translocation of Proteins into Canine Rough Microsomes PDF

Unit 11.5 In Vitro Analysis of SV40 DNA Replication PDF Unit 11.6 In Vitro Transcription PDF Unit 11.7 Nuclear Import in Digitonin-Permeabilized Cells PDF Unit 11.8 In Vitro Translation Using HeLa Extract PDF Unit 11.9 Analysis of Eukaryotic Translation in Purified and Semipurified Systems PDF Unit 11.10 Preparation and Use of Interphase Xenopus Egg Extracts PDF Unit 11.11 Analysis of the Cell Cycle Using Xenopus Egg Extracts PDF Unit 11.12 Analysis of Apoptosis Using Xenopus Egg Extracts PDF Unit 11.13 Mitotic Spindle Assembly In Vitro PDF Unit 11.14 Analysis of RNA Export Using Xenopus Oocytes PDF Unit 11.15 In Vitro Analysis of Peroxisomal Protein Import PDF Unit 11.16 In Vitro Analysis of Chloroplast Protein Import PDF Unit 11.17 In Vitro RNA Splicing in Mammalian Cell Extracts PDF

Chapter 12 Cell Motility Introduction PDF Unit 12.1 Chemotaxis Assays for Eukaryotic Cells PDF Unit 12.2 Invasion Assays PDF Unit 12.3 Cell Traction PDF Unit 12.4 Cell Wound Assays PDF Unit 12.5 Dictyostelium Cell Dynamics PDF Unit 12.6 Optical Microscopy—Based Migration Assay for Human Neutrophils PDF Unit 12.7 Actin-Based Motility Assay PDF

Chapter 13 Organelle Motility Introduction PDF Unit 13.1 Microtubule/Organelle Motility Assays PDF Unit 13.2 In Vitro Motility Assay to Study Translocation of Actin by Myosin PDF Unit 13.3 Organelle Motility in Plant Cells: Imaging Golgi and ER Dynamics with GFP PDF Unit 13.4 Movement of Nuclei PDF Unit 13.5 Measuring Dynamics of Nuclear Proteins by Photobleaching PDF

Chapter 14 Signal Transduction: Protein Phosphorylation Introduction PDF Unit 14.1 Overview of Protein Phosphorylation PDF Unit 14.2 Immunological Detection of Phosphorylation PDF Unit 14.3 Detection of MAP Kinase Signaling PDF Unit 14.4 Labeling Cultured Cells with 32Pi and Preparing Cell Lysates for Immunoprecipitation PDF Unit 14.5 Phosphoamino Acid Analysis PDF Unit 14.6 Determination of Akt/PKB Signaling PDF

Chapter 15 Protein Trafficking Introduction PDF Unit 15.1 Overview of Protein Trafficking in the Secretory and Endocytic Pathways PDF

Unit 15.2 Use of Glycosidases to Study Protein Trafficking PDF Unit 15.3 Endocytosis: Biochemical Analyses PDF Unit 15.4 Determining Protein Transport to the Plasma Membrane PDF Unit 15.5 Analysis of Membrane Traffic in Polarized Epithelial Cells PDF Unit 15.6 Analysis of Protein Folding and Oxidation in the Endoplasmic Reticulum PDF Unit 15.7 Measurements of Phagocytosis and Phagosomal Maturation PDF

Chapter 16 Antibodies as Cell Biological Tools Introduction PDF Unit 16.1 Production of Monoclonal Antibodies PDF Unit 16.2 Production of Polyclonal Antisera PDF Unit 16.3 Purification of Immunoglobulin G PDF Unit 16.4 Fragmentation of Immunoglobulin G PDF Unit 16.5 Antibody Conjugates for Cell Biology PDF Unit 16.6 Production of Antibodies That Recognize Specific Tyrosine-Phosphorylated Peptides PDF

Chapter 17 Macromolecular Interactions in Cells Introduction PDF Unit 17.1 Imaging Protein-Protein Interactions by Fluorescence Resonance Energy Transfer (FRET) Microscopy PDF Unit 17.2 Identification of Protein Interactions by Far Western Analysis PDF Unit 17.3 Interaction Trap/Two-Hybrid System to Identify Interacting Proteins PDF Unit 17.4 Mapping Protein-Protein Interactions with Phage-Displayed Combinatorial Peptide Libraries PDF Unit 17.5 Protein-Protein Interactions Identified by Pull-Down Experiments and Mass Spectrometry PDF Unit 17.6 Measuring Protein Interactions by Optical Biosensors PDF Unit 17.7 Chromatin Immunoprecipitation for Determining the Association of Proteins with Specific Genomic Sequences In Vivo PDF Unit 17.8 Isothermal Titration Calorimetry PDF

Chapter 18 Cellular Aging and Death Introduction PDF Unit 18.1 Current Concepts in Cell Death PDF Unit 18.2 Analysis of Caspase Activation During Apoptosis PDF Unit 18.3 Assessment of Apoptosis and Necrosis by DNA Fragmentation and Morphological Criteria PDF Unit 18.4 Quantitative Fluorescence In Situ Hybridization (Q-FISH) PDF Unit 18.5 Analysis of Mitochondrial Dysfunction During Cell Death PDF Unit 18.6 Analysis of Telomeres and Telomerase PDF Unit 18.7 Nonisotopic Methods for Determination of Poly(ADP-Ribose) Levels and Detection of Poly(ADPRibose) Polymerase PDF Unit 18.8 Flow Cytometry of Apoptosis PDF

Chapter 19 Whole Organism and Tissue Analysis Introduction PDF Unit 19.1 Overview of Metastasis Assays PDF

Unit 19.2 Tail Vein Assay of Cancer Metastasis PDF Unit 19.3 Microanalysis of Gene Expression in Tissues Using T7-SAGE: Serial Analysis of Gene Expression After High-Fidelity T7-Based RNA Amplification PDF Unit 19.4 SAGE Analysis from 1 µg of Total RNA PDF Unit 19.5 The Chick Chorioallantoic Membrane as an In Vivo Angiogenesis Model PDF Unit 19.6 Experimental Metastasis Assays in the Chick Embryo PDF

Chapter 20 Expression and Introduction of Macromolecules into Cells Introduction PDF Unit 20.1 Direct Introduction of Molecules into Cells PDF Unit 20.2 Protein Transduction: Generation of Full-Length Transducible Proteins Using the TAT System PDF Unit 20.3 Calcium Phosphate Transfection PDF Unit 20.4 Transfection Using DEAE-Dextran PDF Unit 20.5 Transfection by Electroporation PDF Unit 20.6 Transfection of Cultured Eukaryotic Cells Using Cationic Lipid Reagents PDF Unit 20.7 Optimization of Transfection PDF

Chapter 21 Fluorescent Protein Technology Introduction PDF Unit 21.1 Measuring Protein Mobility by Photobleaching GFP Chimeras in Living Cells PDF Unit 21.2 Fluorescence Localization After Photobleaching (FLAP) PDF

Chapter 22 Cell Biology of Chromosomes and Nuclei Introduction PDF Unit 22.1 Overview of Cytogenetic Chromosome Analysis PDF Unit 22.2 Preparation of Cytogenetic Specimens from Tissue Samples PDF Unit 22.3 Traditional Banding of Chromosomes for Cytogenetic Analysis PDF Unit 22.4 Fluorescence In Situ Hybridization (FISH) PDF Unit 22.5 Multi-Color FISH Techniques PDF

Appendix 1 Useful Information and Data 1A Useful Measurements and Data PDF 1B Compendium of Drugs Commonly Used in Cell Biology Research PDF 1C Identification of Motifs in Protein Sequences PDF 1D Safe Use of Radioisotopes PDF 1E Absorption and Emission Maxima for Common Fluorophores PDF 1F Importing Biological Materials PDF 1G Centrifuges and Rotors PDF 1H Internet Basics for Biologists PDF

Appendix 2 Laboratory Stock Solutions and Equipment 2A Common Stock Solutions, Buffers, and Media PDF 2B Medium Formulations PDF 2C Standard Laboratory Equipment PDF

Appendix 3 Commonly Used Techniques 3A Molecular Biology Techniques PDF 3B Spectrophotometric Determination of Protein Concentration PDF 3C Dialysis and Concentration of Protein Solutions PDF 3D Quantification of DNA and RNA with Absorption and Fluorescence Spectroscopy PDF 3E Silanizing Glassware PDF 3F Enzymatic Amplification of DNA by PCR: Standard Procedures and Optimization PDF 3G Micro RT-PCR PDF 3H The Colorimetric Detection and Quantitation of Total Protein PDF

SUPPLIERS APPENDIX Selected Suppliers of Reagents and Equipment PDF

DNAthink于2004-10-18

CHAPTER 1 Cell Culture INTRODUCTION

C

ell biology traces its roots to the introduction of the concept of “cells” by Robert Hooke in the second half of the 17th century. However, not until nearly halfway through the 20th century were techniques for the culture of cells developed. In fact, 1998 marked the golden anniversary of the first continuous mammalian cell line. Cell culture has become such an integral part of cell biology that it is somewhat difficult to imagine the field in the B.C. (“Before Culture”) era. Cell culture also represents the primary way in which cell biology reaches into related disciplines, since the maintenance and propagation of cells has become an important component of biochemistry, biophysics, genetics, immunology, physiology, molecular biology, and neuroscience. Accordingly, it is altogether fitting that the first chapter of Current Protocols in Cell Biology should present methods related to the culture of cells. The immediate aim of cell culture is to maintain or expand a population of cells, and the single most important consideration is cell viability. Determining the number of cells and their viability is important in standardizing culture and experimental conditions. As viable cells replicate in culture, passaging of the cells allows their number to be expanded to meet experimental needs. The ability to freeze, store, and recover cells provides an essential safeguard against losing a cell line to contamination, incubator malfunction, or an error on the part of the investigator. In addition to preserving the cells, maintenance of a frozen stock is desirable to avoid cellular senescence and genetic drift. Chapter 1 therefore begins with protocols for passaging cells, freezing and thawing cells, and determining cell number and viability (UNIT 1.1). Success in cell culture is highly dependent on the choice of a medium. At minimum, a medium must provide the nutritional requirements of the cells as well as any required growth factors, and maintain pH and osmolarity compatible with survival. The historical development of a wide variety of culture media has influenced significantly the types of cells that can be studied experimentally, since cell lines that proliferate in a particular environment are always selected at the expense of those that do not. The second unit of Chapter 1 therefore focuses on media used in culturing cells and provides descriptions of standard, serum-free, and selective media, as well as the use of soft agar for anchorageindependent growth (UNIT 1.2).

The next three units of this chapter deal with microbial contamination of cell cultures. describes basic aseptic techniques and the laminar flow hoods that are the main weapons in the constant battle against contamination. UNIT 1.4 provides protocols related to sterilization, namely filtration and heat sterilization (e.g., autoclaving), as well as the use of disinfectants. UNIT 1.5 describes methods for detecting microbial contaminants (bacteria, fungi, and mycoplasmas). While the best way to deal with such contamination may well be to review the previous unit on autoclaving and faithfully apply its precepts, situations do arise where an attempt to salvage a contaminated culture is warranted. UNIT 1.5 details the use of antibiotics for this purpose. UNIT 1.3

Of course there are cell biologists who do not see the growth of fungi as an annoying contamination of their mammalian cell cultures but as a desirable goal. For scientists who Cell Culture Contributed by Joe B. Harford Current Protocols in Cell Biology (2003) 1.0.1-1.0.2 Copyright © 2003 by John Wiley & Sons, Inc.

1.0.1 Supplement 19

wish to propagate yeast, UNIT 1.6 provides recipes for media and descriptions of some basic culture methodologies. UNIT 1.7 represents the first unit of Chapter 1 dealing with culture of plants cells, specifically the culture and transformation of BY-2 cells derived from tobacco. BY-2 cells have been described as the HeLa cell of higher plants.

Future units in Chapter 1 will cover specialized systems for cell culture (e.g., cell cloning, polarized cells, and three-dimensional cultures), as well as additional units on the propagation of plant cells, cells from other so-called simpler organisms, and viruses. For additional information on mammalian cell culture, readers are directed to Freshney (1993). LITERATURE CITED Freshney, R.I. 1993. Culture of Animal Cells. A Manual of Basic Techniques, 3rd ed. Wiley-Liss, New York.

Joe B. Harford

Introduction

1.0.2 Supplement 19

Current Protocols in Cell Biology

Basic Techniques for Mammalian Cell Tissue Culture

UNIT 1.1

Tissue culture technology has found wide application in the field of cell biology. Cell cultures are utilized in cytogenetic, biochemical, and molecular laboratories for diagnostic as well as research studies. In most cases, cells or tissues must be grown in culture for days or weeks to obtain sufficient numbers of cells for analysis. Maintenance of cells in long-term culture requires strict adherence to aseptic technique to avoid contamination and potential loss of valuable cell lines (see UNIT 1.3). An important factor influencing the growth of cells in culture is the choice of tissue culture medium. Many different recipes for tissue culture media are available and each laboratory must determine which medium best suits their needs. Individual laboratories may elect to use commercially prepared medium or prepare their own. Commercially available medium can be obtained as a sterile and ready-to-use liquid, in a concentrated liquid form, or in a powdered form. Besides providing nutrients for growing cells, medium is generally supplemented with antibiotics, fungicides, or both to inhibit contamination. Medium preparation is discussed in UNIT 1.2. As cells reach confluency, they must be subcultured or passaged. Failure to subculture confluent cells results in reduced mitotic index and eventually cell death. The first step in subculturing monolayers is to detach cells from the surface of the primary culture vessel by trypsinization or mechanical means. The resultant cell suspension is then subdivided, or reseeded, into fresh cultures. Secondary cultures are checked for growth, fed periodically, and may be subsequently subcultured to produce tertiary cultures, etc. The time between passaging cells depends on the growth rate and varies with the cell line. The Basic Protocol describes subculturing of a monolayer culture grown in petri plates or flasks; the Alternate Protocol 1 describes passaging of suspension cultures. Support Protocols describe freezing of monolayer cells, thawing and recovery of cells, counting cells using a hemacytometer, and preparing cells for transport. Alternate Protocol 2 describes freezing of suspension cells. CAUTION: When working with human blood, cells, or infectious agents, appropriate biosafety practices must be followed. NOTE: All solutions and equipment coming into contact with living cells must be sterile, and aseptic technique should be used accordingly. TRYPSINIZING AND SUBCULTURING CELLS FROM A MONOLAYER 2

A primary culture is grown to confluency in a 60-mm petri plate or 25-cm tissue culture flask containing 5 ml tissue culture medium. Cells are dispersed by trypsin treatment and then reseeded into secondary cultures. The process of removing cells from the primary culture and transferring them to secondary cultures constitutes a passage, or subculture.

BASIC PROTOCOL

Materials Primary cultures of cells HBSS (APPENDIX 2A) without Ca2+ and Mg2+, 37°C Trypsin/EDTA solution (see recipe), 37°C Complete medium with serum: e.g., supplemented DMEM (APPENDIX 2A) with 10% to 15% (v/v) FBS (complete DMEM-10 or -15), 37°C Cell Culture Contributed by Mary C. Phelan Current Protocols in Cell Biology (1998) 1.1.1-1.1.10 Copyright © 1998 by John Wiley & Sons, Inc.

1.1.1

Sterile Pasteur pipets 37°C warming tray or incubator Tissue culture plasticware or glassware including pipets and 25-cm2 flasks or 60-mm petri plates, sterile NOTE: All culture incubations should be performed in a humidified 37°C, 5% CO2 incubator unless otherwise specified. Some media (e.g., DMEM) may require altered levels of CO2 to maintain pH 7.4. 1. Remove all medium from primary culture with a sterile Pasteur pipet. Wash adhering cell monolayer once or twice with a small volume of 37°C HBSS without Ca2+ and Mg2+ to remove any residual FBS that may inhibit the action of trypsin. Use a buffered salt solution that is Ca2+ and Mg2+ free to wash cells. Ca2+ and Mg2+ in the salt solution can cause cells to stick together. If this is the first medium change, rather than discarding medium that is removed from primary culture, put it into a fresh dish or flask. The medium contains unattached cells that may attach and grow, thereby providing a backup culture.

2. Add enough 37°C trypsin/EDTA solution to culture to cover adhering cell layer. 3. Place plate on a 37°C warming tray 1 to 2 min. Tap bottom of plate on the countertop to dislodge cells. Check culture with an inverted microscope to be sure that cells are rounded up and detached from the surface. If cells are not sufficiently detached, return plate to warming tray for an additional minute or two.

4. Add 2 ml 37°C complete medium. Draw cell suspension into a Pasteur pipet and rinse cell layer two or three times to dissociate cells and to dislodge any remaining adherent cells. As soon as cells are detached, add serum or medium containing serum to inhibit further trypsin activity that might damage cells. If cultures are to be split 1/3 or 1/4 rather than 1/2, add sufficient medium such that 1 ml of cell suspension can be transferred into each fresh culture vessel.

5. Add an equal volume of cell suspension to fresh plates or flasks that have been appropriately labeled. Alternatively, cells can be counted using a hemacytometer or Coulter counter and diluted to the desired density so a specific number of cells can be added to each culture vessel. A final concentration of ∼5 × 104 cells/ml is appropriate for most subcultures. For primary cultures and early subcultures, 60-mm petri plates or 25-cm2 flasks are generally used; larger vessels (e.g., 150-mm plates or 75-cm2 flasks) may be used for later subcultures. Cultures should be labeled with date of subculture and passage number.

6. Add 4 ml fresh medium to each new culture. Incubate in a humidified 37°C, 5% CO2 incubator. If using 75-cm2 culture flasks, add 9 ml medium per flask. Some labs now use incubators with 5% CO2 and 4% O2. The low oxygen concentration is thought to simulate the in vivo environment of cells and to enhance cell growth. For some media it is necessary to adjust the CO2 to a higher or lower level to maintain the pH at 7.4.

7. If necessary, feed subconfluent cultures after 3 or 4 days by removing old medium and adding fresh 37°C medium. Basic Techniques for Mammalian Cell Tissue Culture

8. Passage secondary culture when it becomes confluent by repeating steps 1 to 7, and continue to passage as necessary.

1.1.2 Current Protocols in Cell Biology

PASSAGING CELLS IN SUSPENSION CULTURE A suspension culture is grown in culture flasks in a humidified 37°C, 5% CO2 incubator. Passaging of suspension cultures is somewhat less complicated than passaging of monolayer cultures. Because the cells are suspended in medium rather than attached to a surface, it is not necessary to disperse them enzymatically before passaging. However, before passaging, cells must be maintained in culture by feeding every 2 to 3 days until they reach confluency (i.e., until the cells clump together in the suspension and the medium appears turbid when the flask is swirled).

ALTERNATE PROTOCOL 1

NOTE: All culture incubations should be performed in a humidified 37°C, 5% CO2 incubator unless otherwise specified. Some media (e.g., DMEM) may require altered levels of CO2 to maintain pH 7.4. 1. Feed cells as follows every 2 to 3 days until the cultures are confluent: a. Remove flask of suspension cells from incubator, taking care not to disturb those that have settled to the flask bottom. b. Aseptically remove and discard about one-third of the medium from flask and replace with an equal volume of prewarmed (37°C) medium. If the cells are growing rapidly, add an additional 10% medium by volume in order to maintain optimum concentration of 1 × 106 cells/ml. Gently swirl flask to resuspend cells. c. Return flask to incubator. If there is 10 liters) that are used under positive pressure are more suitable for larger volumes.

8. Add serum to the desired final concentration at the time of use. Basal nutrient medium and the serum supplement should be stored individually at 4°C, and the complete medium should be made up at the time of use and only in the volume necessary. Working volumes of serum should be stored at 4°C and used within several weeks. Serum should not be subjected to repeated freezing and thawing, but it can be stored for at least 2 years at −20°C with little deterioration in growth-promoting activity. In this way, medium components are not wasted, and the chances of detecting, isolating, and eliminating contamination with minimal losses are increased.

PREPARING MEDIA FOR REDUCED-SERUM OR SERUM-FREE GROWTH The most obvious advantages of serum-free cell culture are that it costs less and it simplifies the purification of cell products. However, it is in the increased knowledge of cell physiology that the real value of serum-free medium lies. Both reduced-serum and serum-free media are intermediates in a continuum between completely undefined mixtures of biological fluids and chemically defined, protein-free medium. Although mixtures of biological fluids and protein-free medium have useful applications, neither extreme provides a suitable environment for studies of cellular physiology. At the current time, the most physiologically relevant culture medium for an isolated cell type is a defined, protein-supplemented medium consisting of required components at optimal concentrations and extracellular matrix constituents. This set of conditions can be approached for an increasing number of cell types, but for some cell types, a reduced-serum medium is the best that can be achieved. This state of affairs reflects incomplete knowledge of cell growth requirements, and it suggests that there are novel mediators of cell proliferation and differentiation that remain to be discovered.

BASIC PROTOCOL 2

Cell Culture


Components of Reduced-Serum and Serum-Free Mediuma

Table 1.2.2

Component

Final concentration

Stock concentration

Suggested supplier(s)

Undefined supplements Serab Pituitary extract Conditioned medium

5% to 20% (v/v) 5 to 100 µg/ml 5% to 50% (v/v)

100% (v/v) 1 to 2 mg/ml 100% (v/v)

Hyclone, Sigma, or UBI UBI, Clonetics Not commercially availablec

Energy sources Glucose Glutamine

1 to 4.5 g/liter 1 to 2 mM

None None

Sigma Sigma

Attachment factors Collagen type I Fibronectin Vitronectin

10 to 50 µg/ml 1 to 10 µg/ml 1 to 10 µg/ml

3 to 4 mg/ml 0.5 to 1 mg/ml 0.5 to 1 mg/ml

UBI, Sigma Sigma Sigma

Hormone Insulin

1 to 10 µg/ml

1 mg/ml

Sigma, UBI

Carrier proteins Transferrin BSA, fatty acid–free

5 to 30 µg/ml 0.5 to 4 mg/ml

1 mg/ml 50 mg/ml

Sigma, UBI Bayer

Trace element Selenium, sodium salt

1 to 20 nM

2 µM

Sigma

2 mM 50 mg/ml 20 to 50 mg/ml 2 to 4 mg/ml 1 to 2 mg/ml

Sigma Sigma Sigma Steraloids, Sigma Chemicon International

Lipids and lipid precursors Ethanolamine 1 to 20 µM Fraction V BSA 0.05 to 5 mg/ml Unsaturated fatty acids 1 to 10 µg/ml Sterols 1 to 20 µg/ml Low-density lipoprotein 1 to 20 µg/ml

aNonsterile stock solutions should be sterilized by filtration. Add glucose and glutamine as dry powder (or frozen aliquots that have

been thawed) to reconstituted powdered medium. bSee Critical Parameters for discussion of FBS. cConditioned medium must be made in the investigator’s laboratory, and the choice of cells used depends on the investigator’s purpose.

If all the growth requirements of the cells of interest can be satisfied, the undefined medium supplement can be completely eliminated. The limitation of this approach is that not all of the cell growth regulators operating within tissues have been discovered. As the variety of cell types that can be cultured in vito for extended periods expands, new growth activities may be identified and novel growth factors purified. Materials Basal nutrient medium, such as DMEM, Ham’s F-12, or RPMI 1640 (APPENDIX 2B; see Critical Parameters for discussion of medium selection) Nutrients: inorganic salts, amino acids, and vitamins (e.g., Sigma) Trace elements (e.g., Sigma) Supplements: growth factors and hormones (e.g., Sigma, UBI, Becton Dickinson Labware) and other assorted medium components (Table 1.2.2) Additional reagents and equipment for culture of mammalian cells (UNIT 1.1)

Media for Culture of Mammalian Cells

Optimize nutrient medium 1. Starting with a complete medium empirically determined to best support the growth of the cells of interest, reduce the concentration of the undefined medium supplement until cell proliferation is suboptimal but cell viability remains high.


2. Vary the concentrations of individual components of the nutrient medium in a cell proliferation assay. Vary the concentrations of components in the following groupings, which have been found to be limiting for proliferation of at least one cell type: a. energy sources (glucose and glutamine); b. trace elements and electrolytes (Na+, Cl−, K+, Ca2+, Mg2+, Zn2+, Cu2+, Fe2+, selenium, H2PO4−, and HCO3−); c. amino acids (glutamine, cystine, cysteine, histidine); d. vitamins (biotin, vitamin B12); e. lipids and lipid precursors (oleic acid, linoleic acid, or cholesterol conjugated to fatty acid–free BSA; undefined lipids bound to fraction V BSA; low-density lipoprotein; ethanolamine or phosphoethanolamine). Determine the optimal concentration range for the most limiting factor, and fix its concentration in the center of that range. Repeat the growth assays to sequentially identify and optimize each limiting factor. Concentrations of 0.1×, 1×, and 10× those listed in the chosen medium formulation are initially useful to determine which components are limiting factors for proliferation.

3. Lower the concentration of the undefined supplement and repeat the optimization procedure. Continue optimizing the nutrient medium until reductions in the concentration of the undefined medium supplement can no longer be compensated for. Each new supplement concentration is tested on an individual culture for a finite period (3 to 10 days). The best concentration, supporting the most cell growth, is incorporated into the next round of testing on new cultures.

Optimize medium supplements 4. Optimize medium supplements such as growth factors, hormones, transport proteins, and attachment factors either independently of or in conjunction with the nutrient medium components. The set of defined supplements that are required for an individual cell type may be less complex in the presence of an optimized nutrient medium. If an optimized nutrient medium is not being used, then a nutrient-rich basal medium such as DMEM/F-12 (see recipe) is recommended for supplement optimization (see Critical Parameters). As with nutrient optimization, the strategy is to decrease the concentration of an undefined supplement to give suboptimal growth with high cell viability and then to restore proliferation with optimized concentrations of defined supplements.

PREPARATION OF SELECTIVE MEDIA: HAT MEDIUM Culture medium can be used to intentionally select mammalian cells with desired properties from a mixed population of cells. This strategy is founded on the proven effectiveness of selective media in selecting strains of mutant microorganisms that, for example, cannot grow in minimal media but thrive in a medium supplemented with one particular nutrient. Although mammalian cells with desired characteristics proliferate in selective media, in this case selective pressure is applied to inhibit or, preferably, to kill cells that do not possess those characteristics. Thus, the efficacy of a selective medium depends on the efficiency with which it eliminates unwanted cells and on the degree to which the selected phenotypic trait is expressed. Because monoclonal antibodies are commonly used reagents in cell and molecular biology, the use of selective media that target differences in metabolic pathways is illustrated with two protocols for selecting hybridomas (see Background Information for further discussion). In both protocols, unfused spleen cells do not survive 7 to 10 days.

BASIC PROTOCOL 3

Cell Culture


Materials Spleen cell × myeloma fusion products (10:1) RD medium (Life Technologies) with 10% FBS (Hyclone; see recipe) 4 × 10−5 M aminopterin (A solution; 100× stock in 0.1 N NaOH) 1 × 10−5 M hypoxanthine/1.6 × 10−3 M thymidine in water (HT solution; 100× stock) HAT medium: RD/10% FBS/1× A solution/1× HT solution 96-well tissue culture plates Additional reagents and equipment for culture of mammalian cells (UNIT 1.1) 1. Resuspend the fusion products in RD/10% FBS at a concentration of 1 × 106 myeloma cells per ml. Add 0.01× volumes of A and HT solutions to the cell suspension to make HAT selection medium. 2. Plate 0.1 ml of cell suspension per well into 96-well plates. Incubate the plates in a humidified 37°C, 5% CO2 incubator. 3. Every 2 or 3 days, remove half the existing medium from the wells by aspiration and replace with fresh HAT medium. 4. After 21 days, screen hybridoma supernatants for the presence of the antibodies of interest. 5. Wean hybridomas stepwise from HAT medium by transferring them to HT-supplemented medium and then to RD/10% FBS over a 2-week period. Replace 50% of the medium with HT medium four times at 3-day intervals. Replace 50% of the medium with RD/10% FBS at 3-day intervals. Aminopterin is toxic, so it is advisable to wean hybridomas from HAT medium as soon as possible. ALTERNATE PROTOCOL

PREPARATION OF SELECTIVE MEDIA: CHOLESTEROL-FREE, SERUM-FREE MEDIUM Most of the mouse myeloma cell lines survive in serum-supplemented medium, but they die in the absence of cholesterol (see Background Information for explanation). This conditional lethal defect has been exploited to create an alternative selection process for hybridomas using cholesterol-free medium. Because they are produced by fusion with spleen cells capable of producing cholesterol, NS-1 hybridomas survive in cholesterol-free medium, but the parent NS-1 myeloma cells are selected against. This selective medium allows for the outgrowth of up to 10 times as many hybridomas as HAT medium. This procedure can be used with any myeloma cell line that is unable to synthesize cholestrol. Materials Spleen cell × NS-1 myeloma fusion products (10:1) RD medium with 5F supplement (see recipes) 100× BSA–oleic acid conjugate solution: fatty acid−free BSA (e.g., Bayer; 50 mg/ml) conjugated with oleic acid (e.g., Sigma; 500 µg/ml) in PBS 96-well tissue culture plates Additional reagents and equipment for culture of mammalian cells (UNIT 1.1)


1. Make complete RD/5F medium by adding appropriate volumes of the stock solutions of insulin, transferrin, ethanolamine, 2-mercaptoethanol, and sodium selenite (e.g., Sigma) to RD medium. Supplement the medium with 1/100 vol of 100× BSA−oleic acid conjugate. BSA–oleic acid acid conjugate is available from Sigma. For a protocol, see Kawamoto et al. (1983).


2. Resuspend fusion products at 2 or 3 × 105 NS-1 cells per milliliter of medium. 3. Plate 0.1 ml of cell suspension per well in 96-well plates. Incubate the plates in a humidified 37°C, 5% CO2 incubator. 4. Add 0.5× volume of fresh medium to the wells every 2 or 3 days after removing half the existing medium by aspiration. 5. Screen hybridoma supernatants for antibodies of interest after 10 to 14 days. Maintain hybridomes in RD/SF medium supplemented with BSA–oleic acid. GROWTH OF TRANSFORMED CELLS IN SOFT AGAR Malignantly transformed cells can differ from their normal counterparts in a number of respects. Chief among these differences are a loss of contact-inhibited growth, the acquisition of an infinite life span, and the ability to form tumors in animal hosts. Freedman and Shin (1974) found that there was a general correlation between the tumorigenic potential of transformed cells in vivo and their ability to grow in an anchorage-independent manner in vitro. Although this generalization does not hold for every transformed cell, growth in soft agar can be used as a surrogate in vitro assay for transformation and tumorigenicity. The advantages of this method are that it is relatively easy, it may take much less time than an in vivo tumorigenesis assay, and it does not require the maintenance and care of experimental animals. However, if growth in soft agar is used as a measure of transformation of cells that have been manipulated in vitro, an in vivo tumorigenesis assay is required to determine whether the cells are malignantly transformed.

BASIC PROTOCOL 4

Materials 2% (w/v) agar (e.g., Difco; see recipe) Basal nutrient medium, such as DMEM, Ham’s F-12, or RPMI 1640 (APPENDIX 2B; see Critical Parameters for discussion of medium selection), with 24.6% and 20% FBS Single-cell suspension 12-well culture plates (e.g., Corning Costar) 15-ml polycarbonate conical centrifuge tubes (e.g., Sarstadt), sterile 1. For each set of replicate wells, add 2.25 ml of 2% agar solution to 9.75 ml of medium containing 24.6% FBS to give a solution of 0.375% agar in 20% FBS. Dispense 2-ml aliquots into five sterile 15-ml polypropylene conical tubes and incubate at 45°C. 2. To separate tubes, add 50, 100, 200, 500, or 1000 cells to a final volume of 0.5 ml medium with 20% FBS. Prepare an additional set of tubes for each set of replicate cultures. 3. Add each cell suspension to an aliquot of agar solution, mix, and quickly pour into a well of a 12-well plate. 4. Incubate the plate at 37°C in a humid atmosphere of 5% CO2 until cell colonies appear. 5. Count colonies >32 cells (five doublings) under phase contrast with an inverted microscope, and calculate colony formation efficiency (percentage of plated cells that formed colonies). To prevent any cell attachment to the plastic substratum, cells in 0.3% agar can be overlaid on a preformed layer of 0.5 ml of 0.5% agar in medium supplemented with 20% FBS.

Cell Culture


SUPPORT PROTOCOL 1

pH CONTROL IN MEDIA Most cell lines proliferate in medium with a pH of 7.4, and they exhibit decreased viability and rates of proliferation as the medium becomes progressively more acidic or more basic (Eagle, 1973). Culture media must buffer the CO2 and lactic acid produced as cells metabolize glucose and glutamine. Historically, bicarbonate, HCO3−, in conjunction with atmospheric CO2, has been used as a buffering system. Each basal medium formulation has a recommended concentration for sodium bicarbonate, usually 20 to 40 mM, to maintain pH and osmolarity. Media that are to be incubated in an elevated CO2 atmosphere contain higher concentrations of bicarbonate than those designed to be used at ambient CO2 levels. Most cell culture media require an atmosphere of 5% CO2 to maintain pH 7.4. However, certain media contain levels of bicarbonate that require different amounts of CO2. For example, DMEM containing 3700 mg/liter of sodium bicarbonate equilibrates to ∼pH 7.6 in a 5% CO2 environment and requires 10% CO2 to maintain pH 7.4. The low pKa of bicarbonate (pKa = 6.1) makes it a poor buffer around pH 7.4, and, in the absence of atmospheric CO2, the breakdown of H2CO3 formed from bicarbonate releases CO2 that comes out of solution, causing a rise in pH. With the development of Good buffers (Good et al., 1966), nontoxic buffering agents effective in the pH range of 6 to 8, such as PIPES (pKa = 6.8), MOPS (pKa = 7.2), TES (N-tris[hydroxymethyl]methyl-2-aminoethanesulfonic acid; pKa = 7.5), and HEPES (pKa = 7.55), became available to the research community. HEPES in a concentration range of 10 mM to 25 mM has become a standard buffer in serum-free medium, but it is used in addition to and not in place of the bicarbonate and CO2 system. Phenol red is an indicator dye that is commonly added to medium to provide a visual assessment of pH. Red at pH 7.4, it becomes orange (pH 7.0) and then yellow (pH 6.5) as the pH decreases; it turns violet (pH 7.6) and purple (pH 7.8) as the pH rises. Culture medium should generally be replaced as the phenol red changes from orange to yellow, which reflects the accumulation of lactic acid. Materials Powdered medium without NaHCO3 or HEPES HEPES (e.g., Research Organics) NaHCO3 (e.g., J.T. Baker) 1. Dissolve powdered medium in water with gentle stirring. 2. Add HEPES (mol. wt. 238.3) to give a final concentration of 15 mM, and stir until dissolved. 3. Add NaHCO3 to the recommended concentration for the basal nutrient medium being used and stir. 4. Add other medium components, and adjust the pH to 7.4 (see Basic Protocol 1). 5. Filter sterilize the medium (see Basic Protocol 1, step 7). Filtering medium under vacuum may cause the pH to increase slightly. A small change in pH need not be compensated because other factors such as medium supplements, temperature, atmospheric pressure, and atmospheric CO2 levels can also affect pH.

6. Check the pH of the complete medium after it has equilibrated with incubator temperature and CO2 atmosphere.



USE OF ANTIBIOTICS IN MEDIA Antibiotics can be added to culture media to eliminate microbial contaminants (Perlman, 1979). The most common contaminants encountered are bacteria, yeast, other fungi, and mycoplasma, and the most common routes of contamination are operator error and nonsterile medium components. Of the common microbial contaminants, yeast and other fungi are very difficult to eradicate, and it is recommended that all heavily contaminated cultures be discarded unless the cells cannot be replaced. Thus, it is wise to maintain duplicate cultures of important cells and to cryopreserve cell lines as soon as possible. Penicillin and streptomycin are broad-spectrum antibacterial agents that are often added to culture media (see Basic Protocol 1 and Table 1.2.3). However, the routine use of antibiotics in media is not recommended, because when used to compensate for poor aseptic technique, they may select for antibiotic-resistant strains of microorganisms. Antibiotics may be used to best effect in primary cultures of cells for which the sterility of the tissue samples is in doubt. The antifungal agent amphotericin B should be used sparingly, as it is toxic to mammalian cells (Perlman, 1979), and it may select for the property of cholesterol auxotrophy (Sato et al., 1987). Mycoplasma and viruses are too small to be retained by 0.2-µm sterilization filters; mycoplasma can be treated with gentamicin or kanamycin, but viruses cannot be treated with antibiotics. There is no reliable method for eliminating viral contaminants from cell cultures.

SUPPORT PROTOCOL 2

To be effective in culture, antibiotics must have the following characteristics: they must completely eliminate the microbial contaminant; they must not affect the viability or metabolism of mammalian cells; and they must be compatible with medium components in an aqueous environment. In addition, because the identity of a contaminating species is usually unknown, antibiotics should act on a broad spectrum of mircoorganisms. A list of antibiotics commonly used in culture media is provided in Table 1.2.3. Materials Antibiotic Sterile solvent 0.2-µm-pore-size sterilizing filter 1. Dissolve the antibiotic at a 100× or greater concentration in an appropriate sterile solvent. If it is highly soluble in an aqueous solvent, use water or PBS. 2. If the antibiotic was not obtained in a sterile form, filter the solution through a 0.2-µm filter. 3. Store the antibiotic solution at 4°C prior to use or at −20°C for long-term storage. 4. Add the antibiotic to medium immediately prior to use.

Table 1.2.3

Some Antibiotics Used in Culture Media and Their Microbial Targets

Antibiotic

Concentration

Microbial targets

Amphotericin B Ampicillin Chloramphenicol Gentamicin Kanamycin Penicillin G Streptomycin Tetracycline

2.5 µg/ml 100 µg/ml 5 µg/ml 50 µg/ml 100 µg/ml 100 IU/ml 100 µg/ml 10 µg/ml

Yeast and other fungi Gram-positive and -negative bacteria Gram-negative bacteria Gram-positive and -negative bacteria, mycoplasma Gram-positive and -negative bacteria, mycoplasma Gram-positive bacteria Gram-positive and -negative bacteria Gram-positive and -negative bacteria, mycoplasma

Cell Culture


REAGENTS AND SOLUTIONS Use Milli-Q-purified water or equivalent in all recipes and protocol steps. For common stock solutions, see APPENDIX 2A; for suppliers, see SUPPLIERS APPENDIX.

Agar Dissolve 2% (w/v) agar (e.g., Difco) in water by boiling it or heating it in a microwave. Maintain soft agar at 45°C. DMEM/F12 medium, supplemented Make 1:1 (v/v) mixture of DMEM and F-12. Filter sterilize the medium and store at 4°C. Add the appropriate volume of serum at the time of use. Five factors (5F) supplement Make the following 200× stock solutions: 2 mg/ml insulin in 10 mM HCl 2 mg/ml transferrin in PBS 2 mM ethanolamine in water 2 mM 2-mercaptoethanol in water 2 µM sodium selenite in water Filter sterilize the stock solution and store at 4°C prior to use Add 0.5 ml of each stock solution per 100 ml of medium RD medium, supplemented Make 1:1 (v/v) mixture of RPMI 1640 and DMEM. Add glutamine to 2 mM, penicillin G to 100 IU/ml, and streptomycin to 50 µg/ml. Filter sterilize the medium and store at 4°C. Add the appropriate volume of serum at the time of use. COMMENTARY Background Information


It is beyond the scope of this unit to describe all of the medium formulations that have been developed for mammalian cell culture. The following references, which provide surveys of media and conditions that have been used successfully to culture numerous cell lines and a variety of cell types, are recommended as sources of this information: Bottenstein et al. (1979); Jacoby and Pastan (1979); Barnes et al. (1984); Freshney (1987, 1992); Baserga (1989); Davis (1994); and Sato et al. (1994). Because the immediate aim of the cell culturist is to maintain or expand a population of cells, culture media have been developed with an emphasis on maintaining cell viability and stimulating cell proliferation. These goals have necessarily led to culture media and culture conditions that select for cells that can proliferate at the expense of those that cannot. Thus, the choice of culture medium and the culture conditions used affect the properties and types of cells that can be studied in vitro. As an example of selection, normal rodent cells cultured on a plastic substratum in the presence of fetal bovine serum often go through a period of “crisis” in which a minority population of ane-

uploid cells survive and proliferate as immortalized but not completely transformed cells. Another example of unintentional selection through in vitro cell culture is the outgrowth of basal keratinocytes from cultured epidermal cells; this occurs because keratinocytes in the suprabasal layers of the epidermis differentiate and lose the ability to proliferate. Conversely, for some applications, culture media have been developed to deliberately select for the survival of specific cell types of interest. Low-calcium, serum-free media developed for many epithelial cell types (Sato et al., 1994) are selective in that they promote the proliferation of epithelial cells but inhibit the outgrowth of fibroblasts in primary cultures; surviving fibroblasts are eliminated by dilution as the cultures are repeatedly passaged (UNIT 1.1). Selective media are generally designed to exploit differences in susceptibility to metabolic inhibitors or differences in metabolic pathways. An example of a metabolic inhibitor as a selection agent is the use of the antibiotic G418 (geneticin) to kill nontransfected mammalian cells while allowing the growth of transfectants containing the bacterial gene for aminoglycoside phosphotransferase (neor; Southern and Berg, 1982).


Endogenous synthesis of purines and pyrimidines, which are essential components of nucleosides and nucleotides, can be blocked in mammalian cells by the folic acid analogue aminopterin (A). Such a block is circumvented in normal cells by salvage pathways that use hypoxanthine (H) and thymidine (T). However, cells deficient in the salvage pathway enzymes hypoxanthine-guanine phosphoribosyltransferase (HGPRT−) and thymidine kinase (TK−) cannot produce purines or pyrimidines by means of the salvage pathways in the presence of aminopterin, and die. Thus, HAT medium (Littlefield, 1964) selects for cells that contain functional HGPRT and TK enzymes, and it selects against cells that are HGPRT− or TK−. This selection system has been made applicable to the selection of hybridomas by the generation of HGPRT– mouse myeloma cell lines. When these cells are fused with HGPRT+ spleen cells from an immunized mouse, the resulting HGPRT+ hybridomas survive in HAT medium, and the parental myeloma cells die. Another example of the use of selective media takes advantage of cell line deficiencies. Most of the mouse myeloma cell lines that are commonly used to generate hybridomas (P3X63-Ag8, NS-1-Ag4-1, X63-Ag8.653, and NS-0) are clonally derived from the MOPC21 tumor cell line P3 (Horibata and Harris, 1970), and they are all unable to synthesize cholesterol (Sato et al., 1984, 1987), which is an essential component of the plasma membrane. This trait is not a common characteristic of murine lymphoid cells or of mammalian cells in general, and it may have arisen from prolonged exposure to the antifungal agent amphotericin B (Sato et al., 1987). In all of these related myeloma cell lines, the defect in cholesterol biosynthesis has been traced to the enzyme 3-ketosteroid reductase (Sato et al., 1988). In cholesterol-free medium, NS-1 hybridomas are selected for while the cholesterol auxotrophic NS-1 parent cells are selected against (Myoken et al., 1989). One must bear in mind that it is usually only after a population of cells has been expanded in culture that they can be characterized with respect to their physiological or differentiated properties. At one time it was thought that cells normally dedifferentiated in vitro and that culture techniques could not be used to study differentiated cellular functions. Subsequently, it was found that the dedifferentiation phenomenon resulted from the overgrowth of differentiated cells by contaminating fibroblasts. Once this problem was recognized, the first

differentiated animal cell lines were established (Buonassisi et al., 1962; Yasumura et al., 1966). Thus, it is incumbent upon the cell culturist to choose culture media and culture conditions that not only support the viability and proliferation, if possible, of the cells of interest, but that allow those cells to manifest some or all of their differentiated properties in vitro. From a historical viewpoint, it is of interest that early culture media consisted of undefined mixtures of biological fluids, tissue extracts, and simple salt solutions. The first continuous mammalian cell line, the mouse L cell fibroblast line (Sanford et al., 1948), was established from 20-methylcholanthrene-treated C3H mouse tissue explants grown in chicken plasma clots in 40% horse serum, 20% chick embryo extract, and 40% saline. The first continuous human cell line, the HeLa cervical adenocarcinoma line (Gey et al., 1952), was isolated in a mixture of chicken plasma, bovine embryo extract, and human placental cord serum. At about the same time, attempts were being made to make culture media more defined by creating synthetic nutrient media and by determining the nutritional requirements of cells in culture. An early synthetic nutrient medium, medium 199 (Morgan et al., 1950), was created to increase the longevity of primary chicken muscle cell cultures that were started as tissue explants in Earle’s salt solution with 40% horse serum and 1% chick embryo extract. Medium 199 consisted of a salt solution with amino acids, vitamins, purines, pyrimidines, pentose sugars, adenosine triphosphate (ATP), adenylic acid, Tween 80 as a source of oleic acid, cholesterol, antioxidants, and iron in the form of ferric nitrate. Although medium 199 promoted the outgrowth of “large, flat and spindle-shaped” cells from the original tissue fragments, none of the medium components except glutamine clearly enhanced the life span of the cultures. The pioneering work of Eagle (Eagle, 1955) showed that L cells and HeLa cells had similar, demonstrable nutritional requirements for the thirteen essential amino acids, seven vitamins, glucose or other carbohydrates, and electrolytes. Strikingly, neither L cells nor HeLa cells would grow under these minimal essential conditions without the further addition of a small amount of dialyzed serum protein. Nonetheless, this research marked the beginning of concerted efforts to optimize basal nutrient media and to determine the growth requirements of cells in vitro. Culture media for a number of normal, immortalized, and transformed cells have been

Cell Culture


improved through the application of two complementary strategies. The approach of Ham and his colleagues has been to optimize the compositions of basal nutrient media for individual cell lines or cell types (Ham and McKeehan, 1979; Ham, 1984; Bettger and McKeehan, 1986) in the presence of ever-decreasing concentrations of dialyzed serum protein. These efforts gave rise to Ham’s F-12 nutrient medium and the MCDB media, which are commonly used today. MCDB media are optimized basal media developed for specific call types by R. Ham and colleagues. In studying hormonally responsive cell lines in culture, Sato and his colleagues realized that a major role of serum in culture medium was to provide hormones and hormone-like growth factors that were required for cell proliferation and expression of differentiated functions (Bottenstein et al., 1979; Barnes and Sato, 1980; Barnes, 1987). This understanding led them to replace serum with purified hormones, growth factors, transport proteins, and attachment factors as supplements for preexisting nutrient media. In combination, these two experimental approaches demonstrated (1) that basal nutrient media could be optimized for individual cell types, but optimal cell proliferation required additional hormones or growth factors, transport proteins, and attachment mediators in the absence of serum, and (2) that the combinations of purified medium supplements for individual cell lines could be simplified when using an optimized basal medium. General conclusions that can be drawn from the work of Ham and Sato are: (1) individual cell types require quantitatively balanced sets of nutrients, of which some are cell type specific; (2) cell proliferation and differentiated properties are regulated by hormones, growth factors, protein-bound nutrients, and attachment factors, of which many are present in serum and tissue extracts; (3) most cells in culture are growth stimulated by the serum components insulin and insulin-like growth factors, the iron-transporting protein transferrin, and unsaturated fatty acids or lipoproteins; and (4) because different cell types have similar but different growth requirements, it is unlikely that a single medium formulation will prove optimal for all cells.

Critical Parameters Media for Culture of Mammalian Cells

When choosing or developing a cell culture medium, the single most important parameter is cell viability. This holds true whether the medium is used to maintain a population of

differentiated cells, to stimulate cell proliferation, or to optimize the yield of a cellular product. An adequate serum-containing, serum-reduced, or serum-free medium formulation should promote a high degree of cell viability. Conversely, poor cell viability is a good indication that the culture medium or culture environment is deficient in one or more essential components. Suggestions for improving culture media are provided in Basic Protocols 1 and 2, and the reader is referred to UNIT 1.1 for methods of assessing cell viability. On occasion, changes in culture conditions, such as a switch from serum-supplemented to serumfree medium, may cause the majority of cells in a culture to die, followed by the outgrowth of a surviving subpopulation of cells. Although this phenomenon has been referred to in the literature as adaptation or weaning, it is more likely to be a selective process in which the surviving cells differ phenotypically from the parental population. The investigator can reduce the chances of phenotypic changes becoming fixed in a population of cells by maintaining cells in culture medium that supports a high level of viability, by using low split ratios when passaging cells, and by periodically returning to cryopreserved stocks of low-passage-number cells. The first choice for a basal nutrient medium should be one that other investigators have used successfully to culture the cells of interest and have reported in the literature. If for some reason that medium is not adequate for the purposes at hand, a number of basal media should be tested for the ability to support the proliferation of the cells of interest and to maintain their phenotypic properties. It is useful to start with basal media that have been used with similar or related cell types and in similar culture conditions (e.g., clonal or high-density cultures), but basal media developed for unrelated cell types may also yield good results (Ham, 1984). Commercially available basal media commonly used for continuous cell lines are DMEM; Ham’s F-12 medium; a 1:1 mixture (v/v) of DMEM and Ham’s F-12 medium (DMEM/F-12); and RPMI 1640, which was originally developed for lymphoid cells. The MCDB media were developed by Ham and his colleagues for individual types of normal cells, but they may also be effective on continuous cell lines. Most of these basal media are qualitatively similar but differ quantitatively. The basal medium selected based on empirical testing can be used as a starting point for further optimization. For excellent discussions on cel-


lular nutrition and procedures for optimizing basal nutrient medium see Ham and McKeehan, 1979; Ham, 1984; and Bettger and McKeehan, 1986. The most commonly used serum supplement is fetal bovine serum (FBS). Less expensive alternatives to FBS are calf serum, newborn calf serum, calf serum fortified with transferrin or growth factors (available from Hyclone and Sigma), and horse serum. For normal human lymphocytes, the use of commercial human serum treated at 56°C for 30 min to inactivate complement may be appropriate. The most suitable serum supplement for the cells of interest should be determined empirically. Although variability in efficacy between batches of FBS owing to variations in composition is not as problematic as it once was, it is advisable to test batches of FBS at several concentrations for the ability to support the proliferation of cells of interest at low and high densities. Clonal growth assays are the most stringent tests of the efficacy of batches of serum, but acceptable batches of serum should also be able to support, at reasonable concentrations, the viability of high-density cultures. A number of reduced-serum or serum-free media have been developed for continuous cell lines and nontransformed cells (Bottenstein et al., 1979; Jacoby and Pastan, 1979; Barnes et al., 1984; Freshney, 1987, 1992; Baserga, 1989; Davis, 1994; Sato et al., 1994), and most are optimized for a single cell line or cell type. However, the similarities between media developed for related cells are increasing the understanding of the nutritional and growth factor requirements of individual cell types, which in turn is making the development of serum-free medium a more rational process (Sato et al., 1994). As in choosing a basal nutrient medium for serum-supplemented medium, the best choice for a reduced-serum or serum-free medium is one that has been used by other investigators for the same or a related cell line or cell type. It is desirable to use a defined, serum-free medium whenever possible, as this affords the investigator the greatest degree of control over an in vitro culture−based experiment. Proprietary serum-free media for specific cell types are commercially available from companies such as Clonetics and Cell Systems (see Table 1.2.1). If an optimized, preexisting serum-free medium is not available, then a serum-free or serum-supplemented medium that supports cell viability and suboptimal proliferation can be used as a starting point for

further medium development. As described above (see Background Information), two complementary strategies for improving culture media are to optimize the components of the basal nutrient medium (Ham and McKeehan, 1979; Ham, 1984) and to replace serum or other undefined medium components with defined, purified protein and nonprotein supplements (Bottenstein et al., 1979; Barnes and Sato, 1980; Barnes, 1987; Sato et al., 1994). Under both strategies, concentrations of serum, dialyzed serum, or other undefined supplements are lowered stepwise to reduce the rate of cell proliferation, and the concentrations of defined medium components are individually manipulated until cell proliferation is restored. A completely defined culture medium or a much reduced serum-supplemented medium should be attained after a number of rounds of optimization. See Basic Protocol 2 for description of both these approaches. Clues as to which defined supplements are likely to be growth stimulatory for the cells of interest are provided by the following sources: hormones or growth factors that act on the cell type of interest in vivo; autocrine factors that the cells have been found to produce in vitro; and defined supplements that have been included in serum-free media developed for similar or related continuous cell lines or cell types. Thus, the first place to search for potentially useful supplements is the literature. The following general suggestions are based on the serumfree media that have been developed over the past 20 years. 1. Most cells are growth stimulated by insulin or insulin-like growth factor I and require iron obtained by the iron-transporting protein transferrin. 2. Fatty acid−free BSA is a useful carrier protein for unsaturated fatty acids, sterols, and steroid hormones, which are insoluble in an aqueous solvent. 3. Cells of epithelial origin often respond to epidermal growth factor (EGF), acidic fibroblast growth factor (aFGF or FGF-1), and dexamethasone or hydrocortisone. 4. Mesenchymal cells respond to EGF, aFGF, basic FGF (bFGF or FGF-2), and platelet-derived growth factor (PDGF). 5. In the absence of serum-derived attachment mediators, treating tissue culture plastic with attachment factors, such as type I collagen (UNIT 10.3), fibronectin, vitronectin, and laminin (UNIT 10.2), or with an incompletely defined, natural extracellular matrix (e.g., Matrigel, from Becton Dickinson Labware; UNIT 10.2) may

Cell Culture


enhance the plating efficiencies and growth rates of adherent cells. It is important to use highly pure water in preparing media and medium additives. Double-distilled water was a standard ingredient in medium for many years, but it has been superseded by purification systems that incorporate reverse osmosis. In the Milli-Q system, locally supplied water is subjected to deionization, reverse osmosis, and filtration through activated charcoal and a sterilizing filter. Use a vessel that is thoroughly rinsed with water after use but never washed with detergent.

Bottenstein, J., Hayashi, I., Hutchings, S., Masui, H., Mather, J., McClure, D.B., Ohasa, S., Rizzino, A., Sato, G., Serrero, G., Wolfe, R., and Wu, R. 1979. The growth of cells in serum-free hormone-supplemented media. Methods Enzymol. 58:94-109.

Anticipated Results

Eagle, H. 1973. The effect of environmental pH on the growth of normal and malignant cells. J. Cell. Physiol. 82:1-8.

As the survival and growth requirements of individual cell lines and cell types become better understood, the routine culture conditions for cells of interest become more defined. When all of the growth requirements of cells of interest are understood, any undefined medium supplements that were previously required can be completely eliminated. Defined culture conditions afford the investigator the greatest degree of control over in vitro culture experiments, and they provide more accurate insights into cellular physiology in vivo.

Time Considerations Optimizing a basal nutrient medium or developing a serum-free medium formulation is not a trivial undertaking, and it can be very time-consuming with no guarantee of success. Thus, the investigator should carefully consider how important defined culture conditions are to the experimental goals before taking on either task. However, as optimized and defined media are developed for a wider array of cell types, it is becoming easier and less time-consuming to create defined media for additional types of cells. The amount of time required to mix and sterilize 5- to 20-liter batches of medium should not exceed 2 to 4 hours.

Literature Cited Barnes, D.W. 1987. Serum-free animal cell culture. BioTechniques 5:534-541. Barnes, D.W. and Sato. G.H. 1980. Serum-free cell culture: A unifying approach. Cell 22:649-655. Barnes, D.W., Sirbasku, D.A., and Sato, G.H. (eds.) 1984. Cell Culture Methods for Cell Biology, Vols. 1-4. Alan R. Liss, New York. Baserga, R. (ed.) 1989. Cell Growth and Division: A Practical Approach. Oxford University Press, Oxford. Media for Culture of Mammalian Cells

Bettger, W.J. and McKeehan, W.L. 1986. Mechanisms of cellular nutrition. Physiol. Rev. 66:1-35.

Buonassisi, V., Sato, G.H., and Cohen, A.I. 1962. Hormone-producing cultures of adrenal and pituitary tumor origin. Proc. Natl. Acad. Sci. U.S.A. 48:1184-1190. Davis, J.M. (ed.) 1994. Basic Cell Culture: A Practical Approach. Oxford University Press, Oxford. Eagle, H. 1955. Nutrition needs of mammalian cells in tissue culture. Science 122:501-504.

Freedman, V.H. and Shin, S.-I. 1974. Cellular tumorigenicity in nude mice: Correlation with cell growth in semi-solid medium. Cell 3:355-359. Freshney, R.I. 1987. Culture of Animal Cells: A Manual of Basic Technique, 2nd ed. Alan R. Liss, New York. Freshney, R.I. (ed.) 1992. Culture of Epithelial Cells. Wiley-Liss, New York. Gey, G.O., Coffman, W.D., and Kubicek, M.T. 1952. Tissue culture studies of the proliferative capacity of cervical carcinoma and normal epithelium. Cancer Res. 12:264-265 (Abstr). Good, N.E., Winget, G.D., Winter, W., Connolly, T.N., Izawa, S., and Singh, R.M.M. 1966. Hydrogen ion buffers and biological research. Biochemistry 5:467-477. Ham, R.G. 1984. Formulation of basal nutrient media. In Cell Culture Methods for Cell Biology, Vol. 1 (D.W. Barnes, D.A. Sirbasku, and G.H. Sato, eds.) pp. 3-21. Alan R. Liss, New York. Ham, R.G. and McKeehan, W.L. 1979. Media and growth requirements. Methods Enzymol. 58:4493. Horibata, K. and Harris, A.W. 1970. Mouse myeloma and lymphomas in culture. Exp. Cell Res. 60:61-77. Jacoby, W.B. and Pastan, I.H. (eds.) 1979. Cell Culture. Methods Enzymol., Vol. 58. Kawamoto, T., Sato, J.D., McClure, D.B., and Sato, G.H. 1983. Development of a serum-free medium of growth of NS-1 mouse mycloma cells and its explication to the isolation of NS-1 hybridomes. Anal. Biochem. 130:445-453. Littlefield, J.W. 1964. Selection of hybrids from matings of fibroblasts in vitro and their presumed recombinants. Science 145:709-710. Morgan, J.F., Morton, H.J., and Parker, R.C. 1950. Nutrition of animal cells in tissue culture. 1. Initial studies on a synthetic medium. Proc. Soc. Exp. Biol. Med. 73:1-8. Myoken, Y., Okamoto, T., Osaki, T., Yabumoto, M., Sato, G.H., Takada, K., and Sato, J.D. 1989. An alternative method for the isolation of NS-1 hy-


bridomas using cholesterol auxotrophy of NS-1 mouse myeloma cells. In Vitro Cell Dev. Biol. 25:477-480. Perlman, D. 1979. Use of antibiotics in cell culture media. Methods Enzymol. 58:110-116. Sanford, K.K., Earle, W., and Likely, G.D. 1948. The growth in vitro of single isolated tissue cells. J. Natl. Cancer Inst. 9:229-246. Sato, J.D., Kawamoto, T., McClure, D.B., and Sato, G.H. 1984. Cholesterol requirement of NS-1 mouse myeloma cells for growth in serum-free medium. Mol. Biol. Med. 2:121-134. Sato, J.D., Kawamoto, T., and Okamoto, T. 1987. Cholesterol requirement of P3-X63-Ag8 and X63-Ag8.653 myeloma cells for growth in vitro. J. Exp. Med. 165:1761-1766. Sato, J.D., Cao, H.-T., Kayada, Y., Cabot, M.C., Sato, G.H., Okamoto, T., and Welsh, C.J. 1988. Effects of proximate cholesterol precursors and steriod hormones on mouse myeloma growth in serum-free medium. In Vitro Cell Dev. Biol. 24:1223-1228.

Sato, J.D., Hayashi, I., Hayashi, J., Hoshi, H., Kawamoto, T., McKeehan, W.L., Matsuda, R., Matsuzaki, K., Mills, K.H.G., Okamoto, T., Serrero, G., Sussman, D.J., and Kan, M. 1994. Specific cell types and their requirements. In Basic Cell Culture: A Practical Approach (J.M. Davis, ed.) pp. 181-222. Oxford University Press, Oxford. Southern, P.J. and Berg, P. 1982. Transformation of mammalian cells to antibiotic resistance with a bacterial gene under control of the SV40 early region promoter. J. Mol. Appl. Genet. 1:327-341. Yasumura, Y., Tashjian, A.H., Jr., and Sato, G.H. 1966. Establishment of four functional clonal strains of animal cells in culture. Science 154:1186-1189.

Contributed by J. Denry Sato Adirondack Biomedical Research Institute Lake Placid, New York Mikio Kan Texas A&M University Houston, Texas

Cell Culture


Aseptic Technique for Cell Culture

UNIT 1.3

This unit describes some of the ways that a laboratory can deal with the constant threat of microbial contamination in cell cultures. Microorganisms are ubiquitous. Bacteria can be isolated from nearly any surface including inanimate objects and human skin. Fungal spores and bits of vegetative hyphae drift into a laboratory from air conditioning ducts and open doors. Mycoplasma infections most frequently originate from improperly sterilized media or serum. At the risk of eliciting paranoia in the novice cell culture user who has no training in microbiological techniques, the possibility for microbial contamination exists everywhere. Inherent with successful manipulation of cell cultures is the basic understanding that everything that comes into contact with the cells must be sterile or noncontaminating. This includes media, glassware, and instruments, as well as the environment to which the cultures are briefly exposed during transfer procedures. Because cleaning up a contaminated culture is too frequently a disheartening and unsuccessful experience, the best strategy is to employ procedures to prevent microbial contamination from occurring in the first place. This unit begins with a protocol on aseptic technique (see Basic Protocol 1). This catch-all term universally appears in any set of instructions pertaining to procedures in which noncontaminating conditions must be maintained. In reality, aseptic technique cannot be presented in one easily outlined protocol, but rather encompasses all aspects of environmental control, personal hygiene, equipment and media sterilization, and associated quality control procedures needed to ensure that a procedure is, indeed, performed with aseptic, noncontaminating technique. Although cell culture can theoretically be carried out on an open bench in a low-traffic area, most cell culture work is carried out using a horizontal laminar-flow clean bench (see Basic Protocol 2) or a vertical laminar-flow biosafety cabinet (see Alternate Protocol). Subsequent units within this chapter address these diverse considerations—e.g., sterilization and disinfection, use of antibiotics, and quality control. Where applicable, use presterilized, disposable labware and other equipment. The wide availability and reliability of these products has simplified cell culture, particularly for small-scale laboratory needs. ASEPTIC TECHNIQUE This protocol describes basic procedures for aseptic technique for the novice in cell culture technology. One basic concern for successful aseptic technique is personal hygiene. The human skin harbors a naturally occurring and vigorous population of bacterial and fungal inhabitants that shed microscopically and ubiquitously. Most unfortunately for cell culture work, cell culture media and incubation conditions provide ideal growth environments for these potential microbial contaminants. This procedure outlines steps to prevent introduction of human skin flora during aseptic culture manipulations.

BASIC PROTOCOL 1

Every item that comes into contact with a culture must be sterile. This includes direct contact (e.g., a pipet used to transfer cells) as well as indirect contact (e.g., flasks or containers used to temporarily hold a sterile reagent prior to aliquoting the solution into sterile media). Single-use, sterile disposable plastic items such as test tubes, culture flasks, filters, and pipets are widely available and reliable alternatives to the laborious cleaning and sterilization methods needed for recycling equivalent glass items. However, make certain that sterility of plastic items distributed in multiunit packages is not compromised by inadequate storage conditions once the package has been opened.

Cell Culture Contributed by Rosalie J. Coté Current Protocols in Cell Biology (1998) 1.3.1-1.3.10 Copyright © 1998 by John Wiley & Sons, Inc.

1.3.1

Ideally, all aseptic work should be conducted in a laminar cabinet (see Basic Protocol 2 and Alternate Protocol). However, work space preparation is essentially the same for working at the bench. Flame sterilization is used as a direct, localized means of decontamination in aseptic work at the open bench. It is most often used (1) to eliminate potential contaminants from the exposed openings of media bottles, culture flasks, or test tubes during transfers, (2) to sterilize small instruments such as forceps, or (3) to sterilize wire inoculating loops and needles before and after transfers. Where possible, flame sterilization should be minimized in laminar-flow environments as the turbulence generated by the flame can significantly disturb the sterile air stream. Materials Antibacterial soap 70% ethanol or other appropriate disinfectant 95% ethanol Clean, cuffed laboratory coats or gowns Latex surgical gloves Clean, quiet work area Shallow discard pans containing disinfectant Bunsen burner or pilot-activated burner (e.g., Touch-o-Matic, VWR) Take personal precautions 1. Just prior to aseptic manipulations, tie long hair back behind head. Vigorously scrub hands and arms at least 2 min with an antibacterial soap. Superficial lathering is more prone to loosening than removing flaking skin and microbial contaminants. Loosely adhering skin flora easily dislodge and can potentially fall into sterile containers.

2. Gown appropriately. For nonhazardous sterile-fill applications, wear clean, cuffed laboratory coats and latex gloves. Greater stringencies may be necessary depending upon laboratory regulatory requirements. Work with potentially hazardous agents certainly mandates additional considerations for safety. Front-closing laboratory coats are not recommended for work with hazardous biological agents. Safety glasses should be worn by laboratory personnel when manipulating biological agents outside the confines of a biosafety cabinet.

3. Frequently disinfect gloved hands with 70% ethanol while doing aseptic work. Although the gloves may initially have been sterile when first worn, they will no doubt have contacted many nonsterile items while in use. Note that 70% ethanol may not be an appropriate agent for latex glove disinfection when working with cultures containing animal viruses, as studies have shown that ethanol increases latex permeability, reducing protection for the wearer in the event of exposure. In this case, quarternary ammonium compounds are more appropriate.

4. Dispose of gloves by autoclaving after use. Do not reuse. Bag and autoclave single-use laboratory coats after use. Bag, autoclave (if necessary), and wash other laboratory coats within the laboratory facility or send out for cleaning at a laundry certified for handling biologically contaminated linens. Never take laboratory clothing home for washing.

5. Thoroughly wash hands after removing protective gloves. Aseptic Technique for Cell Culture


Prepare and maintain the work area 6. Perform all aseptic work in a clean work space, free from contaminating air currents and drafts. For optimal environmental control, work in a laminar-flow cabinet (see Basic Protocol 2 and Alternate Protocol). 7. Clear the work space of all items extraneous to the aseptic operation being performed. 8. Wipe down the work surface before and after use with 70% ethanol or other appropriate disinfectant. 9. Wherever feasible, wipe down items with disinfectant as they are introduced into the clean work space. Arrange necessary items in the work space in a logical pattern from clean to dirty to avoid passing contaminated material (e.g., a pipet used to transfer cultures) over clean items (e.g., flasks of sterile media). 10. Immediately dispose of any small contaminated items into a discard pan. 11. When the aseptic task has been completed, promptly remove any larger contaminated items or other material meant for disposal (e.g., old culture material, spent media, waste containers) from the work space and place in designated bags or pans for autoclaving. Disinfect the work space as in step 8. Flame sterilize the opening of a vessel 12. For a right-handed person, hold the vessel in the left hand at ∼45° angle (or as much as possible without spilling contents) and gently remove its closure. Do not permit any part of the closure that directly comes in contact with the contents of the vessel to touch any contaminating object (e.g., hands or work bench). Ideally, and with practice, one should be able to hold the closure in the crook of the little finger of the right hand while still being able to manipulate an inoculating loop or pipettor with the other fingers of the hand. Holding the vessel off the vertical while opening will prevent any airborne particulates from entering the container.

13. Slowly pass the opening of the vessel over the top of (rather than through) a Bunsen burner flame to burn off any contaminating matter. Be careful when flaming containers of infectious material. Any liquid lodged in the threads of a screw cap container will spatter as it is heated. Aerosols thus formed may actually disseminate entrapped biological agents before the heat of the flame is hot enough to inactivate them.

14. While still holding the vessel at a slant, use a sterile pipet and pipettor to slowly add or remove aliquots to avoid aerosol formation. 15. Flame-sterilize again as in step 13, allow the container to cool slightly, and carefully recap the vessel. Flame sterilize small hand instruments 16. Dip critical areas of the instrument (i.e., those that come into contact with the material of concern) in 95% ethanol. Make certain that the alcohol is in a container heavy enough to support the instrument without tipping over. CAUTION: 95% ethanol is flammable; keep the container at a safe distance from any open flame. Cell Culture


17. Remove the instrument from the alcohol, being careful not to touch the disinfected parts of the instrument. Allow excess ethanol to drain off into the container. 18. Pass the alcohol-treated part of the instrument through the flame of a Bunsen burner and allow residual alcohol to burn off. 19. Do not let the sterilized portion of the instrument contact any nonsterile material before use. Let the heated part of the instrument cool for ∼10 sec before use. 20. After use, return the instrument to the alcohol disinfectant until needed again. Flame sterilize inoculating loops and needles 21. Hold the inoculating wire by its handle and begin in the center of the wire to slowly heat the wire with the flame of a Bunsen burner. Proceed back and forth across the wire’s full length until it glows orange. 22. While still holding the handle, allow the inoculating wire to cool back to room temperature (∼10 sec) before attempting any transfer of material. If tranfers are made while the inoculating wire is hot, cells will be killed by the hot wire, and aerosols created from spattering material can disperse biological material throughout the work space.

23. After the transfer is made, reheat the inoculating wire as in step 21 to destroy any remaining biological material. Let cool to room temperature before putting aside for next use. BASIC PROTOCOL 2

USE OF THE HORIZONTAL LAMINAR-FLOW CLEAN BENCH Laminar-flow cabinets (hoods) are physical containment devices that act as primary barriers either to protect the material being manipulated within the hood from workergenerated or environmental sources of contamination, or to protect the laboratory worker and laboratory environment from exposure to infectious or other hazardous materials that are present within the hood. Cell culture applications utilize two types of laminar-flow hoods: (a) the horizontal-flow clean bench (described here) and (b) the biological safety cabinet (see Alternate Protocol). Both types of hoods use a high-efficiency particulate air (HEPA) filter and blowers that generate a nonmixing stream of air. The horizontal laminar-flow clean bench is used to provide a near-sterile environment for the clean (i.e., noncontaminating) handling of nonhazardous material such as sterile media or equipment. Because the air stream pattern directs the flow of air within the hood directly back to the hood operator and the room (Fig. 1.3.1), horizontal flow hoods are never to be used with infectious agents or toxic chemicals. Materials 70% ethanol or other disinfectant Horizontal laminar-flow hood, certified for use Swabs (e.g., cheesecloth, paper towels) Pilot light–activated Bunsen burner (e.g., Touch-o-Matic, VWR) 1. Completely clear the bench of the laminar-flow hood and disinfect the bench working surface and the left and right sides of the hood with 70% ethanol or other disinfectant. Do not spray the back (gridded) wall where the HEPA filter is housed.


Resist the urge to leave frequently used items (e.g., pipet canisters or a bag of disposable plastic tissue culture flasks) in the hood between uses. Their presence makes thorough disinfection of the work space difficult.


HEPA filter HEPA-filtered air

Figure 1.3.1 Horizontal laminar-flow clean cabinet. Solid arrows, dirty room air; open arrows, clean HEPA-filtered air; circled +, positive pressure with respect to room air.

room air

blower

prefilter

2. Turn the hood blower and lights on and let the air circulate within the hood 10 min before use. 3. Place items needed for the specific procedure into the hood, wiping each item with 70% ethanol or other disinfectant just before introducing it into the laminar environment. Do not overcrowd the work space. For horizontal laminar-flow effectiveness, maintain a clear path between the work area and the back wall of the cabinet where the HEPA filter is located.

4. Wash hands well before working in the hood and wear a clean laboratory coat and surgical gloves to further protect the work from shedding of skin flora that can contaminant any product (see Basic Protocol 1). 5. While working in the hood, perform all work at least 4 in. back from the front opening, and avoid rapid movements that might disrupt the laminar air flow. Avoid moving materials or hands in and out of the cabinet as much as possible. 6. If flame sterilization is needed in the hood for a particular application, use a burner that can be activated by a pilot light when needed, rather than one that burns constantly. The open flame of a Bunsen burner causes turbulence that disrupts the unidirectional laminar air flow.

7. When work is completed, remove all material from the laminar work bench, clean any spills, and disinfect the bench working surface by wiping with 70% ethanol or other disinfectant. 8. Turn off hood blower and lights.

Cell Culture


ALTERNATE PROTOCOL

USE OF THE VERTICAL LAMINAR-FLOW BIOSAFETY CABINET Biological safety cabinets provide a clean, safe environment for both the worker and the product. The Class II, Type A biosafety cabinet (Fig. 1.3.2) is frequently encountered in cell culture laboratories, and this protocol describes the use of this type of barrier device. The Class IIA biosafety cabinet is suitable for work with low- to moderate-risk biological agents in the absence of toxic or radioactive chemicals. Materials (also see Basic Protocols 1 and 2) Class II, Type A Biosafety Cabinet (BSC), certified for use Pilot light–activated Bunsen burner (e.g., Touch-o-Matic, VWR) or electronic incinerator (e.g., Bacti-Cinerator III, VWR) Closed-front laboratory gowns (for personnel working with biological agents) 1. Turn the hood blower on and verify air flow by feeling (by hand) the current near the front grill of the work surface. Turn the germicidal UV light off if it is on. Turn the fluorescent light on. Before use, the cabinet should already be empty and clean from prior activity. The view window should be lowered to the proper operating height (normally 8 in.) or as specified by the cabinet manufacturer. UV light is effective only for decontaminating clean, solid surfaces with which it comes in contact. It is not effective in decontaminating the cabinet air flow. UV light is not effective against bacterial spores. UV germicidal light tubes should be replaced frequently (at least every 6 months for biosafety cabinets in use on a daily basis) to assure that they are emitting light at 254 nm and at an intensity appropriate for decontamination. CAUTION: UV light is harmful to the eyes. Laboratory personnel should not be near the cabinet or looking at the UV light when it is in use.

2. Wash and gown as required for the operation (see Basic Protocol 1, steps 1 to 5). 3. Wipe down the entire interior cabinet work surface area with 70% ethanol or other appropriate disinfectant. 4. Let blower run for 10 min to filter the cabinet air of any particulates.

exhausted air (30% of recycled air) HEPA filters

view window filtered air (70% of recycled air)

Figure 1.3.2 Biological safety cabinet, Class II, Type A. Note that filtered air is contaminated after passing through the work space, and is filtered again whether it is recycled to the workspace (70%) or exhausted (30%). Solid arrows, dirty (room/contaminated) air; open arrows, filtered air.

blower Aseptic Technique for Cell Culture


5. Raise the front view window as needed to bring necessary items into the cabinet. Wipe each item with 70% ethanol or other disinfectant as it is placed in the cabinet. Do not crowd the work space and make sure no air vents are blocked by supplies or equipment. Do not position material so that it obscures any of the air vents at the front edges of the laminar hood. One frequent source of air flow restriction in biosafety cabinets is “lost” paper towels that have been drawn into the air ducts at the back of the work surface.

6. Organize the work surface for a clean-to-dirty work flow. Place clean pipets, flasks, and sterile media bottles at one side of the cabinet; place discard pans, spent cultures, and other wastes on the other side. 7. Return the view window to the 8-in. operating level. Wait ∼10 min for the blowers to filter the disturbed cabinet air before starting work. 8. While working, keep all material and perform work ≥4 in. back from the front opening of the cabinet, and minimize rapid movements or activity. Keep the view window opening as close to 8 in. as allows reasonable access to the work surface and equipment. These precautions assure that any drafts caused by arm movements will not disrupt air flow or churn room air currents into the clean work area.

9. If direct flame sterilization of items within the cabinet is necessary, use an electric burner or pilot light–activated flame burner located at the back of the work space. A constant open flame in the cabinet can disturb the laminar air flow.

10. At the end of the procedure, enclose all contaminated materials. Clean the cabinet work surface with 70% ethanol or other disinfectant, being especially careful to wipe any spills of culture suspensions or media that can serve as future contamination points. Clear all material from the cabinet. 11. Let the blower run for ≥10 min with no activity to remove any aerosols that were generated. During this period, turn off the fluorescent light and turn on the germicidal UV light. Allow the UV light to operate ≥30 min. COMMENTARY Background Information Aseptic technique The dictionary definition of asepsis simply implies freedom from pathogenic organisms. However, the practical definition of the term for cell biologists, as well as other biotechnologists working with pure cultures, has come to be synonymous with sterile or noncontaminating conditions. The successful manipulation of cell cultures under any circumstance inherently relies upon the ability to maintain rigorous aseptic (i.e., noncontaminating) working conditions. The concept of aseptic technique is simple in theory: prevention of sterile or uncontaminated material and objects from coming into contact with any nonsterile or contaminated material. Practical application of the theory is often illusive for beginning students. However,

breaches in aseptic technique can also cause significant problems for even well-experienced laboratories, particularly when the source of contamination is not readily evident. A single incident of culture contamination is frustrating in its own right, but repeated contamination (particularly by the same type of organism) invariably results in expensive losses and delays until the localization and source are identified. The critical areas of concern with respect to successful aseptic technique include environmental conditions (laboratory or work space), source material (cell lines, media, and reagents), equipment (labware, instruments, and apparatuses), sterilization procedures and equipment (autoclave, dry heat, filtration), and human (laboratory personnel) considerations. Budgetary constraints aside, technological aids exist to greatly simplify the hardware needed

Cell Culture


for aseptic work. Laminar-flow cabinets create clean working environments (see below); clean, certified cell lines are available from cell repositories; media manufacturers and biotechnology supply companies provide sterile media, sera and reagents; and presterilized disposable labware to satisfy most cell culture needs is available from any large distributor of scientific supplies. Despite all the technological advances, the one weak link remaining in successful laboratory applications of aseptic technique is the human factor. Too frequently, contamination occurs because of the desire to work a little too quickly, the urge to eliminate an “unimportant” step, or lapses in concentration during mundane procedures. The only advice to offer as protection against the human factor is to work slowly and deliberately when performing procedures under aseptic conditions, don’t eliminate procedural steps, and pay attention! Establishing a standard routine of procedures and of placement of materials can help prevent the omission of steps.


Laminar-flow cabinets Laminar-flow cabinets or hoods have replaced the open laboratory bench for aseptic work in almost all cell culture and microbiology laboratories. Their effectiveness as physical barriers to contamination relies on a cabinet design incorporating high-efficiency particulate air (HEPA) filters to trap airborne contaminants, and blowers to move the filtered air at specified velocities and in a nonmixing (laminar) stream across a work surface. As noted in each protocol for the particular type of laminar-flow application, the proper choice of cabinet is imperative. Horizontal laminar-flow cabinets are never used with biological or toxic chemical agents as they are not containment devices but rather serve to provide a strong stream of near-sterile air for particlefree working conditions. As this air is blown directly from the HEPA filter (at the back of the cabinet) across the work surface and out of the cabinet (directly into the operator’s face and the room), the restricted use of the horizontal flow cabinet to nonhazardous material is obvious. The Class IIA biosafety cabinet is a laminar containment device that (1) protects the material being manipulated within the cabinet by HEPA-filtered incoming air and (2) protects the operator and room environment from potentially hazardous material in the cabinet with an air curtain at the front of the cabinet (the view screen) and HEPA-filtered cabinet exhaust air.

As Class IIA biosafety cabinets are not totally leak-proof, they cannot be used for high-risk biological agents (see current Center for Disease Control and NIH guidelines for the status of any biological material used in the laboratory; Richmond and McKinney, 1993). Because Class IIA cabinets operate with ∼70% recirculated air within the cabinet (Fig. 1.3.2), the potential for accumulation of chemicals within the laminar work space limits use to low-level toxic or radioactive material. Laminar-flow cabinets are not replacements for good microbiological aseptic technique and must be used in conjunction with standard concerns for asepsis if full efficiency of the equipment is expected. Similarly, there is a limit to the protection a laminar cabinet can provide if it is operated in an environment not conducive to clean work conditions. The cabinets should be installed and operated in a relatively clean, quiet laboratory environment. Laboratory doors should be kept closed while the cabinet is in use to minimize strong room air currents that could break the laminar air stream within the cabinet. The units should not be located directly near room air ducts or anywhere a strong environmental air flow exists. Additionally, air flow disturbance by personnel or equipment, particularly within a few feet in front of the cabinets, should be limited when the laminar device is in use. Because of the critical nature of their function (particularly for the biosafety laminar cabinet), these devices must be certified at installation by professional laminar flow technicians in accordance with National Sanitation Foundation Standard No. 49 for Class II (laminar flow) Biohazard Cabinetry (NSF International, 1992) or other applicable regulatory and safety guidelines. As HEPA filters are brittle and will crack with normal usage of the unit, laminar cabinets must also be recertified annually or after 1000 hr use, and whenever they are moved.

Critical Parameters and Troubleshooting Human sources of contamination As noted above, bacterial shedding from human skin is a natural occurrence. However, under times of physiological or psychological stress, a human may shed so excessively that routine gowning procedures are inadequate. A clue to this condition can be the veteran technician who suddenly can’t seem to transfer anything without contaminating it, especially when contamination is repeatedly bacterial and


by species of Staphylococcus, Micrococcus, or coryneforms. Alleviation of the problem may be achieved by simply controlling the temperature of the laboratory. Gowned personnel sweat in 27°C (80°F) rooms, and people who sweat shed more than people who don’t. Rigorous attention to gowning details as well as liberal washing of hands and arms with an antimicrobial soap just prior to aseptic work may alleviate the situation. If the problem involves psychological stress or physiological stress due to illness or medication, more rigorous gowning procedures may help. Use fresh, clean laboratory coats for each round of aseptic work and make sure laboratory coat sleeves are tucked inside gloves to prevent exposed wrists. Use disinfectants liberally. For worst-case incidences of excessive shedding, the only recourse may be to move the individual to nonaseptic procedures until the condition clears. Decontamination of a laminar-flow cabinet Any mechanical failure of a laminar cabinet must be evaluated by qualified, trained personnel. Increased incidences of microbial contamination (particularly by the same organism) could originate from (1) poor cleaning and disinfection of the cabinet work space, (2) a source of contamination lodged in the ducts within the cabinet (e.g., media or culture material spilled into the cabinet ducts), or (3) a crack in the HEPA filter. Disinfect the catch basin if culture material has spilled through the vents in the work surface into the catch basin below. Use a strong disinfectant (such as 5% to 10% bleach in a sufficient volume to thoroughly contact the spilled material) and allow the disinfectant to stay in contact with the spill for 30 min. Drain the contents of the catch basin into a container suitable for final sterilization by autoclaving. Visually inspect the working interior of the laminar cabinet for evidence of dried culture material or media, especially in the corners of the cabinet. Clean the interior of the cabinet with a laboratory detergent, rinse with water, dry, and treat the area with an appropriate disinfectant. Be very careful not to wet the exposed HEPA filter located on the back wall of horizontal flow cabinets, as this can compromise the filter integrity. Be careful not to let cleaning solutions enter any vents of the cabinet. After thorough cleaning of the cabinet work surface, operate the cabinet (as detailed in Basic Protocol 2 or Alternate Protocol) using a control procedure for localizing the source of any

remaining contamination. This can be achieved with a series of opened plates of trypticase soy agar and Emmons’ modification of Sabouraud’s agar systematically coded and placed across the work surface. Leave the media plates open and the cabinet operating for 30 min. Close the lids of the agar plates and incubate them at 26°C for 5 days. If significant microbial contamination appears in the plates, consult with a qualified laminar technician. The resolution to the problem will require either caulking leaks in the HEPA filter or sealing the cabinet for total interior decontamination of filter and ducts with formaldehyde gas. A final source of frequent contamination in a laminar working condition can be the “sterile” equipment, labware, or solutions used. A poorly filter-sterilized phosphate-buffered saline solution can give rise to significant numbers of pseudomonad bacteria within weeks when stored at room temperature. Insufficiently processed autoclaved or dry heat–sterilized labware frequently results in contamination of cell culture material by spore-forming bacteria.

Anticipated Results When proper aseptic techniques are used, it should be possible to maintain cell cultures without contamination.

Time Considerations

It takes ∼1/2 hr to properly prepare oneself and the cell culture area for culture procedures and a similar amount of time to properly clean up afterward.

Literature Cited Richmond, J.Y. and McKinney, R.W. (eds.) 1993. Biosafety in microbiological and biomedical laboratories, 3rd ed. U.S. Government Printing Office, Washington, D.C. NSF (National Sanitation Foundation) International. 1992. Class II (laminar flow) biohazard cabinetry (NSF 49-1992). NSF International, Ann Arbor, Mich.

Key References Barkley, W.E. and Richardson, J.H. 1994. Laboratory safety. In Methods for General and Molecular Bacteriology, 2nd ed. (P.E. Gerhardt, R.G.E. Murray, W.A. Wood, and N.R. Krieg, eds.) pp. 715-734. American Society for Microbiology, Washington, D.C. Chapter provides an overview of general concerns for working with biological agents, from a classic publication on general methods in bacteriology that often overlaps to satisfy the technical needs of cell biologists. Cell Culture


Chatigny, M.A. 1986. Primary barriers. In Laboratory Safety: Principles and Practices (B.M. Miller, D.H.M. Gröschel, J.H. Richardson, D.Vesley, J.R. Songer, R.D. Housewright, and W.E. Barkley, eds.) pp. 144-163. American Society for Microbiology, Washington, D.C. Offers detailed considerations on the types and uses of laminar-flow barrier technology. The main publication is well worth its price for anyone (staff, supervisors, administrators) responsible for safety in a biological laboratory.

Freshney, R.I. 1994. Culture of Animal Cells: A Manual of Basic Technique, 3rd ed., pp. 51-69. Wiley-Liss, New York. Offers suggestions for maintaining aseptic conditions while working with cell cultures. A classic cell culture publication that surveys the field while providing enough detail for an individual with intermediate knowledge of microbiology and cell biology.

Contributed by Rosalie J. Coté Becton Dickinson Microbiology Systems Sparks, Maryland



Sterilization and Filtration

UNIT 1.4

This unit describes conditions and procedures for the use of the autoclave and the convection or gravity oven for sterilization of heat-stable laboratory materials, for depyrogenation by heat, and for decontamination of biological waste. Sterilization is not an absolute but rather a probability function. Terminal sterilization processes such as autoclaving or dry heat should have a 10−6 or less probability that an organism will survive treatment. The proper choice of sterilization method, well-maintained equipment, validated procedures, and adherence to protocol are all necessary to keep the statistics in one’s favor. This unit includes protocols for a variety of sterilization and decontamination methods. Moist-heat or steam sterilization is used for liquids, dry goods, and decontamination of biological wastes. Dry heat or depyrogenation is used to sterilize laboratory glassware and equipment. The efficacy of sterilization using these methods should be monitored using biological indicators. Disinfectants such as ethanol, quaternary ammonium compounds, and sodium hypochlorite are used for decontamination of facilities and equipment, and for clean up of certain spills. Vacuum or positive-pressure filtration is an alternative method for sterilization of liquids that do not withstand steam sterilization. AUTOCLAVING LIQUIDS The autoclave is used for sterilization by moist heat. The standard conditions for moist-heat sterilization are exposure to saturated steam under pressure at 121°C for 15 min, although other temperature/time specifications can be utilized for specialized needs. In general, materials suitable for autoclaving as a nondestructive sterilization process must meet the following criteria: (1) stable to the temperature and time of the autoclave cycle, (2) unaffected by moisture, (3) packaged to permit exposure to steam, and (4) hydrophilic, if liquid. Materials are sterilized by autoclaving only if they are wetted with the steam; thus, sealing gaskets on certain types of laboratory equipment may not be effectively sterilized if tightened in place during the autoclave cycle. Materials can be decontaminated by autoclaving providing criteria (3) and (4) are met.

BASIC PROTOCOL 1

The autoclave cycle is based on the time it takes the material being sterilized to be in contact with saturated steam at 121°C for 15 min, and not the time the autoclave itself has been selected to run at that temperature. As autoclave efficiency is machine specific, and steam penetration is container and volume dependent, autoclave cycles and load configurations should ideally be validated to assure that sterilization conditions are achieved. For many small laboratories, the purchase expense of temperature-monitoring thermocouples might be prohibitive. However, frequent use of biological indicators to monitor autoclave conditions (see Support Protocol 1) is strongly recommended for all laboratories, particularly when the machine is used for decontamination of biological waste. Table 1.4.1 lists suggested autoclave times for load configurations in an autoclave with a 20 × 20 × 38–in. (51 × 51 × 97–cm) chamber. Materials Heat-resistant containers and vessels (e.g., borosilicate glass, high-grade stainless steel, noncytotoxic plastic) Liquid to be autoclaved Moisture-resistant labels Paper or aluminum foil Autoclave indicator tape Cell Culture Contributed by Rosalie J. Coté Current Protocols in Cell Biology (1999) 1.4.1-1.4.21 Copyright © 1999 by John Wiley & Sons, Inc.

1.4.1 Supplement 1

Autoclave Autoclavable discard pans 1. Use a heat-resistant vessel that can hold twice the volume of the liquid to be autoclaved to assure that boiling encountered during the heating and cooling periods of the autoclave cycle do not result in a boil-over of the vessel contents. The most frequent laboratory frustrations with autoclaves and sterilization of liquids are boil-over of material or blow-off of closures. When volume-to-container relationships are acceptable, and when the solution is not overly viscous, the problem most often occurs because the autoclave’s slow exhaust or liquid cycle is not properly functioning and the machine is exhausting the chamber pressure too quickly. Adjustments or repairs to the unit should be made by qualified personnel.

2. Fill the vessel to desired volume with liquid to be autoclaved. Indicate the contents of the container with permanent ink and a moisture-resistant label. 3. Loosely cover any opening to the vessel. Do not overtighten screw-cap closures, as this prevents adequate pressure/steam exchange. Overwrap and secure cottonplugged flasks with paper or aluminum foil to prevent plugs from blowing off if the autoclave is exhausted too rapidly. 4. Affix a piece of autoclave indicator tape to each item or package, as a visual reference (following the autoclave cycle) that the material has been processed. The color change of autoclave tape indicates only that the tape has been exposed to a 121°C temperature, and not how long it has been held at that temperature. Thus, it is not an indicator of successful sterilization.

5. Load the autoclave with vessels of similar size, volume, and configuration. Place all liquid-containing vessels inside autoclavable discard pans inside the autoclave. Close and lock autoclave door. The discard pans should be large enough to contain all fluid or glass in the event of boiling over or breakage during autoclaving.

6. Set autoclave controls for liquids or slow exhaust. 7. Select and set autoclave controls for appropriate sterilization time (see Table 1.4.1).

Table 1.4.1 Suggested Autoclave Run Times and Configurations for 121°C Sterilization in a 20 × 20 × 38–in. Autoclavea

Size

Volume (ml)

Minimum time (min)b

Maximum time (min)b

13 × 100 mm 16 × 125 mm 20 × 150 mm

4-6 5-10 12-20

18 18 18

20 20 20

Flasks

100 ml 250 ml 500 ml 1000 ml 2000 ml

25-50 75-100 250 500 1000

20 24 26 28 30

26 28 30 32 32

Media bottles

125 ml 500 ml All

50 250-500 All

20 30 35

22 32 90

Vessel Test tubes

Empty glassware Sterilization and Filtration

aChamber dimensions (metric equivalent, 51 × 51 × 97 cm). bSterilization times indicated are actual autoclave timer settings. Material within the vessels will be exposed to 121°C for 15 min using these process times. See text for further explanantion.

1.4.2 Supplement 1


8. Start and run autoclave cycle to completion. Any interruption in the cycle (e.g., a sudden drop in chamber pressure) invalidates the run, and the sterilization cycle must be rerun to assure efficacy of the process. This is problematical if the liquid has limited stability to prolonged or repeated autoclaving, such as the microbiological media used for quality control in cell culture work. In such cases, it is best to start over with new media.

9. Open the autoclave door only when chamber pressure registers 0 lb/in2 (100°C or less). CAUTION: Never stand in the path of escaping steam when opening an autoclave.

10. Remove flasks or containers only when all bubbling has stopped. CAUTION: Superheated liquids can easily boil violently if even slightly jostled. Resulting boil-over can badly scald laboratory personnel.

11. Cool vessels to ambient or other prescribed temperature in a relatively clean area not subject to excessive air currents. This helps avoid suctioning of heavily contaminated environmental air into the container, which can sometimes occur when a vacuum forms within the container as the liquid cools.

AUTOCLAVING DRY GOODS Heat-stable dry materials (including stainless steel instruments, glassware, fabrics, and plasticware) can be effectively sterilized by autoclaving, providing all surfaces of the dry material come in contact with the saturated steam at 121°C. This can become problematical for small items (such as forceps) that must be packaged in an outer container or wrapping that impedes the flow of steam, or for folded fabrics that tend to harbor pockets of cooler air. For this reason, autoclaving times for dry goods sterilization often rely on overkill, as these materials generally have much higher heat resistance. As noted above, validation studies should be done to determine the most effective times and configurations for a given autoclave. For further information on general autoclaving considerations, see Basic Protocol 1 introduction. For default times for an autoclave with a 20 × 20 × 38–in. (51 × 51 × 97–cm) chamber, see Table 1.4.1.


Additional Materials (also see Basic Protocol 1) Items to be autoclaved Shallow heat-resistant container 1. Loosely arrange small items in a shallow, heat-resistant outer container. Loosely cover the outer container’s opening with paper or aluminum foil. If container has its own lid, apply it loosely so that steam and pressure can penetrate. Cap larger items such as bottles or flasks, making sure that all closures (e.g., screw caps) are loose enough to permit penetration of pressurized steam. Small items can also be individually wrapped in paper or foil.

2. Code each item or package with permanent ink and a moisture-resistant label identifying its contents. Affix a piece of autoclave indicator tape to each item or package as a visual reference (following the autoclave cycle) that the material has been processed. The color change of autoclave tape indicates only that the tape has been exposed to a 121°C temperature, and not how long it has been held at that temperature. Thus, it is not an indicator of successful sterilization. Cell Culture


Supplement 1

3. Where feasible, add a small amount of deionized or distilled water to the outer container or the individual items to assure adequate moisture for effective sterilization. 4. Arrange material in the autoclave to avoid dense overpacking that will impede effective sterilization. Where possible, arrange items to permit downward displacement of cooler, heavier air (e.g., place empty bottles or flasks on their sides rather than upright in the autoclave). This prevents pockets of cool air from being trapped in the bottom of the containers as the hot, pressurized steam flows into the vessels.

5. Set autoclave for a fast exhaust cycle. If the sterilizer is so equipped, a drying cycle that removes moisture from the dry goods under vacuum at the end of the timed sterilization run can also be used.

6. Select and set autoclave controls for appropriate sterilization time (see Table 1.4.1). 7. Start and run autoclave cycle to completion. Note that any interruption in the cycle (e.g., a sudden drop in chamber pressure) invalidates the run, and the sterilization cycle must be rerun to assure efficacy of the process.

8. Open the autoclave door to remove items only when chamber pressure registers 0 lb/in2 (100°C or less). CAUTION: Never stand in the path of escaping steam when opening an autoclave. ALTERNATE PROTOCOL 2

AUTOCLAVING FOR DECONTAMINATION OF BIOLOGICAL WASTE Biological laboratory waste is most frequently decontaminated by autoclaving unless it contains hazardous chemical materials that can volatilize in the sterilization process. In many mid- to large-sized laboratories, biological waste includes varying combinations of spent media, discarded cultures, and solid material. An autoclave load size or configuration can vary dramatically with each run. For this reason, autoclave cycles for decontamination most often employ the overkill approach. Validation studies prior to actual-use procedures must be performed to assure that selected operation procedures are adequate to achieve the desired conditions for successful decontamination. For further information on general autoclaving considerations, see Basic Protocol 1 introduction. CAUTION: Do not dispose of biological material containing hazardous chemicals or radioactive isotopes in the waste stream designated for autoclaving. Additional Materials (also see Basic Protocol 1) Items for decontamination Plastic (polyethylene or polypropylene) autoclavable bags for biohazardous waste 1. Place items for decontamination into plastic autoclavable bags clearly labeled as containing biohazardous material. For greater ease with postautoclaving cleanup procedures, segregate plastic disposable material from reusable labware in separate bags. CAUTION: For the safety of laboratory personnel who sort the autoclaved waste for washing or disposal, do not dispose of sharps or pipets as loose items in the bags. These items must be segregated in their own containers (containing a disinfectant, if appropriate).


2. Add ∼500 ml water to bags containing only dry items (such as empty glassware or contaminated lab coats) to supply sufficient moisture for steam generation.

1.4.4 Supplement 1


3. Support bags by placing them in large, shallow, leak-proof, autoclave-resistant discard pans to prevent tearing of the bags and release of contents. For the safety of personnel carrying the discard pans, use a maximum weight limit of 25 lb per pan.

4. Securely seal each bag to prevent leakage of material. Transport biological waste only in closed containers. 5. Code each bag with permanent ink and a moisture-resistant label to permit general identification of its contents or source-lab should an accident occur (e.g., mycoplasma testing lab; QC lab). 6. Affix a piece of autoclave indicator tape to each item or package as a visual reference (following the autoclave cycle) that the material has been processed. The color change of autoclave tape indicates only that the tape has been exposed to a 121°C temperature, but does not indicate the time held at that temperature. Thus, it is not an indicator of successful sterilization.

7. Transport the waste using a sturdy laboratory cart or autoclave carriage and dolly. 8. Load the discard pans loosely into the autoclave to allow steam to flow over and around the material. While loading, slit each individual bag open in several spots to allow direct exposure of the contents of the bags to the steam from the autoclave. Do not stack pans directly on top of each other.

9. Close and lock autoclave door. 10. Run autoclave cycle for 90 min at 121°C on a fast-exhaust, gravity cycle. The cycle time listed here is for a large (24 × 36 × 48–in.; 61 × 91 × by 122–cm) autoclave. With small loads and smaller-chamber autoclaves, 45 min may be sufficient. Any interruption in the cycle (e.g., a spurious drop in autoclave steam pressure) invalidates the run.

11. Open the autoclave door to remove items only when chamber pressure registers 0 lb/in2 (100°C or less). CAUTION: Never stand in the path of escaping steam when opening an autoclave.

USE OF BIOLOGICAL INDICATORS FOR MONITORING AUTOCLAVE PROCESSES

SUPPORT PROTOCOL 1

Biological indicators are used to effectively monitor the efficacy of moist- or dry-heat sterilization processes. The indicators contain standardized preparations and concentrations of resistant endospores of specific strains of bacteria that will survive suboptimal sterilization conditions, and proliferate when subsequently incubated under normal growth conditions. For greatest control of sterilized material, biological controls should be included with every load. Under general laboratory conditions, biological indicators should be used for validation studies in conjunction with thermocouple temperature-sensing probes, and they should be used for frequently scheduled monitoring of the performance of sterilization equipment and procedures. Bioindicator sources listed in this unit are examples only; other products by other manufacturers can work well. Specific manufacturer instructions for product use supercede general instructions described in this protocol. This protocol describes the use of biological indicator ampules for monitoring sterilization. For monitoring dry-heat sterilization, see Support Protocol 2. Cell Culture


Supplement 1

Additional Materials (also see Basic Protocol 1; see Alternate Protocols 1 and 2) Biological indicator ampules: standardized concentration of Bacillus stearothermophilus (ATCC #7953) spores suspended in growth medium containing bromcresol purple as a pH indicator (e.g., Prospore from Raven Biological Laboratories) 55° to 60°C incubator 1. Label the desired number of biological indicator ampules with permanent ink to indicate location and autoclave run number. 2. Place one or more ampules in the most difficult locations to sterilize in the autoclave, including (1) near the front drain of the autoclave, located on the chamber floor at the door of the autoclave, and (2) suspended within the item being sterilized. For liquids, suspend the ampule in the container of liquid by a string tied around the ampule neck and secured around the opening of the container. For dry goods, tuck the ampule within the samples being autoclaved. Manipulating test ampules in loads containing biohazardous waste must be performed by personnel trained for dealing with the potential hazards of the material.

3. Run autoclave cycle at prescribed conditions (see Basic Protocol 1; see Alternate Protocols 1 and 2) and retrieve ampules. CAUTION: After sterilization, handle ampules with care if still hot, as they are under pressure and might burst if strongly jostled. Validation studies should never be performed in routine process cycles; unfortunately, they are too often used interchangeably in small laboratories. If validation and processing are used simultaneously, the sterilized material must be quarantined until the results of the sterilization monitoring tests are confirmed.

4. Place autoclaved test ampules and a labeled, unautoclaved positive control ampule in a vertical position in a 55° to 60°C incubator. 5. Incubate 48 hr. 6. Analyze results by noting the color of the test ampules and positive control ampule. Growth of the positive control confirms lack of sterilization. The positive-control ampule exhibits a color change from purple (prior to sterilization) to yellow (postincubation), with or without turbidity. Growth of the test sample indicates failed sterilization, and is seen as a color change from purple (prior to sterilization) to yellow (postincubation). Positive sterilization is indicated by a purple-colored test ampule (postincubation). An intermediate yellowish color is suspicious and necessitates additional testing of the autoclave parameters. An intermediate grayish color, without turbidity, usually indicates heat destruction of the bromcresol pH indicator, resulting from prolonged autoclaving conditions. BASIC PROTOCOL 2


DRY-HEAT STERILIZATION AND DEPYROGENATION Dry heat is used for components and materials that are resistant to the 140° to 180°C temperatures needed for effective dry sterilization; it is most often used for the sterilization of laboratory glassware and stainless steel instruments. It is also used for sterilization of nonaqueous, heat-stable liquids such as mineral oil. Depyrogenation of heat-tolerant materials is done with ovens capable of operating at the required processing temperatures of 220° to 350°C. As with autoclaving, standard dry-heat sterilization and depyrogenation times refer to the time the material is held at the prescribed temperature and not to the time the oven has been set to run. Dry-heat sterilization using gravity ovens generally requires a longer time than does sterilization with convection ovens, which evenly

1.4.6 Supplement 1


distribute the heated air throughout the chamber with blowers. In all instances, process validation of any dry-heat sterilization protocol is required. The frequent use of bacterial spore strips (see Support Protocol 2) is advantageous for routine monitoring of the efficacy of an established sterilization process. Materials Items to be sterilized Heat-resistant outer containers (borosilicate glass or stainless steel) for small items Aluminum foil Heat-resistant labels or tape Dry-heat indicator tape Laboratory oven (operating temperature of 140° to 180°C for sterilization; 220° to 350°C for depyrogenation) 1. Place small items to be sterilized into heat-resistant outer containers. Use aluminum foil to cover any openings to larger, individually sterilized items, or to cover any openings to large items that do not have their own closures. 2. Code each item or package as to its contents with permanent ink and a heat-resistant label. 3. Affix a piece of dry-heat indicator tape to each item or package as a visual reference (following the oven cycle) that the material has been processed. The color change of indicator tape shows only that the material has been exposed to a prescribed sterilization temperature, but does not indicate the time held at that temperature. It is not an indicator of successful sterilization.

4. Loosely arrange material in the oven. Do not overpack, as this prevents efficient heat penetration to all items. 5. Close and secure oven door. 6. Select operating temperature and time (see Table 1.4.2 for general guidelines). Note that times designated in Table 1.4.2 do not include temperature buildup time, as this is equipment specific. The actual sterilization time begins when the oven chamber reaches the prescribed temperature. Heating times are long for dry-heat sterilization, and can actually be longer than the sterilization time itself. Thus, a load of material might require 2 hr to reach 180°C, while needing only 0.5 hr at that temperature to be effectively sterilized.

7. Run dry-heat sterilization cycle to completion. Any interruption in the cycle (e.g., opening the door to add just one more item to the load) invalidates the run.

Table 1.4.2 Time-Temperature Relationships for Dry-Heat Sterilization

Oven temperature (°C)

Sterilization time (hr)a

180 170 160 150 140

0.5 1.0 2.0 2.5 3.0

aSterilization time indicated is the amount of time for which material should be raised to a given temperature and does not include heating time.

Cell Culture


Supplement 1

8. Turn oven heating element off and allow material to cool to room temperature before removing items from the oven. This can take several hours for large loads in gravity ovens. SUPPORT PROTOCOL 2

USE OF BIOLOGICAL INDICATORS FOR MONITORING DRY-HEAT STERILIZATION Biological indicator strips are used to monitor dry-heat sterilization. For a general discussion of biological indicators, see Support Protocol 1. Additional Materials (also see Basic Protocol 2) Biological indicator strips containing standardized concentrations of Bacills subtilis (ATCC #9372) spores (e.g., Spore-O-Chex; PyMaH, or VWR) Trypticase soy broth (see recipe) 30°C incubator 1. Label the appropriate number of biological indicator strips with location and cycle number or date. 2. Place one or more strips in the most difficult-to-sterilize areas of the load. 3. Run sterilization cycle at prescribed conditions (see Basic Protocol 2) and retrieve strips when cool. 4. Aseptically open the outer wrapping of the indicator, remove the spore strip, and insert it into an appropriately labeled tube of trypticase soy broth (6 to 10 ml per tube). 5. Prepare a positive control by aseptically inserting an unsterilized spore strip into a separate tube of trypticase soy broth. Prepare an uninoculated tube of trypticase soy broth as a negative control. 6. Incubate tubes at 30°C for 4 days. 7. Analyze results by noting turbidity of the broth. Compare test samples with positive and negative control tubes. Resterilize any material in loads with positive test samples. Bacterial growth in the positive control, indicated by cloudy medium, confirms lack of sterilization. Growth should be absent in the negative control; the medium should remain clear, with no precipitate.

BASIC PROTOCOL 3

USE OF DISINFECTANTS: 70% ETHANOL Ethanol is widely used in many laboratories for benchtop or laminar-space disinfection. The antimicrobial activity of the alcoholic solution is very much dependent upon the working concentration of the solvent, proper preparation, storage, and conditions of its use. Ethanol is an effective disinfectant against vegetative bacterial and fungal cells, but is totally ineffective in germicidal activity against bacterial spores. Ethanol is suitable for spraying or swabbing, but is not recommended for large-volume applications. Ethanol is highly flammable, and spills near the flame of a Bunsen burner are always a possible safety hazard. Similarly, an elevated concentration of vaporized ethanol in a liberally disinfected biosafety cabinet could ignite in the presence of a flame or spark. Furthermore, 70% ethanol is not recommended for use in discard pans or for decontamination of biological spills in the catch basins of biological safety cabinets.


1.4.8 Supplement 1


Materials 100% denatured ethanol Ethanol-resistant spray-type storage container 1. Add 700 ml of 100% denatured ethanol to 300 ml deionized or distilled water. Mix well by stirring. Denatured ethanol is absolute ethyl alcohol to which small amounts of chemicals have been added to render it unsuitable for human consumption. This does not interfere with most industrial uses.

2. Store working solution in a tightly closed container to retard evaporation. Choose a spray bottle rather than a squirt bottle to retard the evaporation of solvent that occurs with the larger opening of a squirt bottle neck.

3. For lab benches or laminar-flow cabinets: Liberally spray the alcoholic solution in a crisscross pattern over the work surface, making certain that the entire area is wetted. Let disinfectant remain in contact with the surface for ≥10 min. Wipe away excess solution with absorbent towels 4. For objects (e.g., media bottles, culture flasks): Wet absorbent towels (cheesecloth or paper towels) with the alcoholic solution. Thoroughly swab the object, being careful not to introduce any of the liquid into the threads of screw caps or other container closures. Let disinfectant remain in contact with the object for 3 to 5 min. Wipe away excess solution with absorbent towels USE OF DISINFECTANTS: QUATERNARY AMMONIUM COMPOUNDS The discovery of the antimicrobial activity of quaternary ammonium compounds during the early 20th century was a major advancement in the development of effective germicides. The inherent antimicrobial activity of these compounds was soon shown to be significantly improved by the addition of long-chain alkyl groups to the nitrogen moiety of the quaternary compound. The various quaternary ammonium compounds commercially used as disinfectants today are chemical modifications of this original concept. The mode of action of quaternary ammonium compounds is as cationic surface-active agents, although this chemical property does not fully explain the germicidal activity of the compounds. All have broad-based antimicrobial activity and have proven effectiveness against algae, gram-positive bacteria, some gram-negative bacteria, fungi, and certain viruses, when used at the manufacturer’s recommended concentrations (0.1% to 2.0% active ingredient, or 200 to 700 ppm). They are relatively nontoxic to humans when used according to manufacturer’s instructions and are not chemically destructive to equipment under normal use. They can be autoclaved without formation of toxic vapors and thus are frequently used as disinfectants in discard pans.


The limitations of quaternary ammonium compounds include lack of effectiveness at low concentrations against some commonly encountered gram-negative bacteria (e.g., Pseudomonas sp.). Like many other disinfectants, they are quickly inactivated by the presence of heavy organic burden. Materials Quaternary ammonium compound disinfectant of choice: e.g., Roccal (Sterling Winthrop), Micro-Quat (Ecolab), Zephirol (Bayer) Tightly closed containers Spray bottles 1-gallon jugs

Cell Culture


Supplement 1

1. Dilute concentrated quaternary ammonium compound according to manufacturer’s instructions in deionized or distilled water and stir well to mix. Depending upon use, normal working concentrations of 235 ppm active ingredient are used for sanitization purposes such as floor mopping, whereas concentrations of 470 to 700 ppm active ingredient are used for disinfection of laminar-flow cabinet work surfaces or for use in discard pans.

2. Store working solution in tightly closed containers to retard evaporation. Prepare fresh working solutions frequently (e.g., once a month; follow manufacturer’s instructions).

3. For work surfaces: Liberally spray the working solution in a crisscross pattern over the work surface, making certain that the entire area is wetted. Let disinfectant remain in contact with the surface for >10 min. Wipe away excess solution with absorbent towels. 4. For use in discard pans: Prepare a solution containing 700 ppm quaternary ammonium compound. Fill discard pan about half full with disinfectant solution. Carefully place used pipets into the pans to avoid splashing. Securely cover pan when moving it. Volume of disinfectant must be enough to fully cover the pipets placed into the pan, but not so full that spilling could occur when the pan is filled with pipets or when it is moved. ALTERNATE PROTOCOL 4

USE OF DISINFECTANTS: SODIUM HYPOCHLORITE Chlorine, in various forms, has a long history of use as a powerful disinfectant, yet the exact mode of germicidal action is unclear. Hypochlorites are the most widely used chlorine compounds for disinfection. Commercial liquid bleach products (e.g., Clorox) are solutions containing 5.25% (w/v) sodium hypochlorite. Sodium hypochlorite is effective against vegetative microbial cells, most spores, and many viruses. It has some residual effect after the treated surface dries. It can be used in sanitization procedures for laboratory floors and in laboratory coat washing. It is strongly germicidal and can be used to decontaminate small- to mid-volume spills of biological material. Despite their germicidal effectiveness, chlorine solutions are limited in their use as laboratory disinfectants because of their corrosiveness to metals and their human toxicity. They should not be routinely used in discard pans or in any solutions that are autoclaved, as the chlorine fumes liberated are significant skin and respiratory irritants. Frequent autoclaving of chlorine solutions will corrode the chamber interior of the sterilizer. One exception to this autoclave ban is the need to sterilize any biological spill material in which bleach was used as a disinfectant during the cleanup process. A solution of 10% (v/v) household bleach is a strongly germicidal, containing ∼0.52% (w/v) sodium hypochlorite. Excess hypochlorite is needed in mopping up spills of biological agents, to supply additional chlorine to replace that consumed by the large amount of organic matter associated with the spill. Sodium hypochlorite solutions can be inactivated by organic matter (which consumes the available free chlorine that constitutes microbiocidal activity), by exposure to UV light, and by inorganic chemical reducing agents (such as ferrous or manganese cations and hydrogen sulfide). Hypochlorite solutions should be stored away from heat to avoid deterioration.


Materials Household liquid bleach (e.g., Clorox, Dazzle) 5% (w/v) sodium thiosulfate solution

1.4.10 Supplement 1


1. Add 100 ml household bleach to 900 ml water to give a 10% (v/v) solution. Mix well. Store in the dark away from heat. 2. For cleanup of open spills: Soak paper towels with bleach and gently cover the spill, being careful not to enlarge the area of the spill. Let bleach stay in contact with the spill for ≥20 min. During this time, decontaminate any nearby areas that may have been subject to spatters from the original spill by swabbing with additional 10% bleach. Collect used paper towels in a suitable container and place in an autoclave bag. 3. For autoclaving bleach-containing waste: Add ∼1 vol of 5% sodium thiosulfate solution to the bleach solution to help neutralize the chlorine. Seal the bag and autoclave. Prominently label the autoclave room door to warn laboratory personnel of the potential for irritating vapors. When the autoclave cycle is finished, crack the autoclave door slightly to allow remaining chlorine fumes to dissipate before removing the bags. 4. For sanitizing solution: Add 14.8 ml household bleach to 3.78 liter water (0.4%). Mix well. Solutions containing 0.4% (v/v) household bleach (200 ppm available chlorine) are suitable for soaking lab coats without being so strong as to harm the fabric. The solution is acceptable for routine floor maintenance.

5. For disinfection solution: Add 44.4 ml household bleach to 3.78 liter water (1.2%). Mix well. Liberally apply to surface with clean absorbent towels and let stand for >10 min. Dry surface with a separate clean towel. This concentration of bleach (600 ppm available chlorine) is suitable for disinfecting biosafety cabinets during scheduled weekly maintenance. More frequent use on stainless steel may not be recommended because of the corrosiveness of the bleach.

FILTER STERILIZATION OF SOLUTIONS All solutions that come in contact with cell cultures must be sterile in order to prevent microbial contamination. This includes non-nutritive preparations such as distilled/deionized water and reagents (e.g., dimethyl sulfoxide used as a cryoprotectant). Although heat-stable solutions can be sterilized by autoclaving, many solutions used in cell culture contain one or more heat-labile components (e.g., antibiotics), or are chemically formulated with ingredients that will form deleterious precipitates if subjected to steam sterilization temperatures (e.g., phosphate-buffered salines). Membrane filtration is the most common cold sterilization method for these types of solutions. Filter membranes with 0.2-µm pore size are used for general sterilization purposes; however, some environmentally stressed bacteria (e.g., Pseudomonas sp.) as well as mycoplasma can pass through filters of this porosity. To provide a greater degree of assurance for complete removal of these common tissue culture contaminants, cell culture media and sera should be sterilized using 0.1-µm filter membranes. Filter manufacturers offer many different types of membranes. With respect to cell culture applications, membranes fabricated from cellulose acetate or cellulose nitrate are used for general purpose filtration of aqueous solutions such as media and buffers, but may need prewashing with hot distilled water to remove extractable substances that may be cytotoxic. Nylon membranes are very low in extractable substances such as surfactants or wetting agents; polyethersulfone membranes are low in extractables and have very low protein binding. Cell Culture


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The availability of presterilized, ready-to-use, disposable filter systems has eliminated much of the labor and risk of failure inherent with earlier filtration methods. Filter units come in a wide variety of sizes to handle small (≤10-ml) to large (≥20-liter) volumes. Disposable systems are available for either vacuum or positive-pressure filtration. Many manufacturers have filter systems designed specifically for cell culture applications: the sterilization membranes, housings, and receiving vessels are certified noncytotoxic and nonpyrogenic. This section outlines selection of filters and filter-sterilization procedures for various types of liquids encountered in cell culture laboratories. The most common small-volume filtration technique is positive pressure using a syringe to force the liquid through the filter membrane (see Basic Protocol 5). Volumes ranging from 50 ml to 1 liter are most efficiently processed with a vacuum (see Basic Protocol 4). Larger volumes should be filter sterilized with positive pressure (see Alternate Protocols 5 and 6). The primary use of membrane filtration is in the preparation of tissue culture media; this topic is treated in depth in UNIT 1.2. This unit focuses on problematic filtration needs that often appear in cell culture applications, such as the need to filter sterilize a hazy solution like the serum/yeast extract additives used in mycoplasma media, or the chemically aggressive reagent dimethyl sulfoxide (DMSO). The methods outlined in this section are equally adaptable for the preparation of tissue culture media or stock solutions of additives such as glutamine or puruvate. For media preparation, use noncytotoxic cellulose acetate/nitrate membranes, or similar membranes specific to the application, and food-grade silicon tubing. For further details on the use of vacuum versus positive-pressure filtration, see Background Information. BASIC PROTOCOL 4

Vacuum Filtration Solutions that are initially clean preparations, in that they are free of particulate debris and are not proteinaceous, can be directly filter sterilized with no difficulty. Solutions with high particle load require centrifugation and/or nonsterile prefiltration through depth filters (see Background Information) and larger-porosity membranes prior to sterile 0.2-µm filtration. This protocol uses 200-ml to 1-liter disposable systems designed for the final vacuum filtration of media, sera, and other aqueous solutions. The protocol also describes nonsterile prefiltration for particulate removal from 200-ml to 2-liter volumes of filtrate. Materials Solution to be filtered 47-mm funnel/support assembly (optional; e.g., Kontes, Millipore) attached to a 1to 2-liter vacuum filtration flask (Fig. 1.4.1) 47-mm glass fiber depth filters (optional; Gelman, Millipore) 47-mm membrane filters (optional; 0.45-µm and 0.2-µm pore sizes) Disposable, sterile filter unit (e.g., Corning, Nalgene) including: Filter funnel, housing an integrally sealed 0.2-µm filter membrane Funnel dust cover Removable receiver bottle and cap Barbed tubing adapter Nonsterile depth prefilters (included by most manufacturers) Vacuum source NOTE: Perform all procedures using aseptic technique (UNIT 1.3).


1. If the solution to be sterilized is a hazy suspension or has a noticeable precipitate, centrifuge 30 min at 10,000 × g to clarify. Alternatively, use a funnel/filter assembly

1.4.12 Supplement 1


to process the solution through a series of nonsterile prefilters: depth filter, followed by 0.45-µm membrane, followed by 0.2-µm membrane. Depending upon the particulate load, the filter membranes may have to be replaced if they clog before all the solution is processed. If a water aspirator is used as the vacuum source, include an in-line catch flask or hydrophobic filter to prevent any water from accidentally being drawn into the vacuum port and into the filtration flask.

2. Remove a disposable, sterile filter unit, barbed tubing adapter, and individual wrapped sterile receiver cap from the plastic bag. 3. Check to make certain that the filter funnel is firmly attached to the receiver. Hand tighten if necessary. 4. Attach barbed tubing adapter to the side vacuum port on the neck of the filter funnel. Attach the vacuum tubing to the adapter. If a water aspirator is used, include an in-line catch flask (see step 1).

5. Set the filter upright and provide support to avoid tipping the unit when it is top-heavy with liquid in the funnel. 6. Remove the funnel dust cover and slowly add solution (centrifuged or prefiltered if necessary) to the funnel. Slowly apply a slight vacuum—5 pounds per square inch gravity (psig)—to prevent excessive foaming of proteinaceous solutions. 7. When filtration is complete, turn the vacuum source off. Carefully disconnect the filter unit from the vacuum tubing. If the tubing is pulled off while the vessel is still under full vacuum, the room air rushes into the receiving vessel at a much higher velocity.

8. Using aseptic technique, carefully remove the filter funnel from the receiver bottle and seal the bottle with the sterile screw-cap closure provided with the filter unit. If the total volume of solution to be filtered exceeds the capacity of the receiver bottle supplied with the filter unit, the initial volume of sterile filtrate can be aseptically transferred to a secondary sterile storage vessel, and the filter funnel can be reattached to the original receiver to process a second volume of solution.

Figure 1.4.1 Funnel/support assembly for vacuum prefiltration.

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BASIC PROTOCOL 5

Small-Volume Positive-Pressure Filtration of Nonaqueous Solutions Dimethyl sulfoxide (DMSO) is used as the cryoprotectant for liquid-nitrogen preservation of cell cultures. The reagent is not stable to autoclaving conditions and must be filter sterilized. DMSO is an aggressive solvent that dissolves general-use filter membranes (such as cellulose acetate or cellulose nitrate) as well as the polystyrene filter units themselves. This protocol describes small-volume positive-pressure filtration using DMSO-resistant syringe-type filter units. For large volumes, see Alternate Protocols 5 and 6. Materials Dimethyl sulfoxide (DMSO) Glass 25-ml syringe with Luer-lok tip Sterile syringe filter: 25-mm-diameter nylon membrane, 0.2-µm pore size, polypropylene housing (Nalgene or equivalent) Laminar-flow cabinet Sterile amber glass storage vessels with polytetrafluoroethylene (Teflon, PTFE)–lined screw-cap closure NOTE: Perform all procedures using aseptic technique (UNIT 1.3). 1. Load a glass 25-ml syringe with deionized or distilled water. 2. Aseptically remove a sterile syringe filter from its blister-package wrapper, being careful not to touch the outlet nipple. 3. Attach the inlet end of the filter to the syringe and finger tighten the Luer-lok connection. 4. Apply a firm, but not forceful, pressure to slowly discharge the water through the filter into a waste container. This initial step is necessary to wet the filter to permit flow of the DMSO through the nylon membrane.

5. Carefully remove the filter from the syringe, and rest the filter on the laminar-flow cabinet work surface, being careful to keep the outlet nipple facing up (i.e., not touching any surface). 6. Load the syringe with ∼25 ml DMSO and replace the filter as in step 3. 7. Apply a firm, but not forceful, pressure to slowly discharge this first volume of DMSO into a waste container. This step clears any water remaining in the syringe and filter.

8. Reload the syringe with ∼25 ml DMSO and discharge the filtrate into a suitable sterile amber glass storage vessel. Cap immediately with a PTFE-lined screw-cap closures. Store up to 6 to 9 months at room temperature. ALTERNATE PROTOCOL 5


Large-Volume Positive-Pressure Filtration of Nonaqueous Solutions This protocol uses a peristaltic pump to provide positive pressure for large-volume filtration. The setup is shown in Figure 1.4.2. Materials Dimethyl sulfoxide (DMSO) Sterile filter capsule: 400-cm2-surface-area nylon membrane, 0.2-µm pore size, polypropylene housing (Whatman Polycap 36AS or equivalent) Glass 25-ml syringe with Luer-lok tip

1.4.14 Supplement 1


Worm drive clamps PTFE tubing: polytetrafluoroethylene (PTFE or Teflon) with 0.25-in. (6.4-mm) i.d., 0.06-in. (1.6-mm) wall thickness, 0.38-in. (9.5-mm) o.d. (Norton or equivalent) Peristaltic pump assembly capable of providing an operating pressure of 15 to 20 lb/in.2 Sterile amber glass storage vessels with PTFE-lined screw caps NOTE: Perform all procedures using aseptic technique (UNIT 1.3). 1. Carefully remove a sterile capsule filter from its plastic bag. 2. Remove the nipple cover from the inlet barb. 3. Hold the filter over a waste container, and use a glass 25-ml syringe to carefully add water to the filter housing through the inlet barb. Fill the syringe as necessary and continue to flush the housing until water begins to drip from the sterile outlet side. This step wets the nylon membrane to allow the DMSO to pass through the filter.

4. Attach a piece of PTFE tubing to the inlet barb from the nonsterile DMSO reservoir. Using worm drive clamps, secure the tubing at all connections when working with positive pressure to prevent sudden blowing off of tubing in case of accidental overpressurization. 5. Secure the capsule filter to an upright support at a height with sufficient clearance to accept any receiving vessels. 6. Connect the tubing as shown in Figure 1.4.2, to the peristaltic pump head according to the pump manufacturer’s instructions. Apply power to the pump and begin pumping. 7. Discharge the first 200 ml of DMSO filtrate to a waste container. This step clears any water remaining in the filter and tubing.

8. Collect sterile DMSO in suitable sterile amber glass storage vessels and cap immediately with PTFE-lined screw-cap closures. Store up to 6 to 9 months at room temperature.

disposable capsule filter unit

peristaltic pump

Figure 1.4.2 Positive-pressure filtration assembly for use with a peristaltic pump.

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Large-Volume Positive-Pressure Filtration Using Pressurized Nitrogen This alternative method for large-volume filtration uses pressurized nitrogen to force the solvent through the filter membrane. It offers an advantage for filtration of DMSO in that it saturates the solvent with an oxygen-free gas phase that helps retard oxidation of the material during storage. The setup is shown in Figure 1.4.3. Additional Materials (also see Alternate Protocol 5) 5-liter pressure vessel (or size to fit application; Gelman or Millipore) Pressurized nitrogen tank Additional tubing to fit pressure vessel and nitrogen tank 1. Prepare a capsule filter (see Alternate Protocol 5, steps 1 to 3). 2. Attach a piece of tubing to the filter inlet barb from the outlet barb of a 5-liter pressure vessel. Using worm drive clamps, secure the tubing at all connections when working with positive pressure to prevent sudden blowing off of tubing in case of accidental overpressurization. 3. Fill pressure vessel with DMSO. Close and secure pressure vessel lid. Open pressure relief valve on vessel. 4. Attach another piece of tubing from a pressurized nitrogen tank to the inlet barb on the pressure vessel and secure with a worm drive clamp. 5. Slowly open nitrogen feed valve and wait until gas can be heard escaping from the pressure relief valve. 6. Close pressure relief valve. Pressure will begin to rise in the vessel as indicated either on the pressure gauge included with the vessel, or by the gauges on the nitrogen tank regulator. To prevent bursting of the filter membrane, keep the operating pressure below the maximum rated pressure specified by the filter’s manufacturer.

7. Collect sterile DMSO filtrate (see Alternate Protocol 5, steps 7 and 8).

disposable capsule filter unit

pressurized nitrogen tank

pressure vessel Sterilization and Filtration

Figure 1.4.3 Positive-pressure filtration assembly for use with pressurized nitrogen.

1.4.16 Supplement 1


REAGENTS AND SOLUTIONS Use deionized or distilled water in all recipes and protocol steps. For common stock solutions, see APPENDIX 2A; for suppliers, see SUPPLIERS APPENDIX.

Trypticase soy broth Suspend 30.0 g trypticase soy broth powder (BBL) in 1.0 liter water and stir to dissolve. Dispense 6 to 10 ml per 16 × 125–mm tube and cap loosely. Sterilize by autoclaving at 121°C for 15 min. Store prepared tubes at 4° to 8°C for up to 6 to 9 months. COMMENTARY Background Information Autoclaving Steam (autoclave) and dry heat are destructive terminal sterilization processes in which the effectiveness of the method is characterized by the rate of the microbial killing. As the order of death in a terminal sterilization process is a logarithmic function, mathematical calculations will never result in a zero survival rate. Thus, theoretically, complete sterilization is impossible. Therefore, rather than expecting an absolute, the effectiveness of terminal sterilization processes is mathematically expressed in terms of the decimal reduction time (D value), which is the time required at a given temperature to destroy 90% of survivors. A corollary term used in discussions of terminal sterilization is the Z value, which is the temperature at which a survival curve decreases by one log. Factors influencing these values include the concentration and type of microbial contaminants initially in the material to be sterilized, the physical nature of the material undergoing sterilization, and the performance characteristics of the sterilization equipment. Disinfectants The use of disinfectants in the cell culture laboratory is directed both to issues of personal safety as well as quality control. In most instances the distinction between the two considerations blurs; however, this unit discusses the use of disinfectants primarily as a means for prevention of microbial contamination from the standpoint of quality control (i.e., maintenance of noncontaminating conditions for cell culture manipulations). Human safety considerations are limited to brief notations regarding potential effects of misuse of specific disinfectants (e.g., chemical incompatibilities, lack of effectiveness against certain biological agents) rather than a reiteration of the need for disin-

fectants as a means of personal defense against etiologic agents. The list of common disinfectants is lengthy: alcohols (ethanol or isopropanol), chlorine compounds (bleach), hydrogen peroxide, phenolics, and iodophores (povidone-iodine), to name a few. Yet from this broad list, there is no universal disinfectant solution that can work effectively for all laboratory situations. Thus, the concern for any laboratory is deciding upon its different needs for disinfection (e.g., spill cleanup, work surface disinfection, discard pan disinfection, routine floor cleaning, lab coat laundering), and then selecting the appropriate disinfectant and concentration for each purpose. It is beyond the scope of this unit to provide a lengthy treatise on all the commercially available liquid germicides. Focus is directed, instead, to the three most commonly encountered disinfectants utilized in tissue culture: 70% ethanol, quaternary ammonium compounds, and sodium hypochlorite (bleach). Filtration Filtration as a method of sterilization has been in use for over one hundred years. Early filters were designed to trap contaminants within the depths of a thick, tortuous maze of filter material. Scintered glass filters and the asbestos (later cellulose) Seitz filters commonly used in the 1950s and 1960s worked on the entrapment principle. These depth filters have significant loading capacity and can retain much particulate matter before clogging. The limitation of depth filters is the structural nature of the filter matrix. At best, the loose matrix of a depth filter permits only a nominal designation of particle size retention. Because the pores of a depth filter are of random size and shape, there is a real probability that small-sized contaminants will successfully channel through the filter through interconnecting pores. In addition, the effects of moisture and pressure associated with autoclave sterilization and/or use of

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depth filters tends to result in a shifting of the filter material (media migration) that compromises the integrity of the fibrous filter matrix. By the mid 1970s membrane filters had essentially replaced depth filters for sterile applications. Membranes are classified as screen filters: they are thin and the pores are uniformly sized and spaced across the sheet. This structural consistency allows predictable retention characteristics. Thus, membrane filters can be rated according to the minimal diameter of the smallest particle they will retain (e.g., 1.2-µm, 0.45-µm, 0.2-µm). Membrane filters have limited loading capacity (i.e., they quickly clog). For this reason, they are most effectively used in tandem with depth filters and membrane filters of larger porosity to conserve the filter surface area on the final sterilization membrane. The type of filtration system, as well as the type of membrane used in the system, depends upon the nature of the filterable material and the volume of material being filtered. A wide variety of configurations is available in both presterilized disposable units containing integrated filter membranes, as well as membrane discs that can be used in conjunction with reusable glass or stainless steel filter housings. In the past, positive pressure was the preferred filtration method for cell culture media because it reduced the foaming and concomitant protein denaturation associated with vacuum filtration systems. Unfortunately, positive-pressure filtration systems are efficient only for small (syringe filtration) or large (pressure vessel) volumes. With recent technological advances in new membrane matrices and improved membrane supports for filter housings that eliminate excessive foaming, vacuum filtration with presterilized disposable units is now the easiest and most effective sterilization method for intermediate volumes (0.1 to 2 liters) of tissue culture media. Filter membranes are made from a number of different materials. The most common filters featured in presterilized, disposable filtration units are those fabricated from esters of cellulose acetate or cellulose nitrate. These generalpurpose hydrophilic membranes are suitable for aqueous solutions such as tissue culture basal media and supplements. These membranes can, however, bind proteins and may be of concern for certain critical applications. With the huge cell culture market as a direct target, filter manufacturers have, in recent years, begun to offer presterilized filtration units customized for cell culture, featuring noncyto-

toxic, low-protein-binding membranes (nylon or polyethersulfone). Not every solution utilized in cell culture applications is truly aqueous or hydrophilic. Nonpolar liquids, such as DMSO, or chemically aggressive solutions, such as concentrated acids or bases, demand special chemically resistant membrane filters. Quite often, filters resistant to nonpolar solvents will need pretreatment with an appropriate wetting agent to quickly render the membrane filterable to a particular nonpolar liquid. For example, PTFE (Teflon) membrane filters require prewashing with methanol prior to use with DMSO. This poses no problem providing all traces of the cytotoxic alcohol are removed by washing prior to collection of any sterile, unadulterated DMSO filtrate. Where residual toxic wetting agents are a concern, nylon filters may be a more appropriate consideration. Nylon membranes, when wetted with water, readily accept DMSO. With wide recognition of the detrimental effects of insidious mycoplasma contamination and the concern about absolute removal of these contaminants from a major point of entry (i.e., serum) into a cell culture system, much emphasis is now placed on 0.1-µm filtration of cell culture media and sera. Unfortunately, this pore size is not widely available in disposable filtration units. Gelman Sciences does, however, offer presterilized filter units with 0.1-µm polyethersulfone (Supor) membranes for small- to large-volume cell culture media sterilization. Expect slower flow rates with 0.1-µm filtration. When lacking presterilized filtration units, a laboratory can turn to individual membrane discs of specified porosity and membrane type, available from major filter manufacturers. These can be sterilized by autoclaving as part of an integrated unit in a small-volume filter assembly (e.g., Fig. 1.4.1), or for large-volume needs in a 142- or 293-mm filter holder (Gelman or Millipore). If this route for sterile filtration is chosen, follow the manufacturer’s instructions precisely with respect to membrane sterilization times, and use a slow exhaust cycle to avoid the membrane cracking that can occur with rapid pressure changes. This unit outlines procedures that are suitable for any number of variations. Large volumes of tissue culture media can be processed with the peristaltic pump or pressurized nitrogen procedures providing a hydrophilic filter is used. Similarly, a hydrophilic syringe filter can be used to sterilize small volumes of media. Sterilization of DMSO is, however, limited to

1.4.18 Supplement 1


those filter membranes and filter unit housings fabricated of materials resistant to the reagent.

totoxic boiler amines and/or chemical softening agents.

Critical Parameters

Dry-heat sterilization Hot dry air is an inefficient means of sterilization and should be reserved for those materials that cannot effectively be exposed to saturated steam in an autoclaving process. Sterilization by dry heat can be accomplished with temperatures as low as 140°C, but 170° to 180°C are more routine operating temperatures because of the difficulty in controlling the rate of heat penetration into the load. The time-temperature relationships for dry-heat sterilization noted in Table 1.4.2 indicate a requirement for longer times and higher temperatures than for autoclaving, because dry heat is less efficient than moist heat. Lag time for conduction of heat into the materials to be sterilized can be significant, as can cool-down periods for large, dense objects. Dry-heat sterilization is applicable only for materials resistant to 140° to 180°C, and thus is unsuitable for paper, many plastics, or rubber. As noted for autoclaving, a dry-heat sterilization procedure should be validated with thermocouples and biological indicators.

Autoclaving Successful autoclave sterilization is dependent upon the contents of a load coming into full contact with saturated steam at 121°C for 15 min. Many operational factors tend to work against these criteria. With respect to the sterilizer itself, if an autoclave is improperly maintained, problems can occur with inadequate removal of air from the chamber or with excessive moisture buildup. Either condition compromises sterilization parameters. Most autoclaves have cool spots that can move about in the chamber like a current depending upon load pattern and configuration. The nature of the load content also influences the time necessary to reach the time-temperature relationship. Small volumes will heat to sterilization temperatures faster than large volumes. Agar solutions that have solidified prior to sterilization will take longer than those that are loaded into the autoclave while still molten. Dry materials take longer to sterilize than those that are moist. For any laboratory investing in as expensive a pursuit as tissue culture, sterilization equipment and procedures should be validated to assure that conditions for sterilization are met. Ideally a laboratory should invest in a thermocouple to monitor temperatures within areas of autoclave loads, in order to determine exactly how long it takes the material of concern to reach sterilization temperature. In many cases, the lag between the time an autoclave temperature gauge indicates 121°C in the autoclave chamber and the time the contents of a large flask or discard pan within the chamber reaches the same temperature is sobering. Where thermocouples are not available, the use of biological indicators can yield useful information about autoclave procedures and machine performance, although the time needed for incubation and interpretation of results is a drawback. A final, but most important, consideration for autoclaving is the source of steam generation. Autoclaves used in the preparation of media or for sterilization of materials that come into contact with cell cultures must be supplied with clean steam (i.e., steam generated from purified water). Steam generated directly from general building physical plant sources (e.g., the building heating system) is frequently produced from water treated with potentially cy-

Disinfectants The term disinfection refers to the treatment of surfaces with chemical solutions to reduce microbial presence. Exposure to a disinfectant may result only in bacteriostatic or fungistatic rather than microbiocidal conditions. Disinfection does not imply sterilization, and it should never be used as an alternative to appropriate sterilization methods (e.g., autoclaving, dry heat, ethylene oxide, incineration). Similarly, disinfectants are not detergents and they should not be used as the sole method for cleaning solid surfaces. Indeed, many disinfectants are quickly inactivated when burdened with organic matter. Thus, routine disinfection of work surfaces in laminar-flow cabinets first requires that the surface be washed with a good detergent to remove dried media or other dirt before application of the germicide. Ethanol. A solution of 70% ethanol is not germicidal against bacterial spores. While freshly prepared solutions are normally free of spores, the working solutions can contain bacterial spores from cross-contamination and poor aseptic technique. Once the spores are separated from the physical presence of the disinfectant (i.e., when the ethanol volatilizes from a surface, leaving the dried spores behind), they can germinate in suitable growth conditions. Thus, fresh working solutions of

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70% ethanol should be prepared frequently (e.g., weekly). The efficacy of ethanol as a disinfectant is highly concentration dependent. The mode of action of alcohol as a disinfectant is protein denaturation. Thus, 95% ethanol is a poor disinfectant because there is not enough water in the preparation to permit effective denaturation of contaminant proteins. As ethanolic concentration drops below 70%, there simply isn’t enough of the solvent present to adequately react with large concentrations of proteinaceous matter. Because of the dilution effect, 70% ethanol is not effective in disinfecting large spills of culture material. Quaternary ammonium compounds. Quaternary ammonium compounds are not effective against spores and, in this aspect, should be used with the same cautions noted for ethanol. Quaternary ammonium compounds are quickly inactivated by organic matter. They must not be used for disinfection of large spills of culture material, nor should they be used for kill pans (see below). Bleach. Chlorine compounds, most frequently sodium hypochlorite, are strongly germicidal. Yet their potential for human toxicity and strong corrosiveness limits their use. Biological spills. Strong disinfectant solutions are the first line of defense in decontaminating small to moderate spills of biological agents in both open areas and biosafety cabinets. The choice and concentration of disinfectant is particularly critical in this application. Use of 70% ethanol would be a poor choice as its germicidal activity is highly concentration dependent and dilution effects associated with spill cleanup would diminish its effectiveness (see above). Quaternary ammonium compounds are germicidal against a broad range of microbes, but they are quickly chemically overwhelmed and inactivated by organic matter (e.g., the culture material or medium in a spill). Chlorine compounds are strongly germicidal. Kill pans. A final note of caution regarding disinfectants is their use in discard pans or pipet pans. Too many laboratories consider these containers kill pans and use them as a convenient way to dispose of excess liquid cultures or other contaminated solutions. Discard pans should never be used in such a manner. Most general disinfectants are inactivated by excess organic matter and/or exhibit diminished germicidal effects with dilution. Kill pans should contain 10% (v/v) household bleach (0.525% sodium hypochlorite).

Filtration When using disposable filtration units, replace the filter funnel immediately with a permanent receiving vessel closure once filtration is complete. If the funnel is left on the receiver, the filter membrane will crack as soon as it dries, thereby compromising the sterility of the filtrate. Be prepared and willing to prefilter any hazy or precipitated suspension. The additional steps will much repay the effort when balanced against the significant monetary expense, time, and frustration spent dealing with prematurely clogged sterilization filters. If one repeatedly filter sterilizes the same type of suspension, a prefiltration scheme can be tailored according to the nature of the particulates. For example, if particulates are retained only by 0.2-µm porosity filters, then omit prefiltration with depth filters and 0.45-µm filters. Process large volumes of slow-filtering liquids (such as serum or other proteinaceous substances) in a cold room, if possible, to retard proliferation of microbial growth during the sometimes time-consuming, nonsterile prefiltration steps.

Anticipated Results If sterilization and disinfection procedures are effective and proper aseptic technique is used, it should be possible to initiate and maintain cell cultures without any incidence of contamination.

Time Considerations Preparation of materials for autoclaving and the autoclaving itself should take 1 to 2 hr; cooling solutions and equipment may require several hours. Dry-heat sterilization should take less than half a day plus cooling time. Monitoring the efficacy of sterilization requires several days to allow time for contaminants to grow. Disinfection of space and equipment requires a variable amount of time. The time required for filter sterilization can be a few minutes to hours depending on the solution.

Key References Barkley, W.E. and Richardson, J.H. 1994. Laboratory safety. In Methods for General and Molecular Bacteriology (P. Gerhardt, R.G.E. Murray, W.A. Wood, and N.R. Krieg, eds.) pp. 715-734. American Society for Microbiology, Washington, D.C. A well-written chapter covering all aspects of safety in any laboratory dealing with biological agents.

1.4.20 Supplement 1


Block, S.S. 1983. Disinfection, Sterilization, and Preservation, 3rd ed. Lea & Febiger, Philadelphia. A substantial text covering all aspects of the discipline. Contributors provide a great deal of information to the individual chapters. Brock, T.D. 1983. Membrane Filtration: A User’s Guide and Reference Manual. Science Tech, Inc., Madison, Wis. Includes detailed information on the principles of membrane filtration, selection of filtration systems, and use of membranes for specific applications. Lacks information on the newer membrane types developed since its publication date, but still a worthwhile reference. Perkins, J.J. 1976. Principles and Methods of Sterilization in Health Sciences. Thomas, Springfield, Ill. A classic reference text that covers all aspects of dry-heat and steam sterilization. U.S. Pharmacopeial Convention. 1995. The U.S. Pharmacopeia XXIII/The National Formulary XVIII. U.S. Pharmacopeial Convention, Rockville, MD.

Vesley, D. and Lauer, J. 1986. Decontamination, sterilization, disinfection, and antisepsis in the microbiology laboratory. In Laboratory Safety: Principles and Practices (B.M. Miller, D.H.M. Gröschel, J.H. Richardson, D. Vesley, J.R. Songer, R.D. Housewright, and W.E. Barkley, eds.) pp. 182-198. American Society for Microbiology, Washington, D.C. Presents substantial information on all aspects of decontamination concerns for the laboratory, including choice of disinfectants, spill containment and cleanup, and routine cleaning. Millipore. 1993. Millipore Direct. Millipore, Bedford, MA. This filter manufacturer’s catalog and reference guide provides a wealth of background, practical, and technical information to assist a user in appropriate choices for filters and filtration equipment.

Contributed by Rosalie J. Coté Becton Dickinson Microbiology Systems Sparks, Maryland

The U.S. official standard for sterilization criteria and sterility testing. Describes methods for using biological indicators.

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Assessing and Controlling Microbial Contamination in Cell Cultures

UNIT 1.5

This unit describes procedures for the detection of bacterial, fungal, and mycoplasmal contaminants in cell cultures. Bacterial and fungal contaminants are detected by direct culture under conditions that specifically favor bacteria, mycelia, and yeast (see Basic Protocol 1). The direct method for detecting mycoplasma contamination similarly involves screening with microbiological media designed to encourage proliferation of mycoplasma (see Basic Protocol 2). The two indirect methods presented are (1) a slight modification of Barile’s adaptation on the use of the Hoechst stain to detect mycoplasma by DNA fluorescence (see Alternate Protocol 1), and (2) the use of polymerase chain reaction in conjunction with a commercially available mycoplasma detection kit (see Alternate Protocol 2 and Support Protocol 1). In addition, a procedure is described for controlling microbial contamination through the use of antibiotics (see Basic Protocol 3). Testing for microbial contamination should be integrated into a cell culture program as part of routine quality control. Microbial and mycoplasma testing should be performed upon arrival of all incoming cell lines and on lot samples of ampules prepared for master or working cell banks and seed stocks. Testing for microbial contamination should also be done whenever contamination is suspected (e.g., unusually slow growth rates for a particular cell line, aberrant appearance of cells). Indirect mycoplasma screening methods should also be done on new lots of serum used in media preparation when first received by the laboratory. Testing for microbial contamination should be performed after the cells have been cultured in the absence of antibiotics for several weeks. TESTING FOR BACTERIAL AND FUNGAL CONTAMINANTS The media and methods described in this protocol are suitable for detection of most bacteria and fungi that would be expected to survive as contaminants in cell lines. Brain heart infusion and trypticase soy agar with sheep blood are used for the cultivation of nutritionally fastidious bacteria of clinical origin that may be present in primary tissue cultures or in material contaminated by bacterial flora from human skin and poor aseptic technique. Fluid thioglycollate supports the growth of bacteria that require reduced oxygen tension; these microaerophilic or slightly anaerobic contaminants are frequently spore formers that originate from inadequately autoclaved or heat-sterilized materials. Soybean/casein digest broth is a general-purpose medium that supports the growth of a wide range of bacteria of human or environmental origin. HEPES/trypticase/yeast extract (HTYE) broth is also a general bacterial growth medium, but has the advantage of supporting growth of nutritionally or physiologically stressed bacteria not easily culturable with other media. These general types of bacteria are primarily environmental in origin and can be found in distilled water carboys, in fouled deionization systems, or as air-borne contaminants. Emmons’ modification of Sabouraud’s agar and YM agar are used for detection of filamentous fungi (molds) and yeasts, respectively. Molds are frequently environmental contaminants that can thrive under a wide variety of conditions. They often take residence in air-handling ducts to generate a constant microscopic rain of spores into a laboratory. Similarly, molds can often be found as films colonizing the dispensing tubing from distilled water reservoirs. The types of yeasts normally found as cell culture contaminants are human in origin. Many of the media listed below may be purchased in their final configurations as sterile plates or test tubes from microbiological media manufacturers such as BBL or Difco, as well as other suppliers. Be that as it may, the cell culture user should carefully evaluate Contributed by Rosalie Coté Current Protocols in Cell Biology (1999) 1.5.1-1.5.18 Copyright © 1999 by John Wiley & Sons, Inc.

BASIC PROTOCOL 1

Cell Culture

1.5.1 Supplement 1

the configurations, performance, and cost of any prepared quality control medium before commencing full-scale use in a cell culture laboratory. Materials Medium for bacterial detection: e.g., brain heart infusion (BBL, Difco), fluid thioglycollate medium (BBL, Difco), HTYE broth (see recipe), soybean/casein digest broth USP (e.g., trypticase soy broth, BBL; tryptic soy broth, Difco), or trypticase soy agar (BBL) Medium for mycelial and yeast fungal detection: e.g., Sabouraud’s dextrose agar (Emmon’s modified; BBL, Difco), or YM agar (Difco) Sterile, defibrinated sheep blood (e.g., Colorado Serum, Waltz Farm) Cell culture test samples Antibiotic-free culture medium (optional) Conductivity meter (Corning model 162 or equivalent), if not integrated with the laboratory water purification system 50°C water bath 16 × 125–mm borosilicate screw-cap test tubes with rubber-lined caps 100 × 15–mm sterile plastic disposable petri dishes Semiautomated repeat-volume filling unit to accurately dispense 5- to 24-ml aliquots (optional) Incubators: 26°C, 35° to 37°C, and 37°C with 5% (v/v) CO2 NOTE: To avoid inadvertent contamination of clean cell lines, bacterial and fungal testing should be segregated to a laboratory not used for general cell culture work. Prepare media For liquid (broth) media: 1a. Reconstitute brain heart infusion, HTYE broth, soybean/casein broth, and fluid thioglycollate medium per manufacturer’s instructions, or per specific recipe instructions, in 10-megaohm (or higher) distilled or deionized water. Heat to ∼50°C with frequent stirring to dissolve components. Heat to boiling with frequent stirring to dissolve any medium containing even small amounts of agar (e.g., fluid thioglycollate). 2a. Dispense medium into 16 × 125–mm borosilicate screw-cap test tubes at 10 ml/tube for fluid thioglycollate medium, and a 5 ml/tube for all other media. 3a. Cap tubes loosely, threading caps securely enough to prevent them from blowing off during autoclaving, but loosely enough to permit pressure exchange within the tube head space during the sterilization process. Because of the large (10-ml) volume required with fluid thioglycollate medium, one should anticipate significant tube blow-outs upon autoclaving, and should prepare ∼25% more tubes than required to compensate for the rejected material.

4a. Sterilize tubes by autoclaving at 121°C for 15 min under slow exhaust or liquid cycle.

Assessing and Controlling Microbial Contamination in Cell Cultures

Autoclaving times indicate the time necessary to hold the medium at the 121°C temperature for 15 min, not the time selected to run an autoclave cycle (UNIT 1.4; Table 1.4.1). Autoclave efficiency is very much machine and maintenance specific. However, an autoclave cycle of 20 min for a single 6 × 12 test tube rack of bacteriological medium, and a 32-min cycle for a 2-liter flask containing 1 liter of bulk medium, provide general guidelines for achieving the 121°C for 15 min sterilization criteria for these medium/vessel configurations.

5a. Remove tubes of medium from autoclave immediately after sterilization cycle is completed and/or when autoclave gauges indicate atmospheric pressure in the auto-

1.5.2 Supplement 1


clave chamber. Allow medium to cool to ambient room temperature in a location not subject to excessive air currents or temperature fluctuations. Oversterilization or prolonged holding of bacteriological media at elevated temperatures will severely affect performance. Cooling of just-autoclaved media in laboratory areas with significant temperature fluctuations or personnel movement can cause environmental contamination of otherwise sterile media.

6a. When tubes of medium reach ambient temperature, fully tighten screw caps and store tubes at 4° to 8°C until further quality control checks or until use (up to 6 to 9 months). Fluid thioglycollate is a medium formulated to detect slightly anaerobic bacteria and contains a small amount of agar to retard atmospheric oxygen diffusion into the medium, cysteine as a reducing agent, and methylene blue as an oxygen indicator. Freshly prepared fluid thioglycollate will have a very small zone of aerobiosis at the medium surface/head space interface, as indicated by a slight purple to orange band of oxidized methylene blue. As oxygen slowly continues to permeate with prolonged storage of the medium, the pigmented, oxidized band will enlarge. Do not use the medium if the color has changed to orange in greater than the top 25% of the medium. Oxidized fluid thioglycollate may be rejuvenated, but only once, by steaming the tubed medium in a boiling water bath to purge gaseous oxygen from the medium, and then cooling just prior to use. Other bacteriological media cited here have a shelf life of 6 to 9 months when stored in the dark at 4° to 8°C. Any of the liquid media cited above may be adapted for agar plate or test tube slant use by the addition of agar as the solidification agent as noted below.

For bulk agar media for plates: 1b. Reconstitute trypticase soy agar, Sabouraud’s dextrose agar, and YM agar per manufacturer’s instructions in 10-megaohm (or higher) distilled or deionized water. Use an autoclavable container capable of holding at least twice the volume of the medium being prepared (e.g., use a 2-liter Erlenmeyer flask to autoclave 1 liter of medium) to avoid boiling over during the auotclave cycle. 2b. Sterilize the bulk medium by autoclaving at 121°C for 15 min under slow exhaust or liquid cycle (see step 4a annotation). 3b. Cool in a water bath to ∼50°C. 4b. Add 50 ml/liter (5%) defibrinated sheep blood aseptically to trypticase soy agar. It is critical that medium be cooled to 45° to 50°C before sheep blood is added.

5b. Dispense medium aseptically in 24-ml aliquots to 100 × 15–mm sterile plastic disposable petri dishes. A semiautomated repeat-volume filling unit fitted with a weight on the inlet line to sink the tubing to the bottom of the flask of medium will greatly alleviate drawing of surface foam from the bulk flask to the petri dishes. Surface bubbles most often occur when postautoclaving agitation of the bulk flask is necessary to incorporate heat-labile additives such as sheep blood. A small number of media bubbles can be removed from the plates by lightly passing the flame of a Bunsen burner across the surface of the plated medium as soon as it is dispensed.

6b. Place plates in stacks of 10 to 20 and allow to cool and solidify overnight at room temperature. Store in vented plastic bags at 4° to 8°C until quality control checks or use (up to 12 weeks). Cooling the plates in stacks of 10 to 20 retards formation of excessive condensation on the lids of the plates. Cell Culture


Supplement 1

Prepare test samples 7a. For lot sample preparation of cryopreserved ampules: Use a 1-ml serological pipet to pool and mix the contents of ∼5% of the cell culture ampules prepared from each freeze lot. 7b. For cell culture vessels: Examine cell culture vessels individually under low power, preferably with phase contrast, using an inverted microscope. Look for aberrant growth or appearance of the cells. Aseptically remove 5-ml aliquots from suspect cultures to use for further examination and testing. Quarantine any suspect cultures or containers to ensure that they will not be inadvertently mixed with and cross-contaminate clean cultures. Unless the cultures are heavily contaminated, microbial growth will not be readily evident under low-power magnification. Mycelial fungal contamination sometimes is first noticed macroscopically by the appearance of small “cottony” or “lint-like” debris in the culture vessel.

8. Prepare wet mounts for microscopic evaluation of test samples and examine under oil immersion with high-power objectives (≥1000× magnification). Bacterial contamination is recognized by the presence of small, uniformly sized spheres, rods, or spirals scattered throughout the field. The organisms may be individual, in clusters, or in chains. Rod-shaped bacteria may contain bright, refractile spores. The bacteria may be motile. Fungal yeast contamination appears as ovoid, fairly regularly sized nucleated cells scattered throughout the microscopic field. The yeast may be individual, in the process of budding off smaller daughter cells, or in short chains. Mycelial fungal contamination is characterized by the presence of long filaments or pieces of broken filaments in the microscopic field. Fungal spores, frequently appearing as spherical objects covered with spines or other protrusions, might be observed. Low-level contamination may not be detected under the microscope even by a trained microbiologist, particularly if the cell culture sample contains much debris.

9. If culture contains antibiotics, wash prior to inoculation of microbiological test media by centrifuging at 2000 × g for 20 min (at room temperature or 4° to 8°C), removing the supernatant, and resuspending the pellet in an equal volume of antibiotic-free medium. Repeat for a total of three washes to eliminate traces of antibiotics that might interfere with microbial cultivation. Inoculate microbiological media with test samples 10. For each test sample, inoculate each of the following with 0.3-ml aliquots of cell suspension: 2 tubes of brain heart infusion 2 tubes of fluid thioglycollate medium 2 tubes of HTYE broth 2 tubes of soybean/casein digest broth 2 plates of trypticase soy agar with 5% sheep blood 2 plates of Sabouraud’s dextrose agar, Emmon’s modified 2 plates of YM agar. 11. Incubate one plate of trypticase soy agar (with 5% sheep blood) at 37°C aerobically and the other at 37°C under 5% CO2. 12. Incubate one sample each of the other media at 26°C and the other sample at 35° to 37°C. Assessing and Controlling Microbial Contamination in Cell Cultures

13. Examine all inoculated media daily for 14 days. Usually, visual evidence of bacterial growth appears within 72 hr; fungal growth within 96 hr. Low-level contamination, or proliferation of contaminants previously stressed by the presence of antibiotics or other adverse culture conditions, may take longer to appear.

1.5.4 Supplement 1


Positive bacterial and yeast growth appears as turbidity or the formation of a precipitate in liquid media. In an undisturbed test tube, growth may be restricted to a pellicle of growth at the surface of the liquid. On solid media, these bacteria and yeast appear as distinct, slightly convex, discretely isolated circular or confluent areas of microbial colonial proliferation, most often off-white or yellow in color. Filamentous fungal colonial growth is characterized by the appearance of typical cottony, whitish-gray to green to black mold on plates.

14. Autoclave and discard any general-use cell culture preparations that are positive for contamination. If a cell culture that has tested positive is critical to maintain, repeat the microbial evaluation. If still positive, autoclave and discard the culture. If it is necessary to attempt to clean a microbially contaminated critical cell culture, see Basic Protocol 3. Reusable glassware from discarded contaminated cultures should be decontaminated by autoclaving, cleaned, and depyrogenated by dry heat (UNIT 1.4).

TESTING FOR MYCOPLASMA CONTAMINATION BY DIRECT CULTURE This protocol describes the direct detection of mycoplasma contamination by screening with microbiological media designed to encourage proliferation of mycoplasma. Total incubation time for this method is ∼35 days. This schedule is necessary to detect low levels of mycoplasma contamination that might otherwise be scored as false negatives.

BASIC PROTOCOL 2

Materials Cell line for testing Mycoplasma broth medium (see recipe): 6 ml medium in 16 × 125–mm screw-cap test tubes Mycoplasma agar plates (see recipe): 10 ml solidified medium in 60 × 15–mm petri dishes 37°C incubators: one without CO2 and one humidified with 5% (v/v) CO2 Inverted microscope with 100 to 300× magnification NOTE: To avoid inadvertent contamination of clean cell lines, mycoplasma testing should be segregated to a laboratory not used for general cell culture work. 1a. For adherent cultures: Select a cell culture that is near confluency and has not received a fluid renewal within the last 3 days. Remove and discard all but 3 to 5 ml of the culture medium. Scrape a portion of the cell monolayer into the remaining culture medium using a sterile disposable scraper. 1b. For suspension cultures: Take the test sample directly from a heavily concentrated culture that has not received a fresh medium supplement or renewal within the last 3 days. Samples can also be taken directly from thawed ampules that have been stored frozen.

2. Inoculate 1.0 ml of the test cell culture suspension into 6 ml mycoplasma broth medium in a 16 × 125–mm screw-cap test tube. Also inoculate 0.1 ml of the test sample onto the center of a 60 × 15–mm mycoplasma agar plate. 3. Incubate the broth culture aerobically at 37°C. Incubate the agar plate in a humidified 37°C, 5% CO2 incubator. Observe broth culture daily for development of turbidity and/or shift in pH (medium becomes redder for alkaline shift, yellower for acid shift). As an alternative, a self-contained anaerobic system such as the GasPak equipment (Becton Dickinson Microbiology Systems) can be used in conjunction with a standard 37°C incubator to provide proper CO2 levels.

Cell Culture


Supplement 1

Figure 1.5.1 Mycoplasma colonies at 230× magnification. Figure provided by W. Siegel, Bio-Whittaker, Inc.

4. After 5 to 7 days of incubation and again after 10 to 14 days, remove a 0.1-ml sample from the broth culture and inoculate a fresh mycoplasma agar plate. Incubate these plates as in step 3. 5. Using an inverted microscope at 100 to 300× magnification, examine the agar plates weekly for at least 3 weeks for mycoplasma colony formation and growth. Assessing and Controlling Microbial Contamination in Cell Cultures

Mycoplasma colonies range from 10 to 55 ìm in diameter and classically look like a fried egg, with the dense center of the colony embedded in the agar and the thinner outer edges of the colony on the surface of the substrate (Fig. 1.5.1). However, much variation in colony morphology occurs between species and culture conditions. Although colonies typically

1.5.6 Supplement 1


appear within 4 days after inoculation, plates should be kept for the full incubation period before scoring them as negative.

6. To confirm presumptive mycoplasma colonies, subculture a small (∼1 cm2) section of the suspicious area of the agar plate into a tube of mycoplasma broth medium, incubate up to 14 days, and observe as in step 3. When other detection methods are not available, the ability to subculture presumptive mycoplasma colonies will help to differentiate authentic mycoplasma presence from artifacts such as air bubbles, tissue culture cells, or pseudocolonies.

INDIRECT TESTING FOR MYCOPLASMA BY STAINING FOR DNA The total time for this protocol, which includes the use of an indicator cell culture, is 6 days. The staining procedure itself takes ∼1 hr. The use of an indicator cell culture provides a number of advantages. The indicator cell line supports the growth of more fastidious mycoplasma species. Both positive and negative controls are thus readily available for direct comparison with the culture samples being tested. Selection of a proper indicator cell is important to the success of this procedure. It must first have good viability. Transformed cell lines are not recommended as indicators as they produce significant nuclear background fluorescence, which interferes with interpretation of results. Cell lines that produce much debris, such as hybridomas, are also not recommended as indicator cells because of the amount of positive staining artifacts that confuse interpretation of results.


Materials Complete EMEM-10: Eagle’s minimum essential medium (EMEM) with Earle’s salts (Life Technologies), 100 U/ml penicillin, 100 µg/ml streptomycin, and 10% (v/v) bovine calf serum (see UNIT 1.2 for media preparation methods) Indicator cell line: e.g., African green monkey cell line Vero (ATCC #CCL81) or 3T6 murine cell line (ATCC #CCL96) Cell culture for testing Mycoplasma hyorhinis (ATCC #29052) or a known mycoplasma-infected cell line to use as a positive control, actively growing Fixative: 3:1 (v/v) absolute methanol/glacial acetic acid Hoechst stain (see recipe) Mounting medium (see recipe) 60 × 15–mm culture dishes, sterile No. 1 or no. 11⁄2 coverslips, sterilized by autoclaving (UNIT 1.4) 37°C, 5% (v/v) CO2/95% air incubator NOTE: To avoid inadvertent contamination of clean cell lines, mycoplasma testing should be segregated to a laboratory not used for general cell culture work. Prepare indicator cell cultures 1. Aseptically place a sterile glass no. 1 or 11⁄2 coverslip into each sterile 60 × 15–mm culture dish. Use two culture dishes for the positive control, two dishes for the negative control, and two dishes for each test sample.

2. Aseptically dispense 3 ml complete EMEM-10 into each culture dish. Make certain that each coverslip is totally submerged and not floating on top of the medium.

3. Prepare a single-cell suspension of the indicator cell line in complete EMEM-10 at a concentration of 1.0 × 105 cells per ml.

Cell Culture


Supplement 1

4. Inoculate 1 ml indicator cell suspension into each culture dish. 5. Incubate overnight at 37°C in a 5% CO2/95% air incubator. 6. Microscopically examine cultures to verify that the cells have attached to the glass coverslip. Code the top of each culture dish for identification purposes (to record the test samples to be inoculated). Inoculate test samples 7. Add 0.5 ml/dish complete EMEM-10 to two culture dishes for negative controls. 8a. For adherent cultures: Select a test cell culture that is near confluency and has not received a fluid renewal within the last 3 days. Remove and discard all but 3 to 5 ml of the culture medium. Scrape a portion of the cell monolayer into the remaining culture medium using a sterile disposable scraper. 8b. For suspension cultures: Take the test sample from a heavily concentrated culture that has not received a fresh medium supplement or renewal within the last 3 days. Samples can also be taken directly from thawed ampules from frozen stocks.

9. For each test sample, add 0.2 to 0.5 ml/dish test sample to two culture dishes. 10. Add 0.5 ml/dish actively growing Mycoplasma hyorhinis to two culture dishes for positive controls. CAUTION: To prevent spread of mycoplasma, infected strains should be destroyed and removed as quickly as possible. Benches and incubators should be cleaned. For safety of the investigator, BSL 2 laboratory conditions are appropriate. Alternatively, a known mycoplasma-infected cell line can be used.

11. Return the cultures to the CO2 incubator and allow to incubate undisturbed for 6 days. Cultures should be at 20% to 50% confluent. Confluence can interfere with microscopic examination for mycoplasma.

Fix, stain, and mount coverslips 12. Remove cultures from incubator. Aspirate medium and immediately add 5 ml fixative to each culture dish. Incubate for 5 min. Do not allow the culture to dry between removal of the culture medium and addition of the fixative.

13. Aspirate fixative from each culture dish and repeat fixation for 10 min. These fixing times are minimal. Additional fixation time will not harm the procedure and can be beneficial for some preparations.

14. Aspirate the fixative and let the cultures air dry. Dry completely and store in a 60-mm petri or tissue culture dish if samples are to be accumulated at this stage for later staining.

15. Add 5 ml Hoechst stain to each culture dish, cover, and let stand at room temperature for 30 min. 16. Aspirate the stain and rinse each culture three times with 5 ml distilled water. 17. Aspirate well so that the glass coverslip is completely dry. Let air dry if necessary. 18. Place a drop of mounting medium on a clean glass microscope slide. Assessing and Controlling Microbial Contamination in Cell Cultures

19. Use forceps to remove the glass coverslip containing the fixed cells from the culture dish and place face up on top of the mounting medium, being careful to eliminate air bubbles.

1.5.8 Supplement 1


20. Add a second drop of mounting medium onto the top of the specimen coverslip and cover with a larger clean coverslip, being careful to eliminate air bubbles. 21. Label each slide to identify the specimen. 22. Observe each specimen by fluorescence microscopy at 500× using immersion oil. Use a blue glass excitation filter (330/380 nm) in combination with a 440-nm barrier filter (see UNIT 4.2). Compare test samples to positive and negative controls. The nuclei of the indicator cells appear as large (∼20 ìm), ovoid fluorescing bodies. Mycoplasma will appear as small fluorescing particles (0.1 to 1.0 ìm) that are regular in shape and size in the cytoplasm or in intercellular spaces. If infection is heavy, the particles may be tightly clustered in some areas. With low-level contamination, not all cells will be infected. Thus, all of the slide should be examined.

INDIRECT TESTING FOR MYCOPLASMA BY PCR Kits are now available from a number of manufacturers for the detection of mycoplasma in cell cultures or other material using PCR. The kits, although expensive and requiring the expertise and equipment needed for molecular procedures, provide advantages with quick results (1 day) and the ability to speciate the contaminant and thus potentially identify its source. The PCR kits are also useful for detecting mycoplasma in cultures prone to forming artifact debris that often obscures definitive interpretations with staining detection methods. The procedure described here uses primers from a commercially available kit and a nested PCR assay that amplifies the spacer region between the 16S and 23S rRNA genes of mycoplasmas. This specific protocol details procedures using cell cultures; however, instructions provided with the kit also explain how to make modifications for analyzing test samples of serum or frozen cells. The novice practitioner of PCR methodologies is strongly urged to consult Sambrook et al. (1989) for specific details (also see APPENDIX 3).


Materials Cells for testing 10× PCR buffer (usually provided with Taq polymerase) 2.5 mM 4dNTP mix: 2.5 mM each dGTP, dCTP, dTTP, and dATP 25 mM MgCl2 5 U/µl Taq DNA polymerase Mineral oil (if needed for thermal cycler) Mycoplasma detection kit (ATCC), containing first- and second-stage primer mixtures (total 7 primers), as well as two positive control mycoplasma DNAs (Mycoplasma pirum, Acholeplasms laidlawii) Thin-wall microcentrifuge tubes Aerosol-preventive micropipettor tips, sterile Positive-displacement micropipettors Picofuge Thermal cycler Additional reagents and equipment for agarose gel electrophoresis (see Support Protocol) NOTE: To avoid inadvertent contamination of clean cell lines, mycoplasma testing should be segregated to a laboratory not used for general cell culture work. NOTE: To avoid amplification of contaminating DNA from laboratory workers, room contaminants, or previous mycoplasma DNA amplifications, all PCR should be performed using aseptic technique (also see special considerations for PCR experiments in APPENDIX 2A).

Cell Culture


Supplement 1

Prepare test sample 1a. For confluent monolayers: Detach cell monolayer from flask surface with a cell scraper. Gently agitate the flask to dispense the cells in the medium. Transfer 0.5 ml cell suspension (∼5 × 104 cells/ml) to a sterile 1.5-ml microcentrifuge tube. 1b. For suspension cultures: Mix the suspension culture by gently pipetting to obtain an even dispersal of cells in the medium. Transfer 0.5 ml of the suspension (∼5 × 104 cells/ml) to a sterile 1.5-ml microcentrifuge tube. 2. Centrifuge at 12,000 × g for 10 min at 4°C. 3. Discard 400 µl supernatant and use a pipet to resuspend the cell pellet in the remaining 100 µl medium. Pipet gently to avoid formation of bubbles. The 100-ìl cell suspension is ready for PCR.

Run first-stage PCR 4. Using a permanent ink marker, label each thin-wall microcentrifuge tube (reaction tube) with appropriate test sample or control DNA codes. 5. Prepare a master mix of PCR reagents in sufficient quantity for all samples to be tested (n + 1 or 2 reactions) plus a minimum of two positive DNA controls and one negative control. Use the following volumes per reaction: 5 µl 10× PCR buffer 1 µl first-stage primer mixture 1 µl 2.5 mM 4dNTP mix 1 µl 25 mM MgCl2 (see annotation) 0.2 µl 5 U/µl Taq DNA polymerase Deionized water to 45 µl. Store the mix on ice until it is aliquotted. The optimal reaction conditions for this procedure are 10 mM Tris⋅Cl (pH 8.3 to 8.8), 50 mM KCl, 2.0 mM MgCl2, 50 ìM of each dNTP, and 1 U Taq polymerase. Check for the inclusion and final concentration of MgCl2 in the 10× PCR buffer supplied with the Taq polymerase and adjust the volume of MgCl2 in the reagent mix, if necessary, to give a final concentration of 2.0 mM. Adjust the amount of water in the mix accordingly for a final reaction mix volume of 45 ìl. NOTE: In this and all subsequent steps, use positive-displacement micropipettors and sterile aerosol-preventive micropipettor tips to prevent contamination of the amplification reaction.

6. Pipet 45 µl reaction mix into each sample and control reaction tube. 7. If the thermal cycler used requires mineral oil to minimize sample evaporation, add 40 to 60 µl of mineral oil to each tube. Keep the reaction tubes closed, except when aliquotting into them, to avoid possible cross-contamination.

8. Add 5 µl test sample (step 3) to the appropriate reaction tube containing reagent mix (final reaction volume 50 µl). If reaction tubes contain mineral oil, pipet the samples directly into the mix below the mineral oil layer (final tube volume 90 to 110 µl). Assessing and Controlling Microbial Contamination in Cell Cultures

9. Add 5 µl of each positive control mycoplasma DNA into separate positive control tubes. 10. Add 5 µl sterile deionized water into the negative control tube.

1.5.10 Supplement 1


11. Mix each tube thoroughly by flicking it with a finger, and centrifuge the tube briefly in a picofuge. 12. Place all the tubes into a thermal cycler and perform amplification using the following program: Initial step: 30 cycles:

Final step: Chill:

30 sec 30 sec 2 min 2 min 5 min indefinitely

94°C 94°C 55°C 72°C 72°C 4°C

(denaturation) (denaturation) (annealing) (extension) (extension) (hold).

Store PCR products at 4°C or on ice until further use (2 to 3 weeks). For longer periods, store at −20°C. Run second-stage PCR 13. Using a permanent ink marker, label each second-stage reaction tube with appropriate test sample or control DNA codes. 14. Prepare a master mix of reagents in sufficient quantity for all samples (n + 1 or 2 reactions), plus positive and negative controls, using the following volumes per reaction: 5 µl 10× PCR buffer 1 µl second-stage primer mixture 1 µl 2.5 mM 4dNTP mix 1 µl 25 mM MgCl2 (see step 5 annotation) 0.2 µl 5 U/µl Taq DNA polymerase Deionized water to 49 µl. Store the mix on ice until it is aliquotted. 15. Pipet 49 µl reaction mix to each sample and control tube. 16. If the thermal cycler used requires mineral oil to minimize sample evaporation, add 40 to 60 µl of mineral oil to each tube. Keep the reaction tubes closed, except when aliquotting into them, to avoid possible cross-contamination.

17. Carefully pipet 1 µl from the first-stage PCR reaction tube (step 12) into the second-stage reaction tube (final reaction volume 50 µl). If mineral oil was used, add the sample to the reagent mix below the oil layer (final tube volume 90 to 110 µl). 18. Mix each tube thoroughly by flicking it with a finger, and centrifuge the tube briefly in the picofuge. 19. Place all the tubes in the thermal cycler and run the program as above (step 12). Store PCR products at 4°C or on ice until further use (2 to 3 weeks). For longer periods, store at −20°C. 20. Analyze reaction products by agarose gel electrophoresis (see Support Protocol).

Cell Culture


Supplement 1

SUPPORT PROTOCOL

AGAROSE GEL ELECTROPHORESIS OF PCR PRODUCTS Agarose gel electrophoresis is used to analyze the products of the second-stage PCR for the presence of mycoplasma sequences. The protocol is written for one 10-lane gel, which can accommodate up to four unknown samples, two positive controls, one negative PCR control, one negative electrophoresis control, and two molecular weight marker lanes. For additional samples, the procedure can be scaled by using a comb with additional wells, or additional gels can be run. Materials Agarose (e.g., NuSieve, FMC Bioproducts) 1× TBE electrophoresis buffer (APPENDIX 2A) 10 mg/ml ethidium bromide solution Second-stage PCR products from test samples and controls (see Alternate Protocol 2) 6× electrophoresis sample buffer (see recipe) Molecular weight marker (100-bp DNA ladder) Electrophoresis apparatus with a 10 × 14–in. gel tray and a 1-mm, 10-tooth comb Power supply UV light box Prepare 2.5% agarose containing ethidium bromide 1. Seal a 10 × 14–in. gel tray from an electrophoresis apparatus according to manufacturer’s instructions and place on a level surface. 2. Weigh 2.25 g agarose and place in a 250-ml Erlenmeyer flask. 3. Add 90 ml of 1× TBE electrophoresis buffer, swirl to mix, and heat to boiling to completely dissolve agarose. 4. Add 5.4 µl of 10 mg/ml ethidium bromide solution, swirl to mix, and cool to ∼55°C. CAUTION: Ethidium bromide is a mutagen and a potential carcinogen. Gloves should be worn and care should be taken when handling ethidium bromide solutions.

Cast gel 5. Pipet 80 ml agarose solution into the center of the gel tray. Remove any bubbles. 6. Gently place a 1-mm, 10-tooth comb into the gel mold. Allow gel to harden until it becomes milky and opaque in appearance (∼1 hr). 7. Remove tape or sealers from the gel mold. Place gel into electrophoresis tank. 8. Pour ∼950 ml of 1× TBE electrophoresis buffer into the electrophoresis tank. Gel should be totally submerged in buffer, but not covered more than 1 cm.

9. Gently remove the gel comb. Prepare sample 10. Add 10 µl of each second-stage PCR product to a separate microcentrifuge tube containing 2.0 µl of 6× electrophoresis sample buffer. Mix well. 11. Add 5 µl of 100-bp DNA ladder to 2 µl of 6× electrophoresis sample buffer. Mix well. Assessing and Controlling Microbial Contamination in Cell Cultures

Load and run the gel 12. Add 7 µl DNA ladder to each of the first and last wells (lanes 1 and 10) of the gel. 13. Add 12 µl of 6× electrophoresis sample buffer to one lane as a negative control.

1.5.12 Supplement 1


14. Add 12 µl unknown test samples and positive and negative PCR controls to the remaining wells. 15. Connect a power supply and run the gel at 75 V for ∼1 hr and 40 min. 16. View the gel on a UV light box. CAUTION: UV light is damaging to eyes and exposed skin. Protective eyewear should be worn at all times while using a UV light source. With the primers used in Alternate Protocol 2, the mycoplasmas commonly encountered as cell culture contaminants should generate a second-stage PCR DNA amplicon that ranges in size from 236 to 365 bp. In contrast, the A. laidlawii positive control should generate two amplicons of 426 bp and 219 bp, and the M. pirum positive control should generate a single 323-bp amplicon. No discrete amplicon band should be seen in the negative controls. Fuzzy bands 90%.

14. Plate 3–10 × 104 viable cells in 5 ml of fresh complete growth medium in a 25-cm2 tissue culture flask. The cells will attach to a new flask within 2 to 3 hr and begin to exhibit the characteristic spindle shape in 24 hr. Contamination occurs less frequently in flasks than in dishes or multiwell plates

15. Change medium every 3 to 4 days until the culture becomes confluent. Harvest fibroblasts as described in steps 10 and 11. See Figure 2.1.4 for the typical growth rate of human fibroblasts; their doubling time varies from 24 to 72 hr, depending upon the culture conditions. Human fibroblasts can be passaged up to 10 times without significant changes in morphology or growth rates. It is recommended, however, that frozen stocks be prepared after the second or third passage (see step 16). Maximal fibroblast growth requires 5% to 10% FBS (Fig. 2.1.5A). It is technically challenging to grow fibroblasts in the absence of added serum. On the other hand, there is minimal, if any, variation among FBS batches purchased from different vendors in their capacity to promote the growth of human fibroblasts (Fig. 2.1.5B). Human fibroblasts also grow well in the presence of heat-inactivated human serum, which may be used instead of FBS (Fig. 2.1.5C).

16. To freeze cells, resuspend in ice-cold 10% DMSO/90% FBS or 10% DMSO/90% complete DMEM at 0.3–1 × 106 cells/ml. Dispense into 1.5-ml cryotubes at 1 ml/tube and freeze first at −20°C, then move on to −80°C, and finally place in liquid nitrogen. The −20°C and −80°C freezing steps may be performed in a styrofoam box to promote a gradual drop in temperature. Commercially available cell-freezing instruments may also be used for this purpose.

Establishment of Fibroblast Cultures

Figure 2.1.2 (at right) Microscopic appearance (magnification, 40×) of fibroblast cultures established from a newborn foreskin sample using the skin explant culture system (panels A to C) or the cell dissociation culture system (panels D to F). (A) Skin explant, day 1; (B) skin explant, day 5; (C) skin explant, day 14. (D) Dissociation culture, day 1; (E) dissociation culture, day 5; (F) dissociation culture, day 14. Note that fibroblasts migrate out from the edge of a skin specimen (panel B) and become confluent, except for the area where the original skin specimen was located (panel C). In dissociated-cell cultures, fibroblasts attach (panel D), spread on culture plates (panel E), and become confluent (panel F).


A

B

C

D

E

F

Preparation and Isolation of Cells


A

B

C

D

Figure 2.1.3 Identification of fibroblasts. Fibroblasts change their morphology depending upon the extent of confluency (or cell density). (A) Morphology at low density (10% to 20% confluence; magnification, 40×); (B) morphology at high density (100% confluence; magnification, 40×). (C) Indirect immunofluorescence staining of human fibroblast cultures with antibodies against type I collagen (magnification, 100×). (D) Indirect immunofluorescence staining of human fibroblast cultures with control antibodies (magnification, 100×). Briefly, fibroblasts were cultured for 2 days on LabTek chamber slides, fixed in 3% paraformaldehyde in PBS, permeabilized with 0.1% Triton X-100, and then subjected to immunofluorescence staining (UNIT 4.3) with rabbit anti-type I collagen (Chemicon), followed by labeling with FITC-conjugated anti-rabbit IgG (Jackson Immunoresearch).



Number of cells (x106)

1

0.1

0.03

0.01 0

2

4

7

9

11

14

Culture period (days)

Figure 2.1.4 Fibroblast growth curve. A third passage of human fibroblast culture was plated on 35-mm dishes at either 30,000 (triangles) or 100,000 (circles) cells per dish, and cultured in complete RPMI. At the indicated time points, cultures were harvested by incubation with 0.3% trypsin/25 mM EDTA and counted to determine cell number. Note that cells grow relatively rapidly, with an approximate doubling time of 24 hr, and then stop dividing as they reach confluency.

B

C [3H]Thymidine uptake (cpm x 10-3)

[3H]Thymidine uptake (cpm x10-3)

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Figure 2.1.5 Serum requirement for fibroblast growth. A third passage of human fibroblasts (3000 cells/well) was cultured in flat-bottom 96-well plates, pulsed with [3H]thymidine on day 3, and harvested on day 4 using an automated cell harvester. The culture media were supplemented with (A) different concentrations of FBS; (B) 10% FBS from different vendors; and (C) different concentrations of human serum. Data shown are the means and standard deviations from triplicate cultures.



ALTERNATE PROTOCOL

DISSOCIATED FIBROBLAST CULTURE Although more complicated than the skin explant culture (see Basic Protocol), the dissociated fibroblast culture described in this protocol is more suitable for those experiments that require relatively large numbers of fibroblasts. After removal of the epidermis by dispase treatment, fibroblasts are released from the remaining dermis by enzymatic treatment with trypsin. The resulting dermal cells are then plated in suspension onto tissue culture plates. The most tricky step is the enzymatic digestion of dermal tissues. Investigators may need to compare various conditions (e.g., batches of trypsin, trypsin concentrations, and incubation periods) to maximize the cell yield while maintaining the cell viability. Alternatively, collagenase, which is less cytotoxic than trypsin, can be used for the same purpose. Additional Materials (also see Basic Protocol) 1000 U/ml collagenase type IA in PBS (see recipe for PBS; store enzyme solution up to 3 months at −20°C) Nylon mesh (85-µm mesh; Tetko; cut into 5-cm square, wrap in aluminum foil, and sterilize by autoclaving) CAUTION: When working with human blood, cells, or infectious agents, appropriate biosafety practices must be followed. NOTE: Use Milli-Q water or equivalent in all protocol steps and for preparing all solutions. Prepare dissociated cell suspension 1. Wash skin samples in PBS, remove the subcutaneous tissues, remove the epidermis by enzymatic digestion, wash the dermal sample in PBS, and cut the sample into small squares (see Basic Protocol, steps 1 to 5). Since the trypsin that is used to digest the dermal connective tissue also dissociates epidermal cells, the epidermis must first be separated from dermal layer. Otherwise, the resulting cultures will be heavily contaminated by epidermal keratinocytes.

2. Place 10 to 20 dermal pieces in a 15-ml polypropylene tube with 3 ml of 0.3% trypsin/PBS and incubate 10 min in a 37°C water bath, inverting the tube several times every 2 to 3 min. Alternatively, incubate 10 to 20 dermal pieces 1 to 2 hr with 3 ml of 1000 U/ml collagenase at 37°C, agitating every 20 to 30 min. 3. Add 3 ml of ice-cold complete growth medium (DMEM or RPMI containing 10% FBS) to stop the reaction. Vortex the tube vigorously several times. Although fibroblasts detach from collagen fibers after treatment with trypsin or collagenase, mechanical agitation is required for releasing them into the solution. Do not vortex before the addition of complete medium.

4. Pass the fibroblast suspension through 85-µm nylon mesh (placed over the top of a tube) to remove dermal debris. 5. Centrifuge 10 min at 150 × g, 4°C. Aspirate the supernatant, then resuspend the pellet in 100 to 200 µl of complete growth medium. 6. Count total and viable cells (see Basic Protocol, step 13).


The cell viability varies depending upon the conditions used for enzymatic digestion. Cutting skin sample into smaller sizes usually increases cell recovery as well as cell viability.


Culture fibroblasts 7. Plate 3–10 × 104 cells in 5 ml of complete growth medium in a 25-cm2 tissue culture flask and begin incubation. Viable fibroblasts will attach to the flask within 24 hr and begin to exhibit the spindle-shape in 2 to 3 days (Fig. 2.1.2D and E).

8. Gently remove the medium containing nonadherent cells and add fresh medium on day 2. Because the presence of dead cells in culture affects the growth of viable fibroblasts, nonadherent, dead cells must be removed from the culture.

9. Change medium every 3 to 4 days until the culture becomes confluent. As fibroblasts become overconfluent, they appear as bundle clusters instead of spindleshaped cells (Fig. 2.1.2 and Fig. 2.1.3). Harvest the cells before they reach this level.

10. Harvest fibroblasts by washing with PBS followed by incubation with trypsin/EDTA solution. Passage fibroblasts and prepare frozen stocks (see Basic Protocol, steps 10 to 16). REAGENTS AND SOLUTIONS Use Milli-Q water or equivalent in all recipes and protocol steps. For common stock solutions, see APPENDIX 2A; for suppliers, see SUPPLIERS APPENDIX.

Complete growth medium 500 ml DMEM or RPMI 1640 (Life Technologies or Sigma) 60 ml FBS (heat-inactivated 60 min at 56°C; APPENDIX 2A) 5 ml 1 M HEPES buffer solution (Life Technologies) 5 ml 100× nonessential amino acid mixture (Life Technologies) 5 ml 100× L-glutamine (Life Technologies) 5 ml 100× penicillin/streptomycin (Life Technologies) 5 ml 100× sodium pyruvate (Life Technologies) Store up to 1 month at 4°C Phosphate-buffered saline (PBS) 4 liters distilled water 32 g NaCl (140 mM final) 0.8 g KH2PO4 (1.5 mM final) 8.7 g Na2HPO4⋅7H2O (8.1 mM final) 0.8 g KCl (2.7 mM final) Adjust the pH to 7.4 with 1 N NaOH Store indefinitely at room temperature Trypsin/EDTA solution Prepare the following stock solutions: 0.3% (w/v) trypsin (from bovine pancreas; Sigma) in PBS (see recipe for PBS) 1% (w/v) tetrasodium EDTA in PBS (see recipe for PBS) Store stock solutions up to 3 months at −20°C Combine 97.5 ml 0.3% trypsin/PBS and 2.5 ml 1% EDTA/PBS. Store trypsin/EDTA solution up to 1 week at 4°C.



COMMENTARY Background Information


Fibroblasts are the major cellular component of connective tissues, where they play an important role in maintaining structural integrity. They produce and secrete a wide array of extracellular proteins, including proteinases, thereby regulating the biochemical composition and remodeling of tissues. Because biopsy samples can be easily obtained from skin, this tissue serves as a most convenient source of fibroblasts. Skin is composed of continually renewing multilayered squamous epithelium (the epidermis), connective tissue (the dermis), and subcutaneous (adipose) tissue. Human epidermis contains, in addition to keratinocytes (epithelial cells producing keratin intermediate filaments), relatively small numbers of Langerhans cells (antigen-presenting cells of the dendritic cell lineage) and melanocytes (which produce pigment granules called melanosomes). By contrast, mouse epidermis contains keratinocytes, Langerhans cells, and resident γδ T cells called “dendritic epidermal T cells.” The dermis, in both human and mouse skin, is a fibrous and filamentous connective tissue that contains fibroblasts, endothelial cells, mast cells, macrophages, and occasionally other leukocyte populations. Despite the complexity of cellular composition, relatively pure fibroblast cultures can be obtained from skin specimens without sophisticated purification processes. This is primarily due to the fact that fibroblasts grow rapidly and continuously when cultured in the presence of serum, whereas other cell types require additional growth factors (e.g., epidermal growth factor or keratinocyte growth factor for keratinocytes), or show very little mitotic activity in vitro (Schuhmachers et al., 1995). Morphological features—e.g., elongated cell bodies, oval nuclei, and linear or bundlelike alignment of cellular distribution—serve as conventional markers of fibroblasts in culture (Fig. 2.1.2 and Fig. 2.1.3). It is important to emphasize, however, that fibroblasts change their morphology dramatically depending upon the culture conditions, especially the extent of confluency (compare Fig. 2.1.3 panels A and B). Unfortunately, there is no antibody available that recognizes fibroblasts selectively. On the other hand, the absence of specific markers that are expressed by other dermal components (e.g., cytokeratin in keratinocytes, VCAM-1 on endothelial cells, IgE receptor on mast cells, and CD14 on macrophages) serves as a pheno-

typic marker of fibroblasts (Xu et al., 1995). Production of large amounts of type I collagen, as detected by immunofluorescence staining (Fig. 2.1.3C and D), can be used as a functional marker (Schuhmachers et al., 1995). Nevertheless, because the fibroblast cultures established by the standard protocols described in this unit are rarely “contaminated” by other cell types, especially after a few passages, it is generally accepted that they can be used as “fibroblasts” without further characterization. Fibroblasts grow rapidly, with a doubling time of 24 to 72 hr (Fig. 2.1.4), and can be passaged successfully >10 times. Because of this outstanding mitotic potential, fibroblasts have been used for a variety of investigative purposes. For example, they serve as useful tools for studying the function and metabolism of extracellular matrix proteins as well as other fundamental aspects of cell biology. Fibroblast cultures established from patients with inherited disorders have often been used to identify genetic abnormalities. Moreover, autologous fibroblasts can be used as a “vector” in gene therapy to deliver transgenes into patients (Suhonen et al., 1996; Nolta and Kohn, 1997). Fibroblast lines generated from healthy human volunteers can be purchased from American Type Culture Collection (ATCC). The protocols described in this unit are also applicable to other animal species. For example, in the author’s laboratory, several fibroblast lines from rats and rabbits have been developed using the same protocols as described for mouse fibroblasts. After enzymatic separation of the dermal compartment, the remaining epidermal portion can be used to grow epidermal cells, such as keratinocytes and melanocytes. As sources of human fibroblast cultures, newborn foreskin (obtained in circumcision), skin samples excised during surgical operations, cadaver skin (obtained from the transplantation unit), or skin biopsies are routinely used in the author’s laboratory. Fibroblast cultures can be established from relatively small skin specimens; the author routinely uses 4-mm punch biopsies for this purpose (Pandya et al., 1995). It is also practical and feasible to establish fibroblast cultures without sacrificing experimental animals; mouse “ear punch” samples are used for this purpose by the author.

Critical Parameters Because fibroblast outgrowth occurs predominantly from sharp edges of skin speci-


mens, it is crucial to use fine razor blades or surgical scalpels for cutting skin into small pieces. The author routinely uses disposable no. 22 surgical blades for this purpose. Drying of skin specimens is another common cause of poor fibroblast outgrowth. This can be avoided by adding a few drops of PBS while cutting skin specimens. If the surgical blades and forceps are to be soaked with 70% ethanol for sterilization, they should be rinsed well in PBS before use. When performed appropriately, outgrowing fibroblasts should become detectable within 3 to 4 days in skin explant cultures (Fig. 2.1.2B). In cell-dissociation cultures (see Alternate Protocol), the enzymatic digestion process is the most critical; if the viable cell count of resulting suspensions is 10 years in liquid nitrogen. Thus, it is suggested that several aliquots be frozen at a relatively early phase in culture (e.g., after the second or third passage). Because the original features of the cells may be altered during extended culture periods, it is not recommended that they be cultured continuously without experimental usage. If the cells suddenly stop dividing, or if the growth rate accelerates, the cultures need to be replaced. In the author’s laboratory the original cultures are routinely discarded after the sixth passage and new cultures are started from a frozen stock. Care should be taken in harvesting fibroblasts from culture plates. Although fibroblasts are more resistant to contact inhibition than other cell types (e.g., keratinocytes), it is suggested that cells be harvested during their exponential growth phase (Fig. 2.1.4). In the author’s laboratory, after removal of culture medium, culture plates are routinely washed briefly with PBS, and then minimal amounts of 0.3% trypsin/25 mM EDTA are added. These plates can be incubated at room temperature under a microscope; as soon as the cells become rounded (before being released spontaneously from plates), the enzymatic reaction is stopped by the addition of ice-cold growth medium containing 10% FBS, and cells are harvested by tapping the culture vessel or gentle pipetting. These cells need to be centrifuged immediately

to remove trypsin and EDTA. The cell viability should be >90% by trypan blue exclusion.

Troubleshooting Bacterial or fungal contamination Clean the skin well with 70% ethanol before taking a biopsy. Check all the culture media and reagents, including PBS, dispase, trypsin, collagenase, and complete growth medium. Sterilize surgical blades and forceps with 70% ethanol. Always keep the incubator clean. Contamination by keratinocytes Remove the epidermis before setting up the fibroblast cultures. Treat the contaminated cultures with 0.5% dispase for 10 to 30 min at 37°C to remove keratinocytes. Because keratinocytes usually require special growth factors for continuous growth, they will eventually disappear in the first or second passage. Low cell yields or low cell viabilities after enzymatic digestion Optimize the concentrations, batches of enzymes, and incubation periods. Cut skin into smaller pieces. Use freshly prepared trypsin (or collagenase) solutions; they will lose enzymatic activity gradually when kept at 4°C. Alternatively, prepare enzyme solutions in large quantities, divide them into 5- to 10-ml aliquots, and freeze them at −20°C. Low cell viabilities during passage Care must be taken not to overtrypsinize cultures. Wash the culture with PBS before trypsin treatment. Use freshly prepared trypsin solutions. Add ice-cold complete growth medium to stop the enzymatic reaction immediately after fibroblasts round up as determined by examination under a microscope. Avoid excessive pipetting. Slow fibroblast growth Increase FBS concentrations (up to 10%); most of the commercially available FBS batches work well for fibroblast cultures (Fig. 2.1.5). Check the temperature, CO2 level, and humidity of the incubator. The growth rates of fibroblasts often slow down after >10 passages; thaw a frozen stock and start new cultures to avoid this.

Anticipated Results In skin explant cultures, outgrowing fibroblasts become detectable within 3 to 4 days



and continue to grow thereafter. When skin pieces from one 4-mm punch-biopsy sample are plated in a 35-mm tissue culture dish, it usually takes 3 to 5 weeks to obtain a confluent culture. Starting from one newborn foreskin sample, one can obtain 5 to 10 confluent 35-mm dishes in 3 to 5 weeks. Approximately 1–2 × 106 cells can be harvested from a confluent 35-mm dish. Cell yields are usually higher in the dissociated fibroblast preparation (see Alternate Protocol); after enzymatic digestion with trypsin, ∼1–3 × 105 cells can be harvested from a 4-mm punch-biopsy sample and 1–3 × 106 cells can be harvested from a newborn foreskin sample. The use of collagenase typically increases the yield up to 2-fold. When 1–2 × 105 cells are originally seeded in a 25-cm2 flask, they will become confluent within 2 to 3 weeks, producing 2–3 × 106 cells. After the first passage to new culture plates, fibroblasts begin to grow much faster, with a typical doubling time of 24 to 72 hr (Fig. 2.1.4).

Time Considerations Skin explant cultures can be set up in 30 to 60 min (without epidermal separation) or 2 to 4 hr (with epidermal separation). Dissociated fibroblast cultures take 3 to 4 hr, depending upon the extent of enzymatic digestion. Although it has been possible in the author’s laboratory to establish fibroblast cultures from 1- to 2-day-old skin samples kept in complete growth medium at 4°C, it is highly recommended that cultures be set up immediately after taking biopsy samples. Subsequent passages can be made in 15 to 30 min.

Literature Cited Nolta, J.A. and Kohn, J.B. 1997. Human hematopoietic cell culture, transduction, and analyses. In Current Protocols in Human Genetics (N.C. Dracopoli, J.L. Haines, B.R. Korf, D.T. Moir, C.C. Morton, C.E. Seidman, J.G. Seidman, and D.R. Smith, eds.) pp. 13.7.1-13.7.35. John Wiley & Sons, New York. Pandya, A.G., Sontheimer, R.D., Cockerell, C.J., Takashima, A., and Piepkorn, M. 1995. Papulonodular mucinosis associated with systemic lupus erythematosis: Possible mechanisms of increased glycosaminoglycan accumulation. J. Am. Acad. Dermatol. 32:199-205. Schuhmachers, G., Xu, S., Bergstresser, P.R., and Takashima, A. 1995. Identity and functional properties of novel skin-derived fibroblast lines (NS series) that support the growth of epidermalderived dendritic cell lines. J. Invest. Dermatol. 105:225-230. Suhonen, J., Ray, J., Blömer, U. and Gage, F.H. 1996. Ex vivo and in vivo gene delivery to the brain. In Current Protocols in Human Genetics (N.C. Dracopoli, J.L. Haines, B.R. Korf, D.T. Moir, C.C. Morton, C.E. Seidman, J.G. Seidman, and D.R. Smith, eds.) pp. 13.3.1-13.3.24. John Wiley & Sons, New York. Xu, S., Ariizumi, K., Caceres-Dittmar, G., Edelbaum, D., Hashimoto, K., Bergstresser, P.R., and Takashima, A. 1995. Successive generation of antigen-presenting, dendritic cell lines from murine epidermis. J. Immunol. 154:2697-2705.

Contributed by Akira Takashima University of Texas Southwest Medical Center Dallas, Texas



Preparation and Culture of Human Lymphocytes

UNIT 2.2

This unit describes procedures for preparation and culture of human lymphocytes and lymphocyte subpopulations obtained from peripheral blood. Because of ease of access, peripheral blood is the primary source of human lymphocytes (mononuclear leukocytes) used in most studies of lymphocyte function. Peripheral blood is a mixture of cells including lymphocytes, granulocytes, erythrocytes, and platelets. Density gradient centrifugation (see Basic Protocol 1) has proven to be an easy and rapid method for separation of lymphocytes from these other peripheral blood cell populations. Lymphocytes and platelets can be separated from granulocytes and erythrocytes according to their lower densities—they will float on top of a density gradient of Ficoll-Hypaque, whereas granulocytes and erythrocytes will traverse this fluid and collect at the bottom of the tube (Fig. 2.2.1). Monocytes/macrophages can then be separated from the other lymphoid-cell populations by adherence to plastic tissue culture vessels (see Basic Protocol 2). The procedures described in this section can be applied to the isolation of peripheral blood lymphocyte populations obtained either from whole blood or via a leukapheresis procedure. Human lymphocyte subpopulations can be purified based on their cell-surface display of specific distinguishing molecules that can be recognized by monoclonal antibodies. The physical basis for such separation procedures involves the coupling of antibody reagents to magnetic beads, which permit the rapid sequestration of cells that have been bound by the specific antibodies. T and B cells can be positively selected using monoclonal antibody–coated magnetic beads (see Basic Protocol 3) or by exposing the cells to monoclonal antibody and then purifying cells that have bound the antibody using magnetic beads coated with anti–immunoglobulin G (anti-IgG; see Alternate Protocol 1).

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Figure 2.2.1 Isolation of human lymphocytes on a Ficoll-Hypaque gradient. (A) Before centrifugation; (B) after centrifugation. Contributed by William E. Biddison Current Protocols in Cell Biology (1998) 2.2.1-2.2.13 Copyright © 1998 by John Wiley & Sons, Inc.


2.2.1

In addition, specific subpopulations can be isolated by negative selection, in which all unwanted subpopulations are removed using monoclonal antibodies and anti-IgG-coated magnetic beads (see Alternate Protocol 2). CAUTION: When working with human blood, cells, or infectious agents, appropriate biosafety practices must be followed. NOTE: All solutions and equipment coming into contact with living cells must be sterile, and proper aseptic technique should be used accordingly. NOTE: All culture incubations are performed in a humidified 37°C, 5% CO2 incubator unless otherwise specified. Some media (e.g., DMEM) may require altered levels of CO2 to maintain pH 7.4. BASIC PROTOCOL 1

PREPARATION OF LYMPHOCYTES BY FICOLL-HYPAQUE GRADIENT CENTRIFUGATION In this procedure, whole blood or white blood cells from leukapheresis donors are centrifuged in the presence of a density gradient medium to separate lymphocytes from other peripheral blood cell populations. Materials Anticoagulated whole blood or white blood cells from leukapheresis donor Phosphate-buffered saline (PBS), without calcium or magnesium (Bio-Whittaker), room temperature Ficoll-Hypaque solution (see recipe), room temperature Lymphocyte culture medium (LCM; see recipe), room temperature Iscove’s modified Dulbecco’s medium (IMDM; Life Technologies) containing 20% heparinized human plasma Freezing medium (see recipe) 50-ml conical centrifuge tubes Sorvall RT-6000B centrifuge with H-1000 rotor (or equivalent) 1.5-ml cryotubes (e.g., Nunc) Controlled-rate freezer (e.g., CryoMed from Forma Scientific) Liquid nitrogen freezer Additional reagents and equipment for counting cells and determining cell viability (UNIT 1.1) and flow cytometry (Robinson et al., 1998) Prepare Ficoll-Hypaque gradient For whole blood 1a. Pipet 15 ml of whole blood into a 50-ml conical centrifuge tube and add 25 ml room temperature PBS 2a. Using a 10-ml pipet, underlay with 10 ml room temperature Ficoll-Hypaque solution. For leukapheresis preparation 1b. Pipet 10 ml of cell suspension from a leukapheresis preparation into a 50-ml conical centrifuge tube and add 30 ml room temperature PBS. 2b. Using a 10-ml pipet, underlay with 7.5 ml room temperature Ficoll-Hypaque solution.


Separate cells 3. Centrifuge 20 min at 800 × g (2000 rpm in H-1000 rotor), 20°C, with the brake off.


4. Aspirate most of the plasma- and platelet-containing supernatant above the interface band (granulocytes and erythrocytes will be in red pellet; Fig. 2.2.1). Aspirate the interface band (which includes the lymphocytes) along with no more than 5 ml of fluid above the pellet into a 10-ml pipet, then transfer to a new 50-ml conical centrifuge tube, combining the bands from 2 to 3 Ficoll-Hypaque gradients into one 50-ml tube. Add PBS to 50-ml mark. 5. Centrifuge 10 min at 600 × g (1500 rpm in H-1000 rotor), 20°C, with the brake on. 6. Aspirate supernatants and resuspend the pellet in each tube with 10 ml room temperature PBS. Combine resuspended pellets into as few 50-ml tubes as possible. Add PBS to 50-ml mark in each tube used. 7. Centrifuge 15 min 300 × g (750 rpm in H-1000 rotor), 20°C, with brake on. This low-speed centrifugation permits as many platelets as possible to remain above the pellet of lymphocytes.

Process cell pellet 8. Aspirate platelet-containing supernatant and resuspend lymphocyte pellet in room temperature LCM. Count cells and determine number of viable cells by trypan blue exclusion (UNIT 1.1).

Counts

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Figure 2.2.2 Flow cytometric analysis of human lymphocytes isolated by Ficoll-Hypaque density gradient centrifugation stained with (A) fluorochrome-labeled anti–mouse Ig antibody alone (negative control), (B) fluorochrome-labeled anti-CD45 monoclonal antibody, (C) fluorochrome-labeled anti-CD3 monoclonal antibody, and (D) fluorochrome-labeled anti-CD19 monoclonal antibodies. FL1-H refers to the pulse height for the fluorochrome.



Supplement 7

9. Determine purity of lymphocyte preparation by flow cytometry (see, e.g., Robinson et al., 1998) using an anti-CD45 (anti–leukocyte common antigen) antibody (see Fig. 2.2.2 for typical results). 10. To cryopreserve cells, resuspend lymphocytes at twice the concentration desired in the freezing vials using MDM/20% human plasma. Add an equal volume of freezing medium, in increments, over a period of 1 to 2 min, mixing after each addition. Divide the cell suspension into 1-ml aliquots in 1.5-ml cryotubes, place cryotubes in a precooled 4°C controlled-rate freezer and freeze at −1°C/min until −50° or −60°C is attained. Finally, place cryotubes in the gaseous phase of a liquid nitrogen freezer. BASIC PROTOCOL 2

PREPARATION OF MONOCYTES/MACROPHAGES AND “DENDRITIC-LIKE” CELLS FROM LYMPHOCYTE POPULATIONS Monocytes/macrophages comprise 5% to 20% of the lymphocytes prepared by density gradient centrifugation (see Basic Protocol 1). To isolate this cell population, or to deplete these cells from the population of lymphocytes, the most expedient mechanism is to exploit the adherence property of monocytes/macrophages. By coating plastic tissue culture vessels with serum as a source of fibronectin and other components of the extracellular matrix, monocytes/macrophages, but not T and B cells, will adhere to these surfaces. This adherence can be significantly enhanced by the presence of recombinant human interleukin 3 (rhIL-3; Biddison et al., 1997). Cells with properties of dendritic cells and a markedly enhanced capacity for antigen processing and presentation can be generated from the monocyte/macrophage population by differentiation in the presence of rhIL-4 and granulocyte/macrophage colony stimulating factor (GM-CSF; Sallusto and Lanzavecchia, 1994). Materials Lymphocyte culture medium (LCM; see recipe), 37°C Lymphocyte population (see Basic Protocol 1) Recombinant human interleukin 3 (rhIL-3), interleukin 4 (rhIL-4), and GM-CSF (PeproTech) 5 mM tetrasodium EDTA in PBS (also available as Versene from Life Technologies), filter-sterilized using 0.22-µm Nalgene filter, prewarmed to 37°C 175-cm2 tissue culture flasks 50-ml conical centrifuge tubes Sorvall RT-6000B centrifuge with H-1000 rotor (or equivalent) Additional reagents and equipment for counting cells and determining cell viability (UNIT 1.1) and flow cytometry (Robinson et al., 1998) Allow monocyte/macrophage population to attach to plastic 1. Add 20 ml of 37°C LCM to 175-cm2 tissue culture flasks. Incubate 30 min. 2. Add 500 × 106 lymphocytes in 10 ml LCM to each flask plus rhIL-3 to a final concentration of 200 U/ml. 3. Incubate 3 hr, gently rocking flasks every hour. 4. Remove nonadherent cells by aspirating the medium. Wash flasks twice, each time with 20 ml 37°C LCM.


2.2.4 Supplement 7


Isolate macrophages/monocytes or “dendritic-like” cells To isolate monocytes/macrophages 5a. Add 20 ml of 5 mM EDTA/PBS to each flask and incubate 20 min to release adherent monocytes/macrophages. 6a. Vigorously pipet medium up and down to detach adherent cells, then transfer to 50-ml centrifuge tube. To generate and isolate “dendritic-like” cells 5b. Add 30 ml LCM containing 200 U/ml rhIL-4 and 200 U/ml GM-CSF to each flask and incubate 60 hr. 6b. Remove nonadherent cells and place in 50-ml centrifuge tubes, then add 20 ml of 5 mM EDTA/PBS to each flask and incubate 20 min to release adherent cells. Vigorously pipet medium up and down to detach adherent cells, then combine with nonadherent cells in 50-ml centrifuge tubes. Separate and analyze isolated cells 7. Centrifuge 10 min at 600 × g (1500 rpm in Sorvall H-1000 rotor), room temperature. Aspirate supernatant and resuspend pellet in LCM. 8. Count cells and determine number of viable cells by trypan blue exclusion (UNIT 1.1). 9. Determine purity of the monocyte/macrophage or “dendritic-like” cell populations by flow cytometry (see, e.g., Robinson et al., 1998). Monocytes/macrophages can be distinguished from T and B cells by the absence of cell-surface CD3 and CD19 and by the presence of CD14 or CD35. “Dendritic-like” cells can be distinguished from monocytes/macrophages by the presence of cell-surface CD1 molecules (Sallusto and Lanzavecchia, 1994).

POSITIVE SELECTION OF T AND B CELLS BY MONOCLONAL ANTIBODY–COATED MAGNETIC BEADS

BASIC PROTOCOL 3

In this procedure, T cells and B cells are positively selected based on their differential cell-surface expression of CD3 (T cells) or CD19 (B cells). Monoclonal anti-CD3 and anti-CD19 antibodies coupled to magnetic beads are commercially available. Aliquots of a lymphocyte population purified as described in Basic Protocol 1 are incubated in separate tubes with anti-CD3-coupled beads and anti-CD19-coupled beads. T cells and B cells will be bound by their specific antibody-coupled beads and are then physically separated from unbound cells by exposure of the tubes to a strong magnetic field. Unbound cells are removed and washed away, and the specifically bound cells are released by incubation with a soluble antiserum specific for mouse Fab fragments (Detachabead solution from Dynal) which competes with the bead-coupled monoclonal antibody that is bound to the surface of the cells and thus causes the cells to come off. The procedure here describes specific reagents for separation of T cells and B cells, but is directly applicable to separation of any lymphocyte subpopulation that can be distinguished by monoclonal antibody–coated beads—e.g., purification of CD4+ and CD8+ T cell populations with anti-CD4- and anti-CD8-coated magnetic beads. Materials Lymphocyte population (see Basic Protocol 1) Anti-CD3 and anti-CD19 antibodies for flow cytometry (Becton-Dickinson or Coulter) Anti-CD3- and anti-CD19-coated magnetic beads (Dynabeads M-450; Dynal) PBS without calcium and magnesium (Bio-Whittaker)



PBS/HSA: PBS without calcium and magnesium (Bio-Whittaker) containing 0.5% (w/v) human serum albumin (American Red Cross Blood Services) IMDM/HSA: Iscove’s modified Dulbecco’s medium (Life Technologies) containing 0.5% (w/v) human serum albumin Polyclonal anti–mouse Fab antiserum (Detachabead; Dynal) 15-ml conical centrifuge tubes Magnetic separation device (e.g., Dynal MPC-I or Advanced Magnetics Biomag Separator) Platform rocker (e.g., Clay Adams Nutator, Becton Dickinson Primary Care Diagnostics) Sorvall RT-6000B centrifuge with H-1000 rotor (or equivalent) Additional reagents and equipment for flow cytometry (Robinson et al., 1998) and for counting cells and determining cell viability (UNIT 1.1) Prepare and wash magnetic beads 1. Determine the approximate number of T cells and B cells in the starting population of lymphocytes by flow cytometry (or indirect immunofluorescence staining) using anti-CD3 and anti-CD19 antibodies, then determine the number of antibody-coated magnetic beads that will be required to purify the required number of T cells and B cells (5 to 10 beads will be needed for each specific target cell). 2. Based on the bead concentration supplied by the manufacturer, remove the required volume of anti-CD3- and anti-CD19-coated beads from the source vials and place each in a 15-ml conical centrifuge tube. Add PBS to the 14-ml mark and resuspend the beads. 3. Place tubes on magnetic separation device (vertical magnet) for 2 min, then gently aspirate supernatant, leaving beads clinging to one side of each tube. 4. Remove tubes from magnet. Add 10 ml PBS to each tube, resuspend beads, then place tubes on vertical magnet for 2 min. Aspirate PBS as in step 3. Perform magnetic separation 5. Resuspend each tube of beads in 2 ml PBS/HSA. Put tubes on ice for 15 min. 6. Resuspend lymphocyte population (≤200 × 106 cells) in 10 ml PBS/HSA. Place on ice for 15 min. 7. Add 5 ml of lymphocyte suspension to the tube with anti-CD3 beads and 5 ml to the tube with anti-CD19 beads. 8. Incubate 45 min with gentle tilting and rotation on a platform rocker at 4°C. 9. Place tubes on vertical magnet for 2 min. Aspirate nonadherent cells, taking care not to disturb the beads that are clinging to one side of each tube. 10. Add 5 ml IMDM/HSA to each tube. Gently resuspend beads, then place tubes on vertical magnet for 2 min. Aspirate nonadherent cells as in step 9. 11. Add 3 ml IMDM/HSA to each tube. Gently resuspend beads, then place tubes with their conical bottoms on the horizontal magnet. Incubate 2 min at room temperature. Release T cells and B cells from beads 12. Carefully remove 2.5 ml of supernatant. Tap tubes gently to resuspend beads. Preparation and Culture of Human Lymphocytes

13. Add 200 µl Detachabead solution. Incubate 30 min at room temperature with gentle resuspension every 5 min. Smaller volumes of Detachabead solution may be used when starting with smaller numbers of lymphocytes; see manufacturer’s instructions.


14. Add 3 ml PBS/HSA to each tube and resuspend beads. Place tubes on vertical magnet for 2 min, then aspirate and save supernatant containing detached cells, taking care not to disturb the beads clinging to the side of each tube. Repeat this step four times, saving and combining detached cell–containing supernatants from each separation. 15. Centrifuge detached cells 10 min at 600 × g (1500 rpm in H-1000 rotor), room temperature. 16. Aspirate supernatants and resuspend cells in IMDM/HSA. Count cells and determine number of viable cells by trypan blue exclusion (UNIT 1.1). 17. Determine purity of the T cell and B cell populations by flow cytometry using anti-CD3 and anti-CD19 antibodies (see, e.g., Robinson et al., 1998). POSITIVE SELECTION OF T AND B CELLS BY MONOCLONAL ANTIBODIES AND ANTI-IgG-COATED MAGNETIC BEADS


This procedure differs from Basic Protocol 3 in that it does not require the acquisition of separate magnetic beads coupled with individual monoclonal antibodies. The procedure is described for T and B cell separation, but can be applied to any lymphocyte subpopulation that can be distinguished by monoclonal antibodies. The principle of the technique is that a human mixed lymphocyte population is separately exposed to saturating amounts of either anti-CD3 or anti-CD19 mouse IgG monoclonal antibodies, unbound antibodies are washed away, then the cells that have bound these antibodies are physically separated from unbound cells by magnetic beads coated with goat anti-mouse IgG. The cells that are specifically bound to the goat anti-mouse IgG–coated beads are then detached by exposure to soluble antiserum against mouse Fab fragments. Additional Materials (also see Basic Protocol 3) Anti-CD3 and anti-CD19 IgG monoclonal antibodies (Becton Dickinson or Coulter) Goat anti-mouse IgG–coated magnetic beads (Dynabeads M-450; Dynal) Prepare cells and antibodies 1. Determine the approximate number of T cells and B cells in the starting population of lymphocytes by flow cytometry using anti-CD3 and anti-CD19 antibodies (e.g., Robinson et al., 1998). 2. Determine the saturating concentration of the anti-CD3 and anti-CD19 antibodies to be used by flow cytometry. Prepare a solution of each antibody in PBS/HSA at ten times (10×) the saturating concentration. Treat cells with antibodies 3. Resuspend the lymphocyte population (≤200 × 106 cells) in 9 ml PBS/HSA and add 4.5 ml of the suspension to each of two 15-ml conical centrifuge tubes. Add 0.5 ml of 10× anti-CD3 antibody to one tube and 0.5 ml of 10× anti-CD19 antibody to the other tube. 4. Incubate 45 min with gentle tilting and rotation on a platform rocker at 4°C. 5. Wash cells twice, each time by centrifuging 10 min at 600 × g (1500 rpm in H-1000 rotor), room temperature, removing the supernatant, resuspending in 10 ml PBS/HSA, and removing the supernatant. Finally, resuspend each tube in 5 ml PBS/HSA and put on ice for 15 min. Preparation and Isolation of Cells


Prepare magnetic beads 6. Based on the number of T and B cells estimated in step 1, determine the number of goat anti-mouse IgG–coated magnetic beads that will be required to purify the required number of T and B cells (5 to 10 beads will be needed for each specific target cell). 7. Based on the bead concentration supplied by the manufacturer, remove two aliquots of the required volume of goat anti-mouse IgG–coated beads from the source vial and place in 15-ml conical centrifuge tubes. Add PBS to the 14-ml mark and resuspend the beads. 8. Wash beads (see Basic Protocol 3, steps 2 to 4). Perform separation 9. Add 5 ml of anti-CD3-coated cells to one tube of washed anti-IgG-coated beads and 5 ml of anti-CD19-coated cell suspension to the other tube of anti-IgG-coated beads. 10. Perform magnetic separation to isolate T and B cells (see Basic Protocol 3, steps 8 to 17). ALTERNATE PROTOCOL 2

ISOLATION OF T AND B CELL SUBPOPULATIONS BY NEGATIVE SELECTION For certain experimental conditions, it is desirable to isolate lymphocyte subpopulations without antibody engagement of cell-surface molecules such as the immunoglobulin receptor on B cells and the CD3 complex on T cells. Therefore, negative selection procedures are followed that aim to maximize the elimination of all lymphocyte subpopulations except the desired one. For T cell isolation, the procedure involves elimination of B cells (CD19+), monocytes/macrophages (CD14+), and NK cells (CD16+); for B cell isolation T cells (CD3+), monocytes/macrophages, and NK cells are eliminated. The procedure is very similar to the above protocol for selection of T and B cells using anti-IgG–coated magnetic beads. Additional Materials (also see Basic Protocol 3) Anti-CD3, anti-CD14, anti-CD16, and anti-CD19 IgG monoclonal antibodies (Becton-Dickinson or Coulter) Goat anti-mouse IgG–coated magnetic beads (Dynabeads M-450; Dynal) Prepare cells and antibodies 1. Determine the approximate number of T cells, B cells, monocytes/macrophages, and NK cells in the starting population of lymphocytes by flow cytometry using anti-CD3, anti-CD19, anti-CD14, and anti-CD16 antibodies (e.g., Robinson et al., 1998). 2. Determine the saturating concentrations of the anti-CD3, anti-CD19, anti-CD14, and anti-CD16 antibodies to be used by flow cytometry. Prepare a solution of each antibody in PBS/HSA at ten times (10×) the saturating concentration. Treat cells with antibodies 3. Resuspend the lymphocyte population (≤ 200 × 106 cells) in 3.5 ml PBS/HSA in a 15-ml conical centrifuge tube.


4a. For T cell isolation: Add 0.5 ml each of 10× anti-CD19, 10× anti-CD14, and 10× anti-CD16 antibody preparations to the tube. 4b. For B cell isolation: Add 0.5 ml each of 10× anti-CD3, 10× anti-CD14, and 10× anti-CD16 antibody preparations to the tube.


5. Incubate 45 min with gentle tilting and rotation on a platform rocker at 4°C. 6. Wash cells twice, each time by centrifuging 10 min at 600 × g (1500 rpm in H-1000 rotor), room temperature, removing the supernatant, resuspending in 10 ml PBS/HSA, and removing the supernatant. Finally, resuspend cells in 5 ml PBS/HSA and put on ice for 15 min. Prepare magnetic beads 7. Based on the number of cells of each subpopulation estimated in step 1, determine the number of goat anti-mouse IgG–coated magnetic beads that will be required to bind the required number of antibody-coated lymphocytes (5 to 10 beads will be needed for each specific target cell). 8. Based on the bead concentration supplied by the manufacturer, remove the required volume of goat anti-mouse IgG–coated beads from the source vial and place in a 15-ml conical centrifuge tube. Add PBS to the 14-ml mark and resuspend the beads. 9. Wash beads (see Basic Protocol 3, steps 2 to 4). Perform separation 10. Add 5 ml of washed antibody-coated cells (from step 6) to the tube with the washed anti-IgG-coated beads. 11. Incubate 45 min with gentle tilting and rotation on a platform rocker at 4°C. 12. Place tube on vertical magnet for 2 min. Aspirate and save supernatant containing nonadherent cells, taking care not to disturb the beads clinging to the side of the tube. 13. Add 5 ml IMDM/HSA to tube. Gently resuspend beads. 14. Repeat steps 12 and 13 twice, saving and pooling nonadherent cells from each wash. 15. Centrifuge nonadherent cells 10 min at 600 × g (1500 rpm in H-1000 rotor), room temperature. Aspirate supernatant and resuspend cells in 5 ml IMDM/HSA. Count cells and determine number of viable cells by trypan blue exclusion (UNIT 1.1). 16. Determine purity of the T cell and B cell populations by flow cytometry using anti-CD3 and anti-CD19 antibodies (see, e.g., Robinson et al., 1998). REAGENTS AND SOLUTIONS Use deionized or distilled water in all recipes and protocol steps. For common stock solutions, see APPENDIX 2A; for suppliers, see SUPPLIERS APPENDIX.

Ficoll-Hypaque solution (density 1.07 to 1.08 g/ml) Dissolve 6.2 g Ficoll (mol. wt. 400,000; e.g., Sigma) in 75 ml distilled water with slow stirring. Add 10.4 g sodium diatrizoate and stir until solution is clear. Add water to 100 ml. Filter-sterilize using 0.22-µm filter (Nalgene). Store up to 6 months at 4°C in the dark. Alternatively, solutions can be purchased from commercial suppliers (e.g., Lymphocyte Separation Medium from Organon Teknika Cappel or equivalent product from Bio-Whittaker).

Freezing medium Mix 20 ml Iscove’s modified Dulbecco’s medium (IMDM; Life Technologies) and 20 ml heparinized human plasma. Cool to 4°C. Slowly add 10 ml dimethylsulfoxide, mixing after each incremental addition. Cool to 4°C and filter sterilize using a 0.22-µm filter.



Lymphocyte culture medium (LCM) Iscove’s modified Dulbecco’s medium (IMDM; Life Technologies) containing: 100 U/ml penicillin 100 µg/ml streptomycin 10% heparinized human plasma Store up to 2 weeks at 4°C COMMENTARY Background Information Density-gradient separation of lymphocytes Isolation of human lymphocytes from peripheral blood by density gradient centrifugation provides the following advantages: it is a simple technique, it is relatively inexpensive, it does not require special laboratory equipment, sterility is easily maintained, and high yields of lymphocytes are provided (Boyum, 1968). A typical yield of lymphocytes from whole blood of healthy adult donors is 1–2 × 106 cells/ml. Figure 2.2.2 shows a typical result from flow cytometric analysis of density gradient–purified lymphocytes in which 98% of cells are CD45+ (positive for common leukocyte antigen), 61% are CD3+ (T cells), and 21% are CD19+ (B cells). The non-T non-B cell population contains monocytes/macrophages and natural killer (NK) cells. Monocytes/macrophages can be easily purified from the lymphocyte population based on their differential adhesiveness to immobilized components of the the extracellular matrix contained in human serum. A typical yield of monocytes/macrophages from a lymphocyte population is 5% to 10%. Flow cytometric analysis shows that ≥95% of these cells are HLA class I+, HLA-DR+, and CD35+, and ≤1% are CD3+ T cells (Fig. 2.2.3). A subset of these monocytes/macrophages can be differentiated into immature antigen-presentationcompetent “dendritic-like” cells by culture in IL-4 and GM-CSF (Sallusto and Lanzavecchia, 1994). These “dendritic-like” cells have proven to be extremely useful antigen-presenting cells for the generation and cloning of rare self-reactive T cells (Biddison et al., 1997).


Magnetic-bead separation of lymphocytes Cell-separation procedures based on magnetic-bead technology have the advantages of easy manipulation, safety from contamination with microorganisms, and avoidance of high shear forces that affect cell viability. The most direct procedures involve positive selection with specific monoclonal antibodies directly

coupled to magnetic beads and the subsequent liberation of the bound cells by competing anti-Fab antibodies. This procedure also has the added advantage of providing a positively selected lymphocyte subpopulation free of any cell-bound selecting antibodies. A variation on this approach involves precoating the lymphocyte subpopulation of choice with a specific monoclonal IgG antibody and subsequent physical sequestration with anti-IgG-coated magnetic beads. In experimental situations in which specific antibody binding to cell-surface structures must be avoided, negative selection procedures are employed, wherein the lymphocyte subpopulations that are not desired are coated with specific IgG antibodies and are then removed by anti-IgG-coated magnetic beads. Each of these procedures has been used successfully to isolate human T cells, monocytes/macrophages, and B cells (Lea et al., 1986; Vartdal et al., 1987; Funderud et al., 1990). Alternative procedures for positive selection are cell sorting of fluoresceinated antibody– coated cells with a flow cytometer and panning of antibody-coated cells on plastic surfaces containing immobilized anti-immunoglobulin reagents. This author’s experiences with these techniques indicate that they present a much higher likelihood of contamination with microorganisms and subject the lymphocytes to conditions that can lead to more loss of viable cells than occurs with the magnetic-bead procedures. Cell sorting also has a major limitation in the number of cells that can be separated. Negative selection can also be provided by complement-fixing antibodies, but this approach involves the need to remove dead-cell debris before the remaining viable cells can be used for experimentation.

Critical Parameters and Troubleshooting To maximize the yield and purity of lymphocytes using the density gradient protocol (see Basic Protocol 1), it is essential to harvest all of the cells at the interface between the


B

Cell number

A

101

102

103

102

103

101

102

103

D

Cell number

C

101

101

102

103

Fluorescence intensity

Fluorescence intensity

Figure 2.2.3 Flow cytometric analysis of isolated monocytes/macrophages. The solid lines represent cells stained with (A) fluorochrome-labled anti-CD3 monoclonal antibodies, (B) fluorochrome-labeled anti-CD35 monoclonal antibodies, (C) fluorochrome-labeled anti–HLA class I monoclonal antibodies, and (D) fluorochrome-labeled anti-HLA-DR monoclonal antibodies. The dotted lines in panels B, C, and D represent cells stained with fluorochrome-labeled anti–mouse IgG alone.

Ficoll-Hypaque solution and the plasma/platelet layer. This may require harvesting up to 5 ml of the underlying Ficoll-Hypaque solution. This is why it is recommended that at least 7.5 ml of this solution be used as an underlay for each tube. Great care should be taken not to disturb or harvest any of the pelleted granulocytes and erythrocytes. One common problem is the accumulation of platelets at the FicollHypaque interface, which produces clumping and contamination with platelets and erythrocytes. This problem can be partially avoided by further dilution of blood with PBS and by not centrifuging the cells at temperatures 24 hr old.

Anticipated Results One umbilical vein should yield two confluent T-75 flasks in 3 to 5 days. By passage 2 there should be 32 confluent 150-mm dishes with ∼1 × 107 cells per dish. The yield from tissues is less than that from umbilical vein.

Time Considerations Preparation of the cells for all three methods should not take more than half a day.

Literature Cited Folkman, J., Haudenshild, C.C., and Zetter, B.R. 1979. Long term culture of capillary endothelial cells. Proc. Natl. Acad. Sci. U.S.A. 76:52175221. Fukuda, K., Imamura, Y., Koshihara, Y., Ooyama, T., Hanamure, Y., and Ohyama, M. 1989. Establishment of human mucosal microvascular endothelial cells from inferior turbinate culture. Am. J. Otolaryngol. 10:85-91. Gimbrone, M. 1976. Culture of vascular endothelium. Prog. Hemostasis Thromb. 3:1-28. Gordon, P.B., Sussman, I.I., and Hatcher, V.B. 1983. Long-term culture of human endothelial cells. In Vitro 19:661-671. Jaffe, E.A. 1980. Culture of human endothelial cells. Transplant. Proc. 12:49-53. Kibbey, M.C., Grant, D.S., and Kleinman, H.K. 1992. Role of the SIKVAV site of laminin in promotion of angiogenesis and tumor growth: An in vivo Matrigel model. J. Natl. Cancer Inst. 84:1633-1638. Kubota, Y., Kleinman, H.K., Martin, G.R., and Lawley, T.J. 1988. Role of laminin and basement membrane in morphological differentiation of human endothelial cells into capillary-like structures. J. Cell Biol. 107:1589-1598. Voest, E.E., Kenyon, B.M., O’Reilly, M.S., Truit, G., D’Amato, R.J., and Folkman, J. 1995. Inhibition of angiogenesis in vivo by interleukin 12. J. Natl. Cancer Inst. 87:581-586.

Contributed by Hynda K. Kleinman National Institute of Dental Research/NIH Bethesda, Maryland Maria C. Cid Hospital Clinic i Provincia Barcelona, Spain

Preparation of Endothelial Cells


Generation of Continuously Growing B Cell Lines by Epstein-Barr Virus Transformation Epstein-Barr virus (EBV) has been widely used to transform human B cells in vitro and to produce continuously growing B cell lines for use in a variety of in vitro studies. Most human B cell lines that are transformed by EBV secrete little or no infectious viral particles. For this reason, a marmoset line transformed with the human Hawley strain of EBV, which secretes active infectious EBV into the culture supernatant, is used as a source of transforming virus (Miller and Lipman, 1973). This protocol explains how to prepare the marmoset-derived EBV and transform human B cells obtained from peripheral blood.

UNIT 2.4

BASIC PROTOCOL

CAUTION: EBV is a known human pathogen. Appropriate biosafety practices must be followed. Materials Complete culture medium (see recipe), 37°C B95-8 EBV-transformed marmoset cell line (ATCC #CRL 1612) Anti-CD3 monoclonal antibody produced by OKT3 hybridoma (ATCC #CRL 8001) 25-cm2 and 75-mm2 tissue culture flasks Sorvall RT-6000B centrifuge with H-1000 rotor (or equivalent refrigerated centrifuge and rotor) and 50-ml centrifuge tubes 0.45-µm sterile filter Additional reagents and equipment for growing cells, determining cell viability by trypan blue exclusion, and cryopreservation of cells (UNIT 1.1) and preparation of peripheral blood lymphocytes (UNIT 2.3) NOTE: All solutions and equipment coming into contact with cells must be sterile, and proper aseptic technique should be used accordingly. NOTE: All culture incubations should be performed in a humidified 37°C, 5% CO2 incubator unless otherwise specified. Some media (e.g., DMEM) may require altered levels of CO2 to maintain pH 7.4. 1. Resuspend B95-8 cells in complete culture medium at 1 × 106 cells/ml and incubate in 75-mm2 tissue culture flasks at 50 ml of culture per flask for 3 days (until ≥95% viable and in exponential growth phase; see UNIT 1.1 for basic culture technique and determination of cell viability). 2. Transfer cultures to 50-ml centrifuge tubes. Centrifuge 10 min at 600 × g (1500 rpm in H-1000 rotor), 4°C. Filter supernatant through 0.45-µm sterile filter, divide into 0.6-ml aliquots, and store at −70°C. The culture supernatants should contain 102 to 103 infectious units/ml (Miller and Lipman, 1973). Determination of EBV titers can be done by quantitative assessment of transformation of umbilical cord leukocytes (Miller and Lipman, 1973); however, because of the difficulty in obtaining such cells, this determination is usually omitted.

3. Prepare peripheral blood lymphocytes as described in UNIT 2.3. Resuspend lymphocytes in 37°C complete culture medium at 1 × 106 cells/ml, then place 5 ml of the lymphocyte suspension in an upright 25-cm2 tissue culture flask. 4. Add anti-CD3 antibody to final concentration of 10 µg/ml. Incubate cells and anti-CD3 antibody for 1 hr with the flask in the upright position. Contributed by William E. Biddison Current Protocols in Cell Biology (1999) 2.4.1-2.4.3 Copyright © 1999 by John Wiley & Sons, Inc.


2.4.1 Supplement 1

5. Add 0.5 ml of the EBV-containing B95-8 supernatant (from step 2) to flask. Incubate with the flasks in the upright position for 1 to 2 weeks (until medium begins to turn orange/yellow and small clumps of cells become visible). 6. Add 5 ml of fresh 37°C complete medium, then incubate 2 to 3 days. After that period, remove 5 ml of the supernatant and add 5 ml of fresh 37°C complete medium and continue incubating. Repeat the feedings as described in this step until total cell number exceeds 5 × 106. 7. Transfer growing cells to 75-cm2 flask in 50 ml of 37°C complete medium and incubate until cell concentration is ≥1 × 106/ml. 8. Cryopreserve aliquots of cells (UNIT 1.1) and maintain B cell line by splitting to 1 × 105 cells/ml in complete medium, incubating to 1 × 106 cells/ml, then splitting again. The cells can be maintained this way indefinitely in the absence of any contamination with microorganisms.


Complete culture medium RPMI 1640 medium containing: 5% fetal bovine serum (FBS, heat-inactivated; APPENDIX 2A) 100 U/ml penicillin 100 µg/ml streptomycin 100 µg/ml gentamycin 2 mM L-glutamine 10 mM HEPES Store up to 2 weeks at 4°C COMMENTARY Background Information

EBV Transformation of B Cells

EBV is able to transform a subset of human resting B cells from peripheral blood (Sugden and Mark, 1977; Aman et al., 1984). The outgrowth of EBV-transformed B cells is prevented by the presence of EBV-immune T cells contained within the peripheral blood lymphocyte population (Rickinson et al., 1979). For this reason, soluble anti-CD3 antibody is included in the transformation process to inhibit the ability of T cells to respond to EBV antigens presented by the transformed B cells. Other procedures to eliminate T cell reactivity that could be utilized include T cell depletion of the lymphocyte population (UNIT 2.3) and functional T cell inactivation by cyclosporin A (Tosato et al., 1982). EBV-transformed B cell lines have proven useful in studies of the cell biology of antigen presentation (Roche and Cresswell, 1990) and in the production of large-scale protein preparations for characterization of MHC antigens

(Orr et al., 1979) and MHC-bound peptides (Falk et al., 1991).

Critical Parameters EBV-transformed B cells will not survive if there is an effective anti-EBV T cell response in the culture. It is essential that T cell functions be blocked with an adequate concentration of anti-CD3 antibody (10 µg/ml is sufficient). Also the number and viability of resting B cells in the culture must be sufficient to permit an efficient transformation by EBV (at least 5 × 106 peripheral blood lymphocytes are recommended). In addition, the titer of infectious EBV in the B95-8 culture fluid must be adequate for transformation. If transformation fails to occur, it will be necessary to obtain umbilical cord leukocytes to measure EBV infectivity (Miller and Lipman, 1973). Different batches of FBS have variable capacities for promoting the growth of EBV-transformed B cells. It is best to predetermine whether the available FBS can support the growth of existing EBV-trans-

2.4.2 Supplement 1


formed B cell lines before using it for establishing new EBV-transformed lines. Great care should be taken to avoid contamination with mycoplasma (also see UNIT 1.5). Mycoplasmacontaining EBV-transformed B cell lines will lose their capacity for continuous growth.

Anticipated Results With the procedure described above, EBVtransformed B cell lines should be obtained which can maintain continuous growth indefinitely in the absence of contamination by microorganisms.

Time Considerations Preparation of B95-8 culture supernatants requires a 3-day incubation. The establishment of the cultures for transformation takes 1 to 2 hr once the peripheral blood lymphocyte population has been obtained. Culture for 2 to 3 weeks is required to produce continuously growing EBV-transformed B cell lines.

set leukocytes. Proc. Natl. Acad. Sci. U.S.A. 70:190-194. Orr, H., Lopez de Castro, J., Parham, P., Ploegh, H., and Strominger, J. 1979. Comparison of amino acid sequences of two human histocompatibility antigens: HLA-A2 and HLA-B7: Location of putative alloantigenic sites. Proc. Natl. Acad. Sci. U.S.A. 76:4395-4399. Rickinson, A., Moss, D., and Pope, J. 1979. Longterm T cell–mediated immunity to Epstein-Barr virus in man. II. Components necessary for regression in virus-infected leukocyte cultures. Int. J. Cancer 23:610-617. Roche, P. and Cresswell, P. 1990. Invariant chain association with HLA-DR molecules inhibits immunogenic peptide binding. Nature 345:615618. Sugden, B. and Mark, W. 1977. Clonal transformation of adult human leukocytes by Epstein-Barr virus. J. Virol. 23:503-508. Tosato, G., Pike, S., Koski, I., and Blaese, R. 1982. Selective inhibition of immunoregulatory cell functions by Cyclosporin A. J. Immunol. 128:1986-1991.

Literature Cited

Key Reference

Aman, P., Ehlin-Henriksson, B., and Klein, G. 1984. Epstein-Barr virus susceptibility of normal human B lymphocyte populations. J. Exp. Med. 159:208-220.

Presents detailed description of B cell transformation and determination of EBV titers using umbilical cord leukocytes.

Falk, K., Rotzschke, O., Stevanovic, S., Jung, G., and Rammensee, H.-G. 1991. Allele-specific motifs revealed by sequencing of self-peptides eluted from MHC molecules. Nature 351:290293. Miller, G. and Lipman, M. 1973. Release of infectious Epstein-Barr virus by transformed marmo-

Miller and Lipman, 1973. See above.

Contributed by William E. Biddison National Institute of Neurological Disorders and Stroke, NIH Bethesda, Maryland



Supplement 1

Laser Capture Microdissection

UNIT 2.5

This unit describes a method for isolating pure populations of cells for biochemical and molecular analysis, called laser capture microdissection (LCM; see Basic Protocol). A second protocol outlines procedures for staining frozen or paraffin-embedded tissue samples for LCM (see Support Protocol). ISOLATION OF A PURE CELL POPULATION FROM TISSUE SECTIONS Laser capture microdissection (LCM) provides the scientific community with a rapid and reliable method for obtaining pure cell populations from tissue sections under direct microscopic visualization. It incorporates both an inverted light microscope (with or without a fluorescent module) and a near-infrared laser to facilitate the visualization and procurement of cells. Briefly, a stained slide is placed under a microscope, and a specific adherence cap with an ethylene vinyl acetate (EVA) film is placed onto the tissue. The user moves the slide until the area of interest lies in the center of the microscope’s field of view. When the cells of interest are located, a near-infrared laser is fired, which melts the EVA film in the targeted area. The EVA film expands into the void of the stained tissue and solidifies within 200 msec as it rapidly cools. The targeted tissue bonds to the EVA film, retaining the exact cellular morphology, DNA, RNA, and proteins intact. Both frozen and fixed tissue samples are successfully microdissected using LCM. Recovered cells can be analyzed for DNA, RNA, and protein content, and used to construct cell-specific cDNA libraries (Emmert-Buck et al., 1996; Krizman et al., 1996; Simone et al., 1998, 2000; Banks et al., 1999). LCM offers a quick means of procuring pure cell populations; however, it is necessary to follow strict protocols pertaining to fixation, preparation, and handling of tissue samples to be microdissected.

BASIC PROTOCOL

Materials Stained tissue samples, either frozen or formalin paraffin-embedded, cut into 2- to 10-µm sections, and mounted on plain, uncharged microscope slides PixCell II Laser Capture Microdissection System (Arcturus Engineering) CapSure transfer film (Arcturus Engineering) Compressed gas duster CapSure pads (Arcturus Engineering) Cap removal tool (Arcturus Engineering) Setup LCM apparatus 1. Load the CapSure cassette module with a CapSure cartridge. 2. Move joystick into vertical position to properly position the cap in relation to the capture zone. Place the microscope slide containing the prepared and stained tissue sample on microscope stage. When the target zone for microdissection appears in the viewing area, turn on Vacuum on front of controller to hold slide in place. 3. Slide CapSure cassette backward or forward to place cap at Load position. Slide transport arm over CapSure cap, then lift transfer film transport arm and place cap onto slide. 4. To enable the PixCell II laser, turn the keyswitch located on front of the controller and press the Laser Enable button. Figure 2.5.1 shows components of the PixCell II. Preparation and Isolation of Cells Contributed by Lu Charboneau, Cloud P. Paweletz, and Lance A. Liotta Current Protocols in Cell Biology (2001) 2.5.1-2.5.7 Copyright © 2001 by John Wiley & Sons, Inc.

2.5.1 Supplement 10

NIH Laser captu re microdissection

transpor t arm

glass slide

plastic cap laser beam

tissue section

transfer film individual on backing cell sample

cell(s) of interest

joystick

plastic cap

transport arm

transfer of selected cell(s)

glass slide

Figure 2.5.1 Schematic overview of the Laser Capture Microdissection Microscope. Through activation of a laser beam and under direct microscopic visualization, individual cells can be isolated for use in molecular analysis of any kind.

5. Verify that target beam is focused by selecting a spot size of 7.5 µm, using the Spot Size Adjust lever found on the left side of the microscope. Rotate microscope objectives until the 10× objective is in place. Reduce light intensity through the optics until field seen on the monitor appears almost dark and target beam is easily viewed. Using the Fine Focus Adjust located below the Spot Size Adjust lever, adjust target beam until it reaches the point of sharpest intensity and most concentrated light with little or no halo effect (Fig. 2.5.2). The laser is now focused for any of the three laser spot sizes.

6. Select the laser spot size suitable to perform the microdissection. 7. Adjust Power and Duration of the laser pulse on front of the controller to vary the diameter of the “capture zone”, using target settings as a reference point. Adjust these settings up or down to customize the laser to the type and thickness of the tissue to be dissected. Suggested settings are listed in Table 2.5.1. Laser Capture Microdissection

2.5.2 Supplement 10


E

HIJKLMN

F

KOHIJKLMN

G

KOHIJKLMN

Figure 2.5.2 Focusing of the target beam. If transfer of cells to the cap is not adequate, improper focus may be the cause. Select a spot size of 7.5 µm using the Spot Size Adjust lever found on the left side of the microscope. Rotate the objectives of the microscope until the 10× objective is in place. Reduce the intensity of the light through the optics until the field viewed on the monitor is almost dark and the target beam is easily viewed. Compare the target beam with this figure. If the target is not focused, refocus the target beam as described (see Basic Protocol, step 2).



Supplement 10

Table 2.5.1

Settings for LCM

Spot size

Power

Duration

7.5 µm 15 µm 30 µm

25 mW 30 mW 30 mW

3.0 msec 5.0 msec 8.0 msec

Perform LCM 8. Microdissect cells of interest. Guide microdissection with target beam and press Pendant switch for a single microdissection. To microdissect multiple shots, hold Pendant switch down. The tissue sample slide must contain no residual xylene, because it melts the transfer film, making microdissection impossible. Dry slide with a compressed gas duster before microdissection to remove residual liquid from staining procedure (see Support Protocol). To adjust the frequency interval, select Repeat on controller, then choose the desired time between laser pulses.

9. Observe wetting as the laser fires, maintaining a distinct clear circle surrounded by a dark ring (Fig. 2.5.3). If proper wetting is not observed, refocus the target beam (step 5). Collect microdissected cells 10. After collecting the desired number of cells, lift cap from slide using the transport arm. Lift and rotate transport arm until cap reaches the Cap Removal Site. Lower transport arm, then rotate it back toward slide, leaving the cap in place at the removal site. Remove cap using the provided cap tool. 11. Blot polymer surface of CapSure transfer film with CapSure pads to remove nonspecific debris adhered to it. 12. Inspect slide and cap for successful microdissection (Fig. 2.5.3). 13. Insert polymer end of cap into top of a 500-µl microcentrifuge tube. The sample is ready for extraction of desired components or can be frozen at −80°C for later analysis. SUPPORT PROTOCOL

HEMATOXYLIN AND EOSIN STAINING OF TISSUES FOR LCM A reaction sequence for performing basic hematoxylin and eosin staining is outlined in Table 2.5.2. Although LCM is not limited to hematoxylin and eosin staining, it is probably the most versatile stain used. Staining can be done using Coplin jars or racks and dishes. DNA can be retrieved from frozen as well as paraffin-embedded tissue that has been fixed in formalin or ethanol. RNA and protein retrieval is best when frozen tissue is used or alternatively when ethanol-fixed paraffin-embedded tissue is used. RNA and protein are cross-linked when fixed in formalin. However, the morphology of formalin- or ethanolfixed paraffin-embedded tissues is almost always better than that of frozen tissues, and this is often the deciding factor as to what tissue fixation to use.

Laser Capture Microdissection

2.5.4 Supplement 10


PMHIQMRNSLLMJTSIO

UHTMQRNSLLMJTSIO

JUVTKQMNRJMWWL

Figure 2.5.3 LCM capture of complex tissue structures. Successful transfer is achieved when nonspecific cells are absent on cap and when microdissection does not leave residual cells behind (see Critical Parameters and Troubleshooting). Example of good microdissection: (A) heterogeneous prostate tissue, (B) removal of normal epithelium, and (C) transfer to EVA film in 25% of the plasma membrane markers being found in the pelleted fraction and by the apical plasma membrane markers having more basolateral-like distributions. This problem can be corrected by increasing the number of sonication bursts until no Y-shaped membrane structures are visible by phase-contrast microscopy. Oversonication is a less prevalent problem, but results in basolateral markers having a more apical-like distribution in the gradient. This problem can be avoided by more closely monitoring

vesiculation (e.g., after every sonication burst) by phase-contrast microscopy. The assays to determine the recovery and purity of the plasma membrane sheets and of the two domains are relatively straightforward and easily interpreted. The most common problem is determining the appropriate dilution of the preparative fractions such that their activity levels fall within the values of the standard curve. In Table 3.2.2, suggested dilutions for preparative fractions are listed for both the enzyme assays and protein concentration determination using BCA reagent (Pierce). Alternatively, different standard curves can be prepared shifting the concentrations either higher or lower, or different incubation times can be used. In general, dilution of the domain gradient fractions is not suggested when assaying enzyme activities. Thus, altering the standard curve concentrations and incubation times are advised if problems are encountered.

Anticipated Results

The plasma membrane sheets purified according to Basic Protocol 1 are enriched 20- to 40-fold in plasma membrane markers. This preparation contains substantial amounts of both domains in continuity with each other in ratios approaching those of intact hepatocytes. The yield is 10% to 20% of total plasma membranes. The protein concentration of the purified sheets generally ranges from 1 to 2 mg/ml, corresponding to ∼1 mg of plasma membrane protein/g of starting liver wet weight. The major contaminant of the isolated sheets is endoplasmic reticulum, which is enriched in these fractions 1-fold (see Table 3.2.1). The vesicles derived from the apical and basolateral domains are partially resolved on the basis of differences in equilibrium density (see Fig. 3.2.1). The density profile for apical vesicles is characterized by a single peak with its center at a density of 1.10 g/cm3 (refractive index = 1.3713). Basolateral vesicles have a bimodal distribution, with a peak centered at 1.14 g/cm3 (refractive index = 1.3859) and a smaller (and variable) amount found in the pelleted fraction. The apical and basolateral plasma membrane antigens, as well as tight junction components, exhibit distinct staining patterns in plasma membrane sheets. As shown in Fig. 3.2.2, the relatively intense staining for HA4 (an apical protein) is restricted to the bile canalicular membranes in an evenly distributed pattern. In contrast, staining for HA321 (a basolateral protein) is excluded from the canalicular structures and is detected in the surrounding membranes as a more diffuse and less intense signal. The tight junction protein ZO-1 is detected in regions immediately adjacent

Subcellular Fractionation and Isolation of Organelles


Supplement 2

Table 3.2.2 Suggested Dilutions for Analysis of Plasma Membrane Preparative Fractions

Fraction

Dilutions for enzyme assaysa

Dilutions for BCA protein assay

H S1 P1 S2 P2 I 1.18 g/m3 P3 S3 PM

1:50; 1:100 1:25; 1:50 1:50; 1:100 1:20, 1:40 1:10; 1:20 1:10; 1:20 1:2; 1:4 1:5; 1:10 1:2; 1:4 1:50; 1:100

1:50; 1:100 1:25; 1:50 1:20; 1:40 1:25; 1:50 1:5; 1:10 No dilution; 1:2 1:2; 1:4 1:50; 1:100 No dilution 1:5; 1:10

aIn these listings, “1:50” indicates 1 part enzyme in a total of 50 parts (i.e., 1 part enzyme plus 49 parts diluent).

to (outside) the apical plasma membrane in a ring-like pattern, indicating the location of the junctional complexes that form the barrier between plasma membrane domains.

Time Considerations

The isolation of plasma membrane sheets starting from the excision of the rat liver to the final plasma membrane pellet takes only 4 to 5 hr. Preparation of the sheets for density centrifugation by sonication generally takes 15 to 30 min. Each of the enzyme assays takes approximately 3 to 4 hr to perform, including setting up the assay, incubating the samples, reading the absorbances, and performing the calculations. Processing the sheets for indirect immunofluorescence is accomplished within 2 to 3 hr. SDS-PAGE also takes 2 to 3 hr and immunoblotting takes 18 to 24 hr.

Literature Cited

Bartles, J.R., Braiterman, L.T., and Hubbard, A.L. 1985. Endogenous and exogenous domain markers of the rat hepatocyte plasma membrane. J. Cell Biol. 100:1126-1138. Blouin, A., Bolender, R.P., and Weibel, E.R. 1977. Distribution of organelles and membranes between hepatocytes and nonhepatocytes in the rat liver parenchyma. J. Cell Biol. 72:441-455. Fujita, H., Tuma, P.L., Finnegan, C.M., Locco, L., and Hubbard, A.L. 1998. Endogenous syntaxins 2, 3 and 4 exhibit distinct but overlapping patterns of expression at the hepatocyte plasma membrane. Biochem. J. 329:527-538.

Isolation of Rat Hepatocyte Plasma Membranes

Hubbard, A.L., Wall, D.A., and Ma, A. 1983. Isolation of rat hepatocyte plasma membranes. I. Presence of the three major domains. J. Cell Biol. 96:217-229. Hubbard, A.L., Bartles, J.R., and Braiterman, L.T. 1985. Identification of rat hepatocyte plasma membrane proteins using monoclonal antibodies. J. Cell Biol. 100:1115-1125. Stieger, B., Marxer, A., and Hauri, H.-P. 1986. Isolation of brushborder membranes from rat and rabbit colonocytes: Is alkaline phosphatase a marker enzyme? J. Membr. Biol. 2:19-31. Touster, O., Aronson, N.N., Dulaney, J.T., and Hendrickson, H. 1970. Isolation of rat liver plasma membranes: Use of nucleotide pyrophosphatase and phosphodiesterase I as marker enzymes. J. Cell Biol. 47:604-618. Weibel, E.R. 1976. Stereological approach to the study of cell surface morphometry. Sixth European Congress on Electron Microscopy, Jerusalem, pp. 6-9. Weibel, E.R., Satubli, W., Gnagi, H.R., and Hess, F.A. 1969. Correlated morphometric and biochemical studies of the liver cell. J. Cell Biol. 42:68-91. Widnell, C.C. and Unkeless, J.C. 1968. Partial purification of a lipoprotein with 5′ nucleotidase activity from membranes of rat liver cells. Proc. Natl. Acad. Sci. U.S.A. 61:1050-1057.

Contributed by Pamela L. Tuma and Ann L. Hubbard Johns Hopkins University School of Medicine Baltimore, Maryland

Hubbard, A.L. and Ma., A. 1983. Isolation of rat hepatocyte plasma membranes. II. Identification of membrane-associated cytoskeletal proteins. J.Cell Biol. 96:230-239.

3.2.16 Supplement 2


Isolation of Mitochondria from Tissues and Cells by Differential Centrifugation

UNIT 3.3

The protocols in this unit are simple and rapid methods for the isolation of a mitochondrial fraction from three different mammalian tissues (liver, heart, and skeletal muscle), from cultured cells, and from yeast. Unlike the protocols in UNIT 3.4, they only require routine differential centrifugation in low- and high-speed centrifuges and should be accessible to any laboratory. These mitochondrial fractions will be contaminated to varying degrees by smaller particles (lysosomes and peroxisomes), although the heavy mitochondrial fraction from rat liver is relatively pure (∼90%). These preparations can be used as starting material for the density-gradient separations described in UNIT 3.4. Basic Protocol 1 describes the isolation of the heavy mitochondrial fraction from rat liver; this fraction has high respiratory control and can be used in oxygen electrode studies for ≥4 hr after preparation. Centrifuging the postnuclear supernatant at only 3000 × g avoids significant contamination of the pellet by other more slowly sedimenting organelles (e.g., lysosomes, peroxisomes, endoplasmic reticulum). Contamination is reduced further by gently washing the 3000 × g pellet. A mannitol-containing buffer is the medium of choice for this preparation. Other protocols describe methods for obtaining crude total mitochondrial fractions from bovine heart (see Basic Protocol 2), rat skeletal muscle (see Basic Protocol 3), cultured cells (see Basic Protocol 4), and yeast (see Basic Protocol 5). The major differences between these protocols is their mode of homogenization. Although the easy availability of rat liver in most laboratories makes it a popular choice as a source of mitochondria, those from bovine heart are also often used for respiratory studies. Indeed, they may even be more tightly coupled than liver mitochondria, and they can generally be stored for longer periods while maintaining good functional integrity. This may be allied to the lower levels of proteases and slower release of fatty acids in this tissue. Heart mitochondria also tend to provide better yields of the various structural components of electron transport and ATP synthesis. Basic Protocol 2 also provides a strategy for large-scale preparation. The increasing use of yeast as a model for mammalian membrane and organelle synthesis points to the importance of Basic Protocol 5. Although rat brain is another widely used source of mitochondria, they are rarely purified by differential centrifugation alone (see UNIT 3.4). Methods for measuring succinate dehydrogenase, catalase, and β-galactosidase (as mitochondrial, peroxisomal, and lysosomal markers, respectively) in density-gradient fractions are given in UNIT 3.4, but they can also be applied to assessing the purity of mitochondria prepared by differential centrifugation. Protease inhibitors (see UNIT 3.4, Reagents and Solutions) can be included in any or all of the media at the discretion of the investigator, except in the protease-containing solutions used in Basic Protocols 3 and 5. NOTE: For all protocols, all g values are given as g. PREPARATION OF THE HEAVY MITOCHONDRIAL FRACTION FROM RAT LIVER Young adult male animals (150 to 200 g) are routinely used, providing livers of ∼10 g wet weight. This protocol is designed for one such liver, but can be scaled up or down proportionally for different amounts of liver. The animals are normally deprived of food overnight to reduce the glycogen content of the liver; this facilitates the separation process. Contributed by John M. Graham Current Protocols in Cell Biology (1999) 3.3.1-3.3.15 Copyright © 1999 by John Wiley & Sons, Inc.

BASIC PROTOCOL 1


3.3.1 Supplement 4

A homogenization medium containing mannitol and sucrose, a chelating agent (either EGTA or EDTA), and a buffer (normally HEPES or MOPS) is best suited to respiratory studies. NOTE: All protocols using live animals must first be reviewed and approved by an Institutional Animal Care and Use Committee (IACUC) or must conform to governmental regulations regarding the care and use of laboratory animals. NOTE: All solutions, glassware, centrifuge tubes, and equipment should be precooled to 0° to 4°C and kept on ice throughout. When handling the glass vessel of the PotterElvehjem homogenizer, a thermally insulated glove or silicone rubber hand protector should be used, not only to avoid heat transfer from the skin, but also to protect the hand in the unlikely event that the vessel breaks. Materials 150- to 200-g male Sprague-Dawley rat Liver homogenization medium (LHM; see recipe), ice cold Dissecting tools Potter-Elvehjem homogenizer (∼0.09-mm clearance; 25-ml working volume) Overhead high-torque electric motor (thyristor-controlled) Low-speed centrifuge with swinging bucket rotor and appropriate tubes High-speed centrifuge with fixed-angle rotor and 40- to 50-ml polycarbonate tubes Vacuum pump Dounce homogenizer (30- to 40-ml volume) with loose-fitting pestle (Wheaton type B) Isolate liver 1. Deprive a 150- to 200-g male Sprague-Dawley rat of food overnight. 2. Sacrifice the animal by cervical dislocation or decapitation. This must be supervised or carried out by an experienced animal technician.

3. Open the abdominal cavity and transfer the liver to a chilled beaker containing ∼20 ml LHM. 4. Decant the medium and finely mince the liver using scissors. The pieces of liver should be no more than ∼25 mm3.

5. Agitate the minced tissue in ∼30 ml ice-cold LHM and allow the pieces to settle out. 6. Decant the medium and replace with ∼40 ml fresh medium. Homogenize liver 7. Transfer half the suspension to the chilled glass vessel of a Potter-Elvehjem homogenizer. 8. Attach the cold pestle to an overhead high-torque electric motor and homogenize the minced liver using five to six up-and-down strokes of the pestle, rotating at ∼500 rpm. Decant the homogenate into a beaker on ice. The motor should be securely mounted either to a wall, to a bench via a G clamp, or in a floor-standing cradle. Attachment to a free-standing retort stand is not adequate. Isolation of Mitochondria by Differential Centrifugation

9. Rinse the homogenizer with medium and wipe the pestle to remove any adhering connective tissue. Repeat the procedure with the other half of the suspension.

3.3.2 Supplement 4


Isolate mitochondria 10. Centrifuge the homogenate 10 min at 1000 × g, 4°C, in a swinging-bucket rotor using a low-speed centrifuge. 11. Aspirate the supernatant and transfer to 40- to 50-ml polycarbonate tubes. It is convenient to use a 20- to 30-ml syringe attached to a metal filling cannula (i.d. 0.8 to 1.0 mm) to aspirate supernatants that are to be recentrifuged.

12. Centrifuge the supernatant 10 min at 3000 × g, 4°C, in a fixed-angle rotor using a high-speed centrifuge. 13. Using a glass Pasteur pipet attached to some form of vacuum pump, aspirate the supernatant from each tube, keeping the tip of the pipet at the meniscus to remove as much of the floating lipid layer as possible. Also remove as much as possible of the loose-packed pinkish layer that overlies the brown mitochondria. 14. Wipe away any remaining lipid adhering to the wall of the tube with a paper tissue. Removal of this lipid is essential, as free fatty acids are potent uncouplers of phosphorylation from electron transport.

15. Add a small amount of LHM (∼8 ml) to each pellet and crudely resuspend the pellet with a glass rod. Then resuspend fully using 3 to 4 very gentle strokes in a Dounce homogenizer. 16. Make up to the original volume with LHM, transfer to new tubes, and recentrifuge 10 min at 3000 × g in the high-speed centrifuge. 17. Repeat steps 13 to 16 twice more. For large-scale preparations, the total volume of LHM used to resuspend the pellet can be reduced by ∼50% for the second and third washes.

18. Resuspend the purified mitochondria in a buffer whose composition is compatible with any subsequent analysis or processing; in many instances, LHM will be satisfactory. See Time Considerations for information about storage of mitochondria prior to further processing.

LARGE-SCALE PREPARATION OF MITOCHONDRIA FROM BOVINE HEART

BASIC PROTOCOL 2

It is important that fresh slaughterhouse material be used for this preparation, and that any adhering connective and adipose tissue be carefully removed. Because of the scale of the preparation and the size of the homogenization equipment, the procedure must be carried out in a cold room. The procedure is suitable for 500 to 600 g of material, and is adapted from Smith (1967) and Rice and Lindsay (1997). NOTE: All solutions, glassware, centrifuge tubes, and equipment should be precooled to 0° to 4°C and kept on ice or in a cold room throughout. Materials Bovine heart, freshly isolated Heart wash buffer (see recipe) 2.0 M Tris base Sucrose/succinate solution (SS; see recipe)



Supplement 4

Commercial mincer with a total capacity of 2 to 3 liters Waring blender or other large-capacity rotating blades homogenizer Cotton muslin Low-speed centrifuge with swinging-bucket rotor and 250- to 750-ml bottles High-speed centrifuge with fixed-angle rotor Glass rod Dounce homogenizer (50-ml volume) with loose-fitting pestle (Wheaton type B) Prepare the heart 1. Cut freshly isolated bovine heart tissue into small cubes (∼4 cm3) and pass once through a commercial mincer. 2. Suspend in 800 ml ice-cold heart wash buffer. While stirring, adjust the pH to 7.8 by adding 2.0 M Tris base. 3. Pour through two layers of cotton muslin and then squeeze to remove as much of the liquid as possible. 4. Transfer minced tissue to a clean beaker and suspend in 800 ml ice-cold SS. Homogenize the heart 5. Transfer half the suspension to a Waring blender and homogenize at high speed for 20 sec. 6. Readjust the pH to 7.8 using 2.0 M Tris base and then reblend for 60 sec. 7. Repeat the procedure with the other half of the suspension. Combine the two homogenates and dilute with ice-cold SS to ∼2.2 liters. Isolate mitochondria 8. Transfer homogenate to 250- to 750-ml centrifuge bottles and centrifuge 20 min at 800 × g, 4°C, in a swinging-bucket rotor using a low-speed centrifuge. 9. Carefully decant the supernatants and recentrifuge 20 min at 26,000 × g, 4°C, in a fixed-angle rotor using a high-speed centrifuge. A rotor such as the Sorvall SLA-1500 will allow this to be carried out in two centrifugations.

10. Decant and discard the supernatant. The pellet is clearly tripartite.

11. Tilting the bottles, gently pour ∼10 ml SS on top of each pellet and gently swirl the contents to resuspend the top light-brown layer of partially disrupted mitochondria. Discard this material. 12. Crudely resuspend the remaining dark-brown mitochondria in ∼20 ml SS using a glass rod, avoiding the almost-black hard-packed button at the bottom. 13. Completely resuspend mitochondria using 2 to 3 gentle strokes in a Dounce homogenizer. 14. Dilute the suspension to ∼300 ml with SS and recentrifuge 20 min at 26,000 × g, 4°C. 15. Collect and resuspend the middle layer of the pellet as in steps 10 to 13. The upper and lower layers of this second pellet should be relatively minor components.

16. If the composition of the SS medium is incompatible with any subsequent analysis, centrifuge the suspension 20 min at 26,000 × g and resuspend the pellet in a suitable medium. Isolation of Mitochondria by Differential Centrifugation

See Time Considerations for information about storage of mitochondria prior to further processing.

3.3.4 Supplement 4


PREPARATION OF MITOCHONDRIA FROM SKELETAL MUSCLE This protocol uses a commercially available protease to facilitate the homogenization of the muscle tissue. Using this approach, the severity of the shear forces used to disrupt the tissue can be reduced, thus minimizing any damage to the mitochondria. The protocol is adapted from Bhattacharya et al. (1991). The protocol is designed for 4 to 5 g muscle tissue, but can be scaled up or down proportionally.

BASIC PROTOCOL 3

NOTE: All protocols using live animals must first be reviewed and approved by an Institutional Animal Care and Use Committee (IACUC) or must conform to governmental regulations regarding the care and use of laboratory animals. NOTE: All solutions, glassware, centrifuge tubes, and equipment should be precooled to 0° to 4°C and kept on ice or in a cold room throughout. When handling the glass vessel of the Potter-Elvehjem homogenizer, a thermally insulated glove or silicone rubber hand protector should be used, not only to avoid heat transfer from the skin, but also to protect the hand in the unlikely event that the vessel breaks. Materials 150- to 200-g male Sprague-Dawley rat Muscle wash buffer (see recipe) Muscle homogenization medium I (see recipe) Muscle homogenization medium II (see recipe) Dissecting tools Potter-Elvehjem homogenizer (∼0.3-mm clearance, 40-ml working volume) Overhead high-torque electric motor (thyristor-controlled) Fine nylon mesh (200-µm pore size) Low-speed centrifuge with swinging-bucket rotor and appropriate tubes High-speed centrifuge with fixed-angle rotor Dounce homogenizer (30-ml volume) with loose-fitting pestle (Wheaton type B) Isolate muscle tissue 1. Sacrifice a 150- to 200-g male Sprague-Dawley rat by cervical dislocation or decapitation. This must be supervised or carried out by an experienced animal technician.

2. Rapidly dissect out 4 to 5 g of striated leg muscle and wash it twice in ∼50 ml muscle wash buffer. 3. Mince the muscle finely on a cold surface (e.g., a glass plate on crushed ice) using two scalpels. The muscle pieces should be 15 amino acids from the upstream hydrophobic segment. However, since the hydrophilic domain located between hydrophobic segments 1 and 2 consists of only 4 amino acids (EQFG), it is not possible to place the fusion joint more than 15 amino acids from hydrophobic segment 1 without capturing most of hydrophobic segment 2 in fusion b. Therefore, the fusion joint is positioned such that only 5 amino acids of hydrophobic segment 2 are present in fusion b. In placing the fusion joint at this position, the charged residues immediately following hydrophobic segment 1 are left undisturbed. Fusion a lacks a putative transmembrane segment and will remain cytosolic regardless of the topology of the N terminus of intact SPC12 protein. One exception to this is that, if the polypeptide chain contains an N-terminal signal sequence (von Heijne, 1984), fusion a would probably be transported to the ER lumen.


be studied (Walter and Blobel, 1983; Mize et al., 1986; see UNIT 11.4 for analysis of protein translocation into canine microsomes). The protocol can also be adapted to microsomes prepared from metabolically labeled cells (UNIT 7.1; Bonnerot et al., 1994). As a control to determine whether the target protein is inherently sensitive to the protease used, proteolyzed membranes are compared to membranes solubilized in a mild detergent, such as Triton X-100, prior to protease addition. In this protocol, proteinase K is employed as a representative protease due to its broad substrate specificity. In addition, proteinase K is active under the buffer conditions described (as are many other proteases), and proteinase K retains activity in the presence of Triton X-100. The buffer also contains sucrose to help maintain the integrity of membrane vesicles. Another control, which tests for the presence of intact membranes, examines a luminal protein native to the membrane vesicles used. Provided the membranes are intact, the luminal protein control should be resistant to proteinase K in the absence of detergent. Examples of ER proteins used for control purposes are TRAPβ (Kalies and Hartmann, 1996) and BiP (Mullins et al., 1995). Materials Canine pancreatic microsomal membranes (see UNIT 11.4) Magnesium/sucrose/BSA (MSB) buffer (see recipe) 20% (w/v) Triton X-100 10 mg/ml proteinase K (see recipe) 100% (w/v) trichloroacetic acid (TCA) (see recipe) 2× SDS sample buffer (APPENDIX 2A) Antibodies directed against a series of peptides corresponding to specific hydrophilic regions of the target protein Control antibodies directed against a luminal protein or a luminal domain of a membrane protein in the membrane system to be analyzed Additional reagents and equipment for preparing canine pancreatic microsomes (UNIT 11.4), separating proteins by SDS-PAGE (UNIT 6.1), and detecting proteins by immunoblotting (UNIT 6.2) Prepare microsomal membranes 1. Purify microsomal membranes according to the method described in UNIT 11.4. Microsomes are stored at −80°C at a final concentration of 50 A280 units per ml. One equivalent is defined as 1 ìl of this mixture. When stored under these conditions, microsomes are stable ≥1 year. Be sure that inhibitors of the protease to be used are omitted from the original membrane preparation (such as PMSF in the case of proteinase K; see APPENDIX 1B). If protease inhibitor is present, wash the membranes by centrifuging 20 min at ∼100,000 × g, 4°C. Suspend the pellet in MSB buffer at the same concentration by pipetting up and down in a plastic tip.

2. Slowly thaw ∼50 µl of microsomes by placing a tube containing the microsomes in ice. This amount of membranes is needed for examinations that use one antipeptide antibody and a control antibody. An additional 50-ìl aliquot is needed for each additional examination.

3. Prepare ten samples, each containing 5 µl (5 equivalents) rough microsomes diluted with MSB buffer to a final volume of 20 µl in microcentrifuge tubes. Steps 3 to 8 should be performed using tubes kept in ice. Polypropylene microcentrifuge tubes have been found to work satisfactorily. Characterization of Cellular Proteins


4. Add 1 µl of 20% (w/v) Triton X-100 to five of the tubes and mix by pipetting the solution up and down. These detergent-treated aliquots serve as controls to demonstrate that the target protein is inherently sensitive to the protease used. The five tubes lacking Triton X-100 are sample tubes and the five tubes containing Triton X-100 are control tubes.

Perform protease digestion 5. Add 10 mg/ml proteinase K to four of the sample tubes and four of the control tubes to yield final concentrations of 1, 20, 100, and 500 µg/ml. The amount of protease is varied to find the concentration which allows good digestion of the target protein yet is low enough that the protease does not destroy the membrane structure (thus making the membrane permeable to the protease added). Protease should be omitted from the fifth tube in the sample and control sets.

6. Mix samples by pipetting the solutions up and down a few times and incubate 30 min. 7. Stop proteolysis by adding 100% (w/v) TCA to a final concentration of 15%. Analyze protein digests 8. Incubate all ten tubes 15 min on ice and collect TCA pellets by microcentrifuging 5 min at ∼10,000 rpm (8,000 × g), room temperature. 9. Remove supernatant and suspend the pellets in 10 µl of 2× SDS sample buffer and 10 µl H2O by vortex mixing at room temperature. Be sure that all of the pellet has been suspended. The mixtures should be blue because of bromphenol blue, which is present in the sample buffer. If the tubes are yellow (because of high acidity), 1-ìl aliquots of 1 M Tris base can be added successively and then mixed until the mixture turns blue.

10. Load 10 µl from each tube into sample wells of a SDS-PAGE gel. The lanes should be organized as follows. The first five lanes contain one-half of the mixture from the five sample tubes, and the next five lanes contain one-half of the mixture from the five control tubes. The next five lanes contain one-half of the mixture from the five sample tubes, and the final five lanes contain one-half of the mixture from the five control tubes. A 12.5% polyacrylamide gel can be used to identify protein fragments of ∼2 to 30 kDa, whereas a 7% acrylamide gel can be used to identify protein fragments of 30 to 100 kDa.

11. Analyze samples using SDS-PAGE (UNIT 6.1). 12. Examine the separated protein fragments by immunoblotting (UNIT 6.2), using one of the antipeptide antibodies and the control antibody directed against a luminal protein or a luminal domain of a membrane protein in the membrane system to be analyzed. The blot should be cut in half. The gel portion containing the first ten lanes is to be immunoblotted with an antibody directed against the target protein. The gel portion containing the second group of ten lanes should be subjected to immunoblotting using an antibody directed against the luminal protein control. The data should be interpreted by following examples described in Figure 5.2.2A and Anticipated Results.

13. Repeat steps 3 to 12 using each of the antipeptide antibodies and the control antibody.

Determining the Topology of an Integral Membrane Protein


IMMUNOFLUORESCENCE STAINING This protocol illustrates use of an immunofluorescence technique to probe the topology of membrane proteins. The protocol is designed specifically to examine proteins confined to the plasma membranes of cells lacking an outer cell wall, such as cultured mammalian cells. This protocol has been optimized for using human embryonic kidney (HEK) 293 cells, although other cultured cell lines can be used with only minor modifications (noted below). These cells can be grown on glass coverslips and permeabilized with low concentrations of a mild detergent (Nonidet P-40 is used here), which allows antibodies access to the cytoplasmic compartment. The permeabilized cells and a second nonpermeabilized cell preparation are incubated with an antipeptide antibody followed by incubation with a fluorophore-conjugated secondary antibody. Cells are then examined by fluorescence microscopy (UNIT 4.2). As a control, cells expressing a plasma membrane protein of known topology should be examined using antibodies directed to a cytosolic domain and an extracytoplasmic domain.

BASIC PROTOCOL 2

This approach can employ primary antibodies directed against peptides corresponding to the membrane protein under study, such as the antipeptide antibodies described in Basic Protocol 1. However, antipeptide antibodies are found often not to recognize their corresponding epitopes within a native membrane protein (Carrasco et al., 1986). The procedure described here therefore uses a membrane protein that has been tagged with the HA epitope (see Support Protocol for methods regarding epitope tagging). Use of the HA epitope can minimize cost and time associated with the preparation of antipeptide antibodies. The primary and secondary antibodies used in this protocol are the anti-HA epitope mouse monoclonal antibody 12CA5 and a rhodamine-conjugated rabbit antimouse IgG, respectively, but other combinations of antibodies can be used as well (Canfield and Levenson, 1993). Materials HEK 293 cells (ATCC #CRL 1573) 4% (w/v) paraformaldehyde (see recipe) Nonidet P-40/goat serum/BSA (NGB) solution (see recipe) Anti-HA mouse monoclonal antibody 12CA5 (Boehringer Mannheim) Rhodamine-conjugated rabbit anti-mouse immunoglobulin G (IgG) DMEM/FBS/HEPES (DFH) solution (see recipe) Fluoromount G mounting medium (Fisher) 6-well tissue culture plates Glass coverslips, 22-mm diameter Additional reagents and equipment for immunofluorescence staining of fixed mammalian cells (UNIT 4.3), epifluorescence (UNIT 4.2) or confocal laser microscopy, and growing cultured mammalian cells (UNIT 1.1) 1. Seed HEK 293 cells stably expressing the polypeptide of interest on glass coverslips and grow 2 to 3 days at 37°C in 6-well plates before processing for immunofluorescence. Alternatively, transient transfections can be performed on cells grown on glass coverslips in 6-well plates (UNIT 1.1). Cells should be handled using typical aseptic technique required for growing cells in culture (UNIT 1.3). The glass coverslips should be cleaned thoroughly using a strong detergent and washed extensively with distilled water. Duplicate cultures are needed in order to provide sets to be examined in the absence and presence of permeabilizing agents and in the absence or presence of the polypeptide of

Characterization of Cellular Proteins


interest. A single 6-well plate thus provides a triplicate analysis of cells to be treated with permeabilizing agents and containing the polypeptide of interest and a triplicate analysis of cells treated with permeabilizing agents and lacking the polypeptide of interest. A distinct plate provides a triplicate analysis for the set not treated with permeabilizing agents and either containing or not containing the polypeptide of interest (see below). All steps except the antibody incubations can easily be carried out in 6-well tissue culture plates, using volumes of 2 to 3 ml per well for incubations and washes. Methods for HA tagging, and testing for function of tagged proteins in transiently transfected HEK 293 cells are described in the Support Protocol.

2. For the permeabilized set, aspirate culture medium and wash coverslips once in PBS. Replace PBS with 4% paraformaldehyde (2 to 3 ml) and fix 10 min at room temperature. 3. Aspirate fixing reagent and rinse cells three times with PBS. Permeabilize and block cells in NGB solution (2 to 3 ml) for 15 min at room temperature. 4. Dilute monoclonal antibody 12CA5 to 1 µg/ml in NGB. Centrifuge antibody dilution 2 min at 8000 × g to remove aggregates. Although this antibody is supplied at a specified concentration, different batches appear to have different characteristics, and it may be necessary for individual users to optimize their antibody dilutions.

5. Place coverslips cell-side-up on Parafilm and immediately add primary antibody (100 µl per coverslip). Incubate 1 hr at room temperature in a covered, humidified chamber. A small petri dish or any small covered container will serve as a humidified chamber. For more tightly adherent cells, the volume of antibody used can be reduced further as follows. Pipet antibody (50 ìl) directly onto Parafilm. Place coverslips on droplet, cellside-down.

6. Return coverslips to NGB (2 to 3 ml) in the 6-well plate. Wash three times with PBS (10 min each wash) and once with NGB. 7. Incubate cells 1 hr at room temperature in NGB solution (2 to 3 ml) containing a rhodamine-conjugated rabbit anti-mouse IgG. The secondary antibody should be diluted according to the supplier’s recommendations. Optimization of antibody dilution may be necessary.

8. Return coverslips to NGB solution and wash three times with PBS. 9. Dip coverslips in distilled water and mount on slides using Fluoromount-G (15 µl per slide). 10. For the nonpermeabilized set, replace growth medium with DFH solution and chill 15 min to 4°C. These cells provide the set that is nonpermeable in the presence of primary antibody. These steps can most conveniently be performed concurrently with processing of permeabilized cells.

11. Incubate with monoclonal antibody 12CA5 (2 µg/ml) for 1 hr in a humidified chamber at 4°C as described in step 5. Determining the Topology of an Integral Membrane Protein

Note that a higher concentration of antibody is used for this low-temperature incubation.

12. Wash three times with cold DFH and once with cold PBS.


13. Fix, permeabilize, and incubate “nonpermeabilized” cells with secondary antibody, as described in steps 2 to 3 and 7 to 9. The nonpermeabilized set is permeabilized immediately before addition of secondary antibody. This step serves not only to bind the secondary antibody to the primary antibody but also to determine whether the secondary antibody binds nonspecifically to either surface of the plasma membrane.

14. Examine permeabilized and nonpermeabilized cells using epifluorescence or confocal laser scanning microscopy. Fluorescence staining of only the permeabilized set is suggestive of an antibody bound to a cytosolic domain, whereas fluorescence appearing in both the permeabilized and nonpermeabilized sets reveals a domain placed at the extracytoplasmic side of the plasma membrane. A range from 5% to 25% of the cells on a particular coverslip may be stained well. The control cells lacking the polypeptide of interest are needed because one difficulty associated with the use of 12CA5 is that it is not absolutely specific for the HA epitope. Cross-reactivity is seen both on immunoblots and by immunofluorescence in nontransfected or nontagged cells.The resulting background may decrease the sensitivity with which the transfected, tagged polypeptide can be visualized. If the background signal is too high to detect the protein of interest, even after diluting the primary and secondary antibodies further than that described above, try other anti-HA antibody preparations such as those available from BAbCO. Note, however, that some of these antibodies may not recognize HA tags placed internally within the protein sequence.

EPITOPE TAGGING In this protocol, a series of molecular biological manipulations are used to place a foreign epitope into a hydrophilic region of a membrane protein. Integration of the expressed protein leads to placement of the epitope on one side of the membrane. The orientation of the tagged domain can be assessed by monitoring the accessibility of the epitope to a protease (see Basic Protocol 1) or to its cognate antibody (see Basic Protocol 2).

SUPPORT PROTOCOL

The HA epitope, YPYDVPDYA, is recommended for use in this protocol because it contains only two charged amino acids. As shown in Figure 5.2.3, a set of three DNA fragments (differing by addition of 0, 1, or 2 base pairs at one or both ends) can be used to place the HA epitope internally within the sequence of a protein, N-terminally, or C-terminally. The fragments are inserted into a blunt-end restriction site occurring naturally within the protein’s gene or into a restriction site constructed by site-directed mutagenesis (see APPENDIX 3). Depending on the reading frame of the site into which the epitope is inserted, only one of the three DNA fragments is needed to permit in-frame fusion between the membrane protein sequence and the epitope at its N and/or C termini. These fragments also contain the blunt-end PvuII restriction site. This site provides a convenient place to insert a second fragment encoding the HA epitope, which is used if the single tag is found not to present the membrane protein adequately to the added antibody. Alternatively, DNA fragments containing a blunt-end restriction site other than PvuII, such as EcoRV, can be synthesized if this PvuII site is not unique on the plasmid containing the gene of interest. The DNA sequence encoding the HA epitope depicted in Figure 5.2.3 also contains an AatII restriction site that can be used to verify insertion of the fragment into the plasmid used. It is usually necessary to verify orientation of the inserted fragment by DNA sequencing. This is not the only way to introduce an epitope into a membrane protein. A DNA sequence encoding the epitope can be introduced by site-directed mutagenesis. Having introduced a DNA fragment encoding the HA epitope into a cloned gene, the tagged protein should be tested for function. If the tagged protein exhibits normal functional properties, then it is likely that the epitope introduced has not altered the overall



structure of the protein. To test for function, many types of assays can be performed. The type of assay used is dependent on the particular protein under study. For example, the HA-tagged rodent Na,K-ATPase α subunit can be tested for its ability to confer ouabain resistance to HEK 293 cells transfected with the tagged DNA construct (see Basic Protocol 2; Canfield et al., 1996). A tagged protein can be introduced into yeast cells, testing for complementation of a mutant lacking that protein or an evolutionarily conserved protein (Kurihara and Silver, 1993). Alternatively, a tagged protein may be expressed to high levels, enriched, then tested for activity using an in vitro assay. Materials TE buffer (APPENDIX 2A) Plasmid DNA encoding the protein of interest E. coli cells to be transformed Additional reagents and equipment for synthesizing oligonucleotides (APPENDIX 3), ligating DNA (APPENDIX 3), transforming E. coli (APPENDIX 3), isolating plasmid DNA from E. coli (APPENDIX 3), identifying plasmids by restriction endonuclease digestion (APPENDIX 3), and sequencing oligonucleotides (APPENDIX 3) 1. Determine whether fragment I, II, or III (Fig. 5.2.3) is needed to produce an in-frame fusion between the N- and/or C-terminal ends of the HA epitope and the target protein. 2. Synthesize the appropriate DNA fragment(s) identified from step 1. Complementary oligonucleotides should be synthesized.

3. Mix two complementary DNA strands together, each to a final concentration of 1 A260 unit per 50 µl in TE buffer. Oligonucleotides are usually synthesized with their 5′ ends unphosphorylated. The oligonucleotides used should be left unphosphorylated to prevent insertion of multiple oligonucleotides into the linearized plasmid.

4. Anneal the strands by heating the solution 5 min at 70°C and then letting the solution cool in air at room temperature. 5. Mix 5 µl of the oligonucleotide solution with 0.1 µg of linearized plasmid DNA encoding the protein of interest, ligate the mixture in 10 µl of ligation buffer, and transform E. coli with the ligation mixture (APPENDIX 3). The DNA should be linearized at a restriction site contained within a sequence corresponding to the hydrophilic domain to be probed. If such a restriction site does not exist, one can be introduced by site-directed mutagenesis (as illustrated below, blunt-end sites are preferable). If possible, the epitope should be placed ≥15 amino acids from the nearby transmembrane segment(s). If the restriction site used produces sticky ends, the site should be made blunt-ended using standard molecular biological manipulations. To minimize religation of the plasmid without the DNA fragment to be inserted, the ligation mixture should be treated with the restriction enzyme used to produce the blunt end, provided a restriction enzyme producing blunt ends was used. The above-described ratio of fragment and linear plasmid is acceptable in many instances. However, sometimes this ratio may need to be varied in order to achieve efficient ligation of the fragment to the plasmid. Try increasing the concentration of oligonucleotides from 1 A260 unit to 2, 5, and 10 A260 units per 50 ìl in TE buffer (step 3). Determining the Topology of an Integral Membrane Protein

Any of the E. coli strains commonly used for plasmid transformations is suitable, such as HB101 and MC1061.

6. Isolate plasmids from individual E. coli transformants and identify plasmids containing the insert using restriction analysis (APPENDIX 3).


If a fragment shown in Figure 5.2.3 is synthesized, restriction enzymes AatII and PvuII are useful for identifying the desired construct.

7. Determine the DNA sequence of the fragment inserted and residues surrounding this fragment to ensure correct orientation and in-frame fusion. 8. Express the epitope-tagged protein in an appropriate cell and test for its function. Examine its topology using Basic Protocols 1 or 2. If the tagged protein is not recognized by antibodies directed against the HA epitope, double-HA-tagged proteins can be constructed by inserting the appropriate DNA fragment (Fig. 5.2.3) into the PvuII site of the construct tagged with only one HA epitope. Successive insertions into each new PvuII site will produce proteins tagged with increasing numbers of HA epitopes. Plasmid DNA can be stored indefinitely in TE buffer at −20°C and should be introduced into the cell examined prior to use.

REPORTER GENE FUSIONS In this protocol, a series of gene fusions is constructed in which a reporter moiety is fused to various truncated fragments of a target membrane protein. To construct these fusions, DNA fragments encoding a series of N-terminal fragments of a membrane protein should be synthesized by the polymerase chain reaction (PCR; see APPENDIX 3). For each PCR amplification, two primers are required: one corresponding to a sequence upstream of the promoter of the relevant gene and one corresponding to sequences at the desired fusion joint. Only one upstream primer is required, whereas a downstream primer corresponding to each of the fusion joints is usually needed. For construction of PCR fragments, the upstream primer contains a BamHI site or a site compatible with the BamHI sticky end, such as BglII or BclI. The downstream primer contains a XhoI site or a site compatible with XhoI, such as SalI. Compatible sites are needed when the DNA fragment to be inserted contains an internal BamHI or XhoI site. To ensure that the reading frame is maintained across the fusion joint, the XhoI site in the downstream primer is placed in the following reading frame: C TCG AG (where TCG encodes an in-frame serine). A vector that can be used for expression of fusions in yeast is pA189invHD (Green and Walter, 1992; Fig. 5.2.7). pA189invHD is used for construction of gene fusions encoding the C-terminal moiety histidinol dehydrogenase (HD). HD is a cytoplasmic enzyme that is enzymatically inactive when fused to a luminal domain of a membrane protein inserted into the ER membrane. This shuttle vector contains the 2µm DNA fragment for replication in yeast and the ColE1 replication origin for replication in E. coli. pA189invHD can be selected in yeast cells containing a mutation in the URA3 gene due to the presence of plasmid-borne URA3. The vector can also be maintained in E. coli, as it confers ampicillin resistance. pA189invHD contains a single BamHI site and a single XhoI site. Restriction of pA189invHD with BamHI and XhoI generates two DNA fragments (13 kb and 0.8 kb). The larger of these fragments contains the selectable markers and origins of replication described above. The smaller fragment encodes a portion of arginine permease, whose topology was analyzed previously using this vector system (Green and Walter, 1992). The fragment encoding arginine permease is therefore replaced with PCR-amplified fragments encoding N-terminal truncations of the membrane protein to be examined. For examining the topology of a protein without its native promoter, the yeast ADH1 promoter (Bennetzen and Hall, 1982) is included on pA189invHD. When using the ADH1 promoter, the upstream primer should contain a BamHI site in front of the initiation codon for the membrane protein. For efficient expression from the ADH1 promoter in pA189invHD, it is important that no ATG codon is present between the BamHI site and the initiation codon.

BASIC PROTOCOL 3



Materials Reporter plasmid (Fig. 5.2.7): pA189invHD (available from Neil Green, Vanderbilt University) S. cerevisiae strain FC2-12B (MATα trp1-1 leu2-1 ura3-52 his4-401 HOL1-1 can1-1; available from Neil Green, Vanderbilt University) SD +HIS agar plates (see recipe) SD +HOL agar plates (see recipe) Thermocycler Additional reagents and equipment for the polymerase chain reaction (PCR; APPENDIX 3), agarose gel electrophoresis (APPENDIX 3), restriction endonuclease digestion (APPENDIX 3), and transformation of E. coli and S. cerevisiae (APPENDIX 3) Construct gene fusions 1. Amplify by PCR a series of DNA fragments encoding truncations of a membrane protein, using appropriate primers. The DNA primers used in the PCR amplification can be synthesized by one of a number of companies offering services in oligonucleotide synthesis. The upstream primer should

BLA

2µm circle DNA

pA189invHD ori URA3 BamHI

BamHI

ADH1

ADH1 promoter

XhoI

inv-HD

XhoI C TCG AGA TCC TTC SER ARG SER PHE inv-HD


Figure 5.2.7 Gene fusion vector pA189invHD. The direction of transcription is indicated by an arrow. Abbreviations: BLA, β-lactamase gene; ADH1, alcohol dehydrogenase gene promoter; URA3, orotidine-5′-phosphate decarboxylase gene; ori, ColE1 replication origin; inv-HD, fusion of invertase fragment-histidinol dehydrogenase genes.


contain a 5′ BamHI site (or compatible site) followed by at least 20 nucleotides complementary to the DNA sequence to be amplified. The downstream primer should contain a 5′ XhoI site (or compatible site) followed by at least 20 nucleotides from the reverse strand of the DNA sequence to be amplified. The BamHI site should be placed immediately before the initiation codon if the ADH1 promoter of pA189invHD is to be used or before the promoter of the gene to be studied. The XhoI site should correspond to a site in a hydrophilic domain of the membrane protein studied. It is also recommended that the primers contain 2 to 3 nucleotides at the 5′ end of the restriction site for efficient cutting by the restriction enzyme after PCR amplification.

2. Purify the amplified fragments by agarose gel electrophoresis and digest with BamHI and XhoI. Overnight restrictions are sometimes needed for efficient cutting of XhoI sites located near the ends of DNA fragments.

3. Digest the reporter plasmid pA189invHD with BamHI and XhoI. 4. Purify the larger, 13-kb DNA fragment produced from the restriction digestion by agarose gel electrophoresis (see APPENDIX 3). Ligate the 13-kb fragment to each of the fragments produced in step 2. The smaller fragment (0.8 kb) encodes a part of arginine permease and should be discarded.

fusion of HD to a luminal domain

SD +HIS

SD +HOL fusion of HD to a cytosolic domain

SD +HIS

SD +HOL

Figure 5.2.8 Cell growth assay for determining membrane protein topology. Cells of yeast strain FC2-12B bearing a fusion protein that fuses the HD moiety to a luminal domain of an integral membrane protein or bearing a fusion protein that fuses the HD moiety to a cytosolic domain of an integral membrane protein are streaked for single colonies on SD +HIS agar plates and SD +HOL agar plates, then incubated 4 to 5 days at 30°C. When large colonies appear on SD +HIS agar plates and little growth is detected on SD +HOL agar plates, the reporter is interpreted to be fused to a luminal domain of the membrane protein under study. On the other hand, when large colonies appear on both the SD +HIS and SD +HOL agar plates, the reporter is interpreted to be fused to a cytosolic domain.



5. Transform a standard Amps E. coli strain and isolate the desired plasmid construct. A variety of E. coli strains can be used, such as strains HB101 and MC1061.

Perform genetic assay 6. Transform the construct isolated from E. coli into S. cerevisiae strain FC2-12B and select transformants on SD +HIS agar plates. Yeast cells should be handled using aseptic technique. Strain FC2-12B contains a ura3 mutation that permits selection for pA189invHD-derived plasmids. Transformants should appear as individual colonies after 4 days on agar plates incubated at 30°C.

7. Test transformed yeast cells for growth by streaking cells on SD +HOL agar plates and SD +HIS agar plates. Individual colonies should appear after 4 to 5 days at 30°C. An example of the results expected is shown in Figure 5.2.8. As shown in this figure, fusion of HD to a luminal or cytosolic domain does not affect cell growth on SD +HIS agar plates. However, fusion of HD to a luminal domain inhibits growth on SD +HOL agar plates. Fusion of HD to a cytosolic domain does not inhibit cell growth on either type of agar plate.


DMEM/FBS/HEPES (DFH) solution 10% (v/v) fetal bovine serum 20 mM N-2-hydroxyethylpiperazine-N′-2-ethanesulfonic acid (HEPES), pH 7.4 Prepare in supplemented Dulbecco’s modified Eagle medium (DMEM; APPENDIX 2A) Prepare fresh The fetal bovine serum should not be heat-inactivated.

Magnesium/sucrose/BSA (MSB) buffer 150 mM potassium acetate 5 mM magnesium acetate 50 mM N-2-hydroxyethylpiperazine-N′-2-ethanesulfonic acid (HEPES), pH 7.6 200 mM sucrose 1 mM dithiothreitol (APPENDIX 2A) Prepare fresh Nonidet P-40/goat serum/BSA (NGB) solution 0.05% (v/v) Nonidet P-40 (Igepal AC-630) 5% (v/v) goat serum 1% (w/v) BSA Prepare in PBS (APPENDIX 2A), pH 7.4 Prepare fresh


Paraformaldehyde, 4% Heat 900 ml water to 55° to 60°C on a stirring hot plate in a fume hood. Add 40 g paraformaldehyde powder and stir 30 min. If powder has not dissolved, add a few NaOH pellets one at a time (waiting a few minutes between pellets) until the paraformaldehyde dissolves. Add 100 ml of 10× PBS (APPENDIX 2A), filter, cool to room temperature, and adjust to pH 7.4 with HCl. Store up to 1 week at 4°C. CAUTION: The fume hood is used because paraformaldehyde fumes are toxic. The “Prill” form of paraformaldehyde from EM Sciences is safest to use, as it pours without creating a cloud.


Proteinase K, 10 mg/ml 10 mg proteinase K (lyophilized powder, ∼80% protein, 10 to 20 U/mg protein; Sigma) 1 ml 50 mM HEPES, pH 7.6 Prepare fresh on ice No activation of the proteinase K is necessary; the purchased enzyme is ready to use.

SD +HIS agar plates 0.7 g/liter yeast extract (without amino acids; Difco) 20 g/liter glucose 20 g/liter agar 0.1 mg/ml L-tryptophan 0.1 mg/ml L-leucine 0.1 mg/ml L-histidine Mix the ingredients together before autoclaving. Pour the autoclaved solution into 100 × 15–mm petri dishes. Store plates @GHFDQ`\O SKHQ\OSKRVSKDWH$3

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Immunoprecipitation

UNIT 7.2

Immunoprecipitation is a technique in which an antigen is isolated by binding to a specific antibody attached to a sedimentable matrix. The source of antigen for immunoprecipitation can be unlabeled cells or tissues, metabolically or extrinsically labeled cells (UNIT 7.1), subcellular fractions from either unlabeled or labeled cells (see Chapter 3), or in vitro–translated proteins (UNIT 11.2). Immunoprecipitation is also used to analyze protein fractions separated by other biochemical techniques such as gel filtration or sedimentation on density gradients (UNIT 5.3). Either polyclonal or monoclonal antibodies from various animal species can be used in immunoprecipitation protocols. Antibodies can be bound noncovalently to immunoadsorbents such as protein A– or protein G–agarose, or can be coupled covalently to a solid-phase matrix. Immunoprecipitation protocols consist of several stages (Fig. 7.2.1; see Basic Protocol 1). In stage 1, the antigen is solubilized by one of several techniques for lysing cells. Soluble and membrane-associated antigens can be released from cells grown either in suspension culture (see Basic Protocol 1) or as a monolayer on tissue culture dishes (see Alternate Protocol 1) with nondenaturing detergents. Cells can also be lysed under denaturing conditions (see Alternate Protocol 2). Soluble antigens can also be extracted by mechanical disruption of cells in the absence of detergents (see Alternate Protocol 3). All of these lysis procedures are suitable for extracting antigens from animal cells. Yeast cells require disruption of their cell wall in order to allow extraction of the antigens (see Alternate Protocol 4). In stage 2, a specific antibody is attached, either noncovalently or covalently, to a sedimentable, solid-phase matrix to allow separation by low-speed centrifugation. This unit presents the noncovalent attachment of antibody to protein A– or protein G–agarose beads (see Basic Protocol 1). Stage 3 consists of incubating the solubilized antigen from stage 1 with the immobilized antibody from stage 2, followed by extensive washing to remove unbound proteins (see Basic Protocol 1). Immunoprecipitated antigens can be dissociated from antibodies and reprecipitated by a protocol referred to as “immunoprecipitation-recapture” (see Basic Protocol 2). This protocol can be used with the same antibody for further purification of the antigen, or with a second antibody to identify components of multisubunit complexes or to study protein-protein interactions (Fig. 7.2.2). Immunoprecipitated antigens can be analyzed by one-dimensional electrophoresis (UNIT 6.1), two-dimensional electrophoresis, or immunoblotting (UNIT 6.2). In some cases, immunoprecipitates can be used for structural or functional analyses of the isolated antigens. Immunoprecipitates can also be used as sources of immunogens for production of monoclonal or polyclonal antibodies. IMMUNOPRECIPITATION USING CELLS IN SUSPENSION LYSED WITH A NONDENATURING DETERGENT SOLUTION

BASIC PROTOCOL 1

In this protocol, unlabeled or labeled cells in suspension are extracted by incubation in nondenaturing lysis buffer containing the nonionic detergent Triton X-100 (steps 1 to 7). This procedure results in the release of both soluble and membrane proteins; however, many cytoskeletal and nuclear proteins, as well as a fraction of membrane proteins, are not efficiently extracted under these conditions (see UNIT 5.1). The procedure allows immunoprecipitation with antibodies to epitopes that are exposed in native proteins. For immunoprecipitation, a specific antibody is immobilized on a sedimentable, solidphase matrix (steps 8 to 14). Although there are many ways to attach antibodies to matrices (see Commentary), the most commonly used methods rely on the property of immunoglobulins to bind Staphylococcus aureus protein A, or protein G from group G Streptococcus (Table 7.2.1). The best results are obtained by binding antibodies to protein A or Contributed by Juan S. Bonifacino and Esteban C. Dell’Angelica Current Protocols in Cell Biology (1998) 7.2.1-7.2.21 Copyright © 1998 by John Wiley & Sons, Inc.

Protein Labeling and Immunoprecipitation

7.2.1

protein G that is covalently coupled to agarose beads. In this protocol, Sepharose beads are used (Sepharose is a more stable, cross-linked form of agarose). Immunoprecipitation is most often carried out using rabbit polyclonal or mouse monoclonal antibodies, which, with some exceptions (e.g., mouse IgG1), bind well to protein A (Table 7.2.1). Antibodies that do not bind to protein A–agarose can be adsorbed to protein G–agarose (Table 7.2.1) using exactly the same protocol. For optimal time management, incubation of antibodies with protein A–agarose can be carried out either before or during lysis of the cells. The final stage in immunoprecipitation is combining the cell lysate with the antibodyconjugated beads and isolating the antigen (steps 18 to 26). This can be preceded by an optional preclearing step in which the lysate is absorbed with either “empty” protein A–agarose beads or with an irrelevant antibody bound to protein A–agarose (steps 15 to

animal cell

protein A- agarose bead specific antigen

antibody

1

cell lysis (see Basic Protocol 1, steps 1 to 7; Alternate Protocols 1 to 4)

2

antibody binding to protein A- agarose bead (see Basic Protocol 1, steps 8 to 14)

3 antigen isolation on antibody bead (see Basic Protocol 1, steps 18 to 21)

wash (see Basic Protocol 1, steps 22 to 26) and analysis

Immunoprecipitation

Figure 7.2.1 Schematic representation of the stages of a typical immunoprecipitation protocol. (1) Cell lysis: antigens are solubilized by extraction of the cells in the presence or absence of detergents. To increase specificity, the cell lysate can be precleared with protein A–agarose beads (steps 15 to 17, not shown). (2) Antibody immobilization: a specific antibody is bound to protein A–agarose beads. (3) Antigen capture: the solubilized antigen is isolated on antibody-conjugated beads.


17). The need for preclearing depends on the specific experimental system being studied and the quality of the antibody reagents. The protocol described below incorporates a preclearing step using protein A–agarose. Protein fractions separated by techniques such as gel filtration or sedimentation on sucrose gradients (UNIT 5.3) can be used in place of the cell lysate at this stage. After binding the antigen to the antibody-conjugated beads, the unbound proteins are removed by successive washing and sedimentation steps. Materials Unlabeled or labeled cells in suspension (UNIT 7.1) PBS (APPENDIX 2A), ice cold Nondenaturing lysis buffer (see recipe), ice cold 50% (v/v) protein A–Sepharose bead (Sigma, Pharmacia Biotech) slurry in PBS containing 0.1% (w/v) BSA and 0.01% (w/v) sodium azide (NaN3)

Table 7.2.1 Ga,b,c

Binding of Antibodies to Protein A and Protein

Protein A binding

Protein G bindingd

Monoclonal antibodiese Human IgG1 Human IgG2 Human IgG3 Human IgG4 Mouse IgG1 Mouse IgG2a Mouse IgG2b Mouse IgG3 Rat IgG1 Rat IgG2a Rat IgG2b Rat IgG2c

++ ++ − ++ + ++ ++ ++ + − − ++

++ ++ ++ ++ ++ ++ ++ ++ + ++ ++ ++

Polyclonal antibodies Chicken Donkey Goat Guinea pig Hamster Human Monkey Mouse Rabbit Rat Sheep

− − + ++ + ++ ++ ++ ++ + +

− ++ ++ + ++ ++ ++ ++ ++ + ++

Antibody

a++, moderate to strong binding; +, weak binding; −, no binding. bA hybrid protein A/G molecule that combines the features of protein A and

protein G, coupled to a solid-phase matrix, is available from Pierce. cInformation from Harlow and Lane (1988), and from Pharmacia Biotech,

Pierce, and Jackson Immunoresearch. dNative protein G binds albumin from several animal species. Recombinant variants of protein G have been engineered for better binding to rat, mouse, and guinea pig IgG, as well as for avoiding binding to serum albumin. eProtein A binds some IgM, IgA, and IgE antibodies in addition to IgG, whereas protein G binds only IgG.



Specific polyclonal antibody (antiserum or affinity-purified immunoglobulin) or monoclonal antibody (ascites, culture supernatant, or purified immunoglobulin) Control antibody of same type as specific antibody (e.g., preimmune serum or purified irrelevant immunoglobulin for specific polyclonal antibody; irrelevant ascites, culture supernatant, or purified immunoglobulin for specific monoclonal antibody; see Critical Parameters) 10% (w/v) BSA (APPENDIX 2A) Wash buffer (see recipe), ice cold Microcentrifuge with fixed-angle rotor (Eppendorf 5415C or equivalent) Tube rotator (capable of end-over-end inversion) CAUTION: When working with radioactivity, take appropriate precautions to avoid contamination of the experimenter and the surroundings. Carry out the experiment and dispose of wastes in an appropriately designated area, following the guidelines provided by the local radiation safety officer (also see APPENDIX 1D). NOTE: All solutions should be ice cold and procedures should be carried out at 4°C or on ice. Prepare cell lysate 1. Collect cells in suspension by centrifuging 5 min at 400 × g, 4°C, in a 15- or 50-ml capped conical tube. Place tube on ice. Approximately 0.5–2 × 107 cells are required to yield 1 ml lysate, which is generally used for each immunoprecipitation. Labeled cells are likely to have been pelleted earlier as part of the labeling procedure. If the cells are frozen, they should be thawed on ice before solubilization.

2. Aspirate supernatant with a Pasteur pipet attached to a vacuum trap. CAUTION: Dispose of radioactive materials following applicable safety regulations (APPENDIX 1D).

3. Resuspend cells gently by tapping the bottom of the tube. Rinse cells twice with ice-cold PBS as in steps 1 and 2, using the same volume of PBS as in the initial culture. 4. Add 1 ml ice-cold nondenaturing lysis buffer per ∼0.5–2 × 107 cells and resuspend pellet by gentle agitation for 3 sec with a vortex mixer set at medium speed. Do not shake vigorously as this could result in loss of material or protein denaturation due to foaming.

5. Keep suspension on ice 15 to 30 min and transfer to a 1.5-ml conical microcentrifuge tube. Tubes can have flip-top or screw caps. Screw-capped tubes are preferred because they are less likely to open accidentally during subsequent procedures. They are also recommended for work with radioactivity.

6. Clear the lysate by microcentrifuging 15 min at 16,000 × g (maximum speed), 4°C. Centrifugation can be carried out in a microcentrifuge placed in a cold room or in a refrigerated microcentrifuge. Take precautions to ensure that the 4°C temperature is maintained during the spin (e.g., use a fixed-angle rotor with a lid, as the aerodynamics of this type of rotor reduces generation of heat by friction). If it is necessary to reduce background, the lysate can be spun for 1 hr at 100,000 × g in an ultracentrifuge. Immunoprecipitation


7. Transfer the supernatant to a fresh microcentrifuge tube using an adjustable pipet fitted with a disposable tip. Do not disturb the pellet, and leave the last 20 to 40 µl of supernatant in the centrifuge tube. Keep the cleared lysate on ice until preclearing (step 15) or addition of antibody beads (step 18). NOTE: Resuspension of even a small amount of sedimented material will result in high nonspecific background due to carryover into the immunoprecipitation steps. A cloudy layer of lipids floating on top of the supernatant will not adversely affect the results of the immunoprecipitation. When the lysate is highly radioactive—as is the case for metabolically labeled cells—the use of tips with aerosol barriers is recommended to reduce the risk of contaminating internal components of the pipet. Cell extracts can be frozen at −70°C until used for immunoprecipitation. However, it is preferable to lyse the cells immediately before immunoprecipitation in order to avoid protein degradation or dissociation of protein complexes. If possible, freeze the cell pellet from step 3 rather than the supernatant from step 7.

Prepare antibody-conjugated beads 8. In a 1.5-ml conical microcentrifuge tube, combine 30 µl of 50% protein A–Sepharose bead slurry, 0.5 ml ice-cold PBS, and the following quantity of specific antibody (select one): 1 to 5 µl polyclonal antiserum 1 µg affinity-purified polyclonal antibody 0.2 to 1 µl ascitic fluid containing monoclonal antibody 1 µg purified monoclonal antibody 20 to 100 µl hybridoma culture supernatant containing monoclonal antibody. The quantities of antibody suggested are rough estimates based on the expected amount of specific antibodies in each preparation. Quantities can be increased or decreased, depending on the quality of the antibody preparation (see Commentary). Substitute protein G for protein A if antibodies are of a species or subclass that does not bind to protein A (see Table 7.2.1). If the same antibody will be used to immunoprecipitate multiple samples (e.g., samples from a pulse-chase experiment; UNIT 7.1), the quantities indicated above can be increased proportionally to the number of samples and incubated in a 15-ml capped conical tube. In this case, the beads should be divided into aliquots just prior to the addition of the cleared cell lysate (step 18). Antibody-conjugated beads can be prepared prior to preparation of the cell lysate (steps 1 to 7), in order to minimize the time that the cell extract is kept on ice.

9. Set up a nonspecific immunoprecipitation control in a 1.5-ml conical microcentrifuge tube by incubating 30 µl of 50% protein A–Sepharose bead slurry, 0.5 ml ice-cold PBS, and the appropiate control antibody (select one): 1 to 5 µl preimmune serum as a control for a polyclonal antiserum 1 µg purified irrelevant polyclonal antibody (an antibody to an epitope that is not present in the cell lysate) as a control for a purified polyclonal antibody 0.2 to 1 µl ascitic fluid containing irrelevant monoclonal antibody (an antibody to an epitope that is not present in the cell lysate and of the same species and immunoglobulin subclass as the specific antibody) as a control for an ascitic fluid containing specific monoclonal antibody



1 µg purified irrelevant monoclonal antibody as a control for a purified monoclonal antibody 20 to 100 µl hybridoma culture supernatant containing irrelevant monoclonal antibody as a control for a hybridoma culture supernatant containing specific monoclonal antibody The amount of irrelevant antibody should match that of the specific antibody and the antibody should be from the same species as the specific antibody.

10. Mix suspensions thoroughly. Tumble incubation mixtures end over end ≥1 hr at 4°C in a tube rotator. Addition of 0.01% (w/v) Triton X-100 may facilitate mixing of the suspension during tumbling. Incubations can be carried out for as long as 24 hr. This allows preparation of the antibody-conjugated beads prior to immunoprecipitation.

11. Microcentrifuge 2 sec at 16,000 × g (maximum speed), 4°C. 12. Aspirate the supernatant (containing unbound antibodies) using a fine-tipped Pasteur pipet connected to a vacuum aspirator. 13. Add 1 ml nondenaturing lysis buffer and resuspend the beads by inverting the tube 3 or 4 times. For lysates prepared with detergents (this protocol and see Alternate Protocols 1 and 2), use 1 ml nondenaturing lysis buffer; for lysates prepared by mechanical disruption (see Alternate Protocol 3), use detergent-free lysis buffer (see recipe). Use of a repeat pipettor is recommended when processing multiple samples.

14. Wash by repeating steps 11 to 13, and then steps 11 and 12 once more. At this point the beads have been washed twice with lysis buffer and are ready to be used for immunoprecipitation. Antibody-bound beads can be stored up to 6 hr at 4°C until used.

Preclear lysate (optional) 15. In a microcentrifuge tube, combine 1 ml cell lysate (from step 7) and 30 µl of 50% protein A–Sepharose bead slurry. The purpose of this step is to remove from the lysate proteins that bind to protein A–Sepharose, as well as pieces of insoluble material that may have been carried over from previous steps. If the lysate was prepared from cells expressing immunoglobulins—such as spleen cells or cultured B cells—the preclearing step should be repeated at least 3 times to ensure complete removal of endogenous immunoglobulins. If cell lysates were frozen and thawed, they should be microcentrifuged 15 min at 16,000 × g (maximum speed), 4°C, before the preclearing step.

16. Tumble end over end 30 min at 4°C in a tube rotator. 17. Microcentrifuge 5 min at 16,000 × g (maximum speed), 4°C. Immunoprecipitate 18. Add 10 µl of 10% BSA to the tube containing specific antibody bound to protein A–Sepharose beads (step 14), and transfer to this tube the entire volume of cleared lysate (from step 7 or 17). If a nonspecific immunoprecipitation control is performed, divide lysate in two ∼0.4-ml aliquots, one for the specific antibody and the other for the nonspecific control.

Immunoprecipitation

In order to avoid carryover of beads with precleared material, leave 20 to 40 ìl of supernatant on top of the pellets in the preclearing tubes. Discard beads and remaining supernatant. The BSA quenches nonspecific binding to the antibody-conjugated beads during incubation with the cell lysate.


19. Incubate 1 to 2 hr at 4°C while mixing end over end in a tube rotator. Samples can be incubated overnight, although there is an increased risk of protein degradation, dissociation of multiprotein complexes, or formation of protein aggregates.

20. Microcentrifuge 5 sec at 16,000 × g (maximum speed), 4°C. 21. Aspirate the supernatant (containing unbound proteins) using a fine-tipped Pasteur pipet connected to a vacuum aspirator. The supernatant can be kept up to 8 hr at 4°C or up to 1 month at −70°C for sequential immunoprecipitation of other antigens or for analysis of total proteins. To reutilize lysate, remove the supernatant carefully with an adjustable pipet fitted with a disposable tip. Before reprecipitation, preabsorb the lysate with protein A–Sepharose (as in steps 15 to 17) to remove antibodies that may have dissociated during the first immunoprecipitation. CAUTION: Dispose of radioactive materials following applicable safety regulations.

22. Add 1 ml ice-cold wash buffer, cap the tubes, and resuspend the beads by inverting the tube 3 or 4 times. Use of a repeat pipettor is recommended when processing multiple samples.

23. Microcentrifuge 2 sec at 16,000 × g (maximum speed), 4°C. 24. Aspirate the supernatant, leaving ∼20 µl supernatant on top of the beads. 25. Wash beads three more times (steps 22 to 24). Total wash time (steps 22 to 26) should be ∼30 min, keeping the samples on ice for 3 to 5 min between washes if necessary (see Critical Parameters).

26. Wash beads once more using 1 ml ice-cold PBS and aspirate supernatant completely with a drawn-out Pasteur pipet. The final product should be 15 ìl of settled beads containing bound antigen. Immunoprecipitates can either be processed immediately or frozen at −20°C for later analysis. For subsequent analysis of the isolated proteins prior to electrophoresis (e.g., comparison of the electrophoretic mobility of the antigen with or without treatment with glycosidases), samples can be divided into two or more aliquots after addition of PBS. Transfer aliquots of the bead suspension to fresh tubes, centrifuge and aspirate as in the previous steps. Immunoprecipitates can be analyzed by one-dimensional electrophoresis (UNIT 6.1), two-dimensional electrophoresis, or immunoblotting (UNIT 6.2).

IMMUNOPRECIPITATION USING ADHERENT CELLS LYSED WITH A NONDENATURING DETERGENT SOLUTION


Immunoprecipitation using adherent cells can be performed in the same manner as with nonadherent cells (see Basic Protocol 1). This protocol is essentially similar to steps 1 to 5 of Basic Protocol 1, but describes modifications necessary for using the same nondenaturing detergent solution to lyse cells attached to tissue culture plates. It is preferable to use cells grown on plates rather than in flasks, because the cell monolayer is more easily accessible. Additional Materials (also see Basic Protocol 1) Unlabeled or labeled cells grown as a monolayer on a tissue culture plate (UNIT 7.1) NOTE: All solutions should be ice cold and procedures should be carried out at 4°C or on ice.



1. Rinse cells attached to a tissue culture plate twice with ice-cold PBS. Remove the PBS by aspiration with a Pasteur pipet attached to a vacuum trap. CAUTION: Dispose of radioactive materials following applicable safety regulations.

2. Place the tissue culture plate on ice. 3. Add ice-cold nondenaturing lysis buffer to the tissue culture plate. Use 1 ml lysis buffer for an 80% to 90% confluent 100-mm-diameter tissue culture plate. Depending on the cell type, a confluent 100-mm dish will contain 0.5–2 × 107 cells. For other plate sizes, adjust volume of lysis buffer according to the surface area of the plate.

4. Scrape the cells off the plate with a rubber policeman, and transfer the suspension to a 1.5-ml conical microcentrifuge tube using an adjustable pipettor fitted with a disposable tip. Vortex gently for 3 sec and keep tubes on ice for 15 to 30 min. Tubes can have flip-top or screw caps. Screw-capped tubes are preferred because they are less likely to open accidentally during subsequent procedures. They are also recommended for work with radioactivity.

5. Clear the lysate and perform immunoprecipitation (see Basic Protocol 1, steps 6 to 26). ALTERNATE PROTOCOL 2

IMMUNOPRECIPITATION USING CELLS LYSED WITH DETERGENT UNDER DENATURING CONDITIONS If epitopes of native proteins are not accessible to antibodies, or if the antigen cannot be extracted from the cell with nonionic detergents, cells should be solubilized under denaturing conditions. This protocol is based on that for nondenaturing conditions (see Basic Protocol 1, steps 1 to 7), with the following modifications. Denaturation is achieved by heating the cells in a denaturing lysis buffer that contains an ionic detergent such as SDS or Sarkosyl (N-lauroylsarcosine). The denaturing lysis buffer also contains DNase I to digest DNA released from the nucleus. Prior to immunoprecipitation, the denatured protein extract is diluted 10-fold with nondenaturing lysis buffer, which contains Triton X-100; this step protects the antigen-antibody interaction from interference by the ionic detergent. Immunoprecipitation is performed as described (see Basic Protocol 1). The following protocol is described for cells in suspension culture, although it can be adapted for adherent cells (see Alternate Protocol 1). Only antibodies that react with denatured proteins can be used to immunoprecipitate proteins solubilized by this protocol. Additional Materials (also see Basic Protocol 1) Denaturing lysis buffer (see recipe) Heating block set at 95°C (Eppendorf Thermomixer 5436 or equivalent) 25-G needle attached to 1-ml syringe 1. Collect cells in suspension culture (see Basic Protocol 1, steps 1 to 3). Place tubes on ice. 2. Add 100 µl denaturing lysis buffer per ∼0.5–2 × 107 cells in the pellet. 3. Resuspend the cells by vortexing vigorously 2 to 3 sec at maximum speed. Transfer suspension to a 1.5-ml conical microcentrifuge tube. The suspension may be very viscous due to release of nuclear DNA.

Immunoprecipitation

Tubes can have flip-top or screw caps. Screw-capped tubes are preferred because they are less likely to open accidentally during subsequent procedures. They are also recommended for work with radioactivity.


4. Heat samples 5 min at 95°C in a heating block. 5. Dilute the suspension with 0.9 ml nondenaturing lysis buffer. Mix gently. The excess 1% Triton X-100 in the nondenaturing lysis buffer sequesters SDS into Triton X-100 micelles.

6. Shear DNA by passing the suspension five to ten times through a 25-G needle attached to a 1-ml syringe. If the DNA is not digested by DNase I in the denaturing lysis buffer or thoroughly sheared mechanically, it will interfere with the separation of pellet and supernatant after centrifugation. Repeat mechanical disruption until the viscosity is reduced to manageable levels.

7. Incubate 5 min on ice. 8. Clear the lysate and perform immunoprecipitation (see Basic Protocol 1, steps 6 to 26). IMMUNOPRECIPITATION USING CELLS LYSED WITHOUT DETERGENT Immunoprecipitation of proteins that are already soluble within cells (e.g., cytosolic or luminal organellar proteins) may not require the use of detergents. Instead, cells can be mechanically disrupted by repeated passage through a needle, and soluble proteins can be separated from insoluble material by centrifugation. The following protocol describes lysis of cells in a PBS-based detergent-free lysis buffer. Other buffer formulations may be used for specific proteins.


Additional Materials (also see Basic Protocol 1) Detergent-free lysis buffer (see recipe) 25-G needle attached to 3-ml syringe NOTE: All solutions should be ice-cold and procedures should be carried out at 4°C or on ice. 1. Collect and wash cells in suspension (see Basic Protocol 1, steps 1 to 3). 2. Add 1 ml of ice-cold detergent-free lysis buffer per ∼0.5–2 × 107 cells in a pellet. 3. Resuspend the cells by gentle agitation for 3 sec with a vortex mixer set at medium speed. 4. Break cells by passing the suspension 15 to 20 times through a 25-G needle attached to a 3-ml syringe. Extrusion of the cell suspension from the syringe should be rapid, although care should be exercised to prevent splashing and excessive foaming. Cell breakage can be checked under a bright-field or phase-contrast microscope. Repeat procedure until >90% cells are broken. It is helpful to check ahead of time whether the cells can be broken in this way. If the cells are particularly resistant to mechanical breakage, they can be swollen for 10 min at 4°C with a hypotonic solution containing 10 mM Tris⋅Cl, pH 7.4 (APPENDIX 2A) before mechanical disruption.

5. Clear the lysate and perform immunoprecipitation (Basic Protocol 1, steps 6 to 26).




IMMUNOPRECIPITATION USING YEAST CELLS DISRUPTED WITH GLASS BEADS Unlike animal cells, yeast cells have an extremely resistant, detergent-insoluble cell wall. To allow extraction of cellular antigens, the cell wall needs to be broken by mechanical, enzymatic, or chemical means. The most commonly used procedure consists of vigorous vortexing of the yeast suspension with glass beads. The breakage can be done in the presence or absence of detergent, as previously described for animal cells (see Basic Protocol 1, Alternate Protocol 2, and Alternate Protocol 3). The protocol described below is suitable for mechanical disruption of most yeast species, including Saccharomyces cerevisiae and Schizosaccharomyces pombe. A protocol for metabolic labeling for yeast has been described by Franzusoff et al. (1991). Additional Materials (also see Basic Protocol 1) Unlabeled or radiolabeled yeast cells Lysis buffer, ice cold: nondenaturing, denaturing, or detergent-free lysis buffer (see recipes) Glass beads (acid-washed, 425- to 600-µm diameter; Sigma) NOTE: All solutions should be ice-cold and procedures should be carried out at 4°C or on ice. 1. Collect 10 ml of yeast culture at 1 OD600 per immunoprecipitation sample, and centrifuge 5 min at 4000 × g, 4°C. Place tube on ice. 2. Remove supernatant by aspiration with a Pasteur pipet attached to a vacuum trap. CAUTION: Dispose of radioactive materials following applicable safety regulations.

3. Loosen pellet by vortexing vigorously for 10 sec. Rinse cells twice with ice-cold distilled water as in steps 1 and 2. Radiolabeled yeast cells are likely to have been pelleted earlier as part of the labeling procedure. If the pellets are frozen, they should be thawed on ice prior to cell disruption.

4. Add 3 vol ice-cold lysis buffer and 3 vol at 1 OD600 glass beads per volume of pelleted yeast cells. Use nondenaturing lysis buffer or detergent-free lysis buffer as required for the antigen under study. If the experiment requires denaturation of the antigen, the procedure can be adapted (see Alternate Protocol 2 for higher eukaryotic cells); however, the yeast cells must be broken with glass beads before heating the sample at 95°C.

5. Shake cells by vortexing vigorously at maximum speed for four 30-sec periods, keeping the cells on ice for 30 sec between the periods. Check cell breakage under a bright-field or phase-contrast microscope. It is helpful to check ahead of time if the cells can be broken in this way.

6. Remove the yeast cell lysate from the beads using a pipettor with a disposable tip. Transfer to a fresh tube. 7. Add 4 vol (see step 4) lysis buffer to the glass beads, vortex for 2 sec, and combine this supernatant with the lysate from step 6. 8. Clear the lysate and perform immunoprecipitation (see Basic Protocol 1, steps 6 to 26). Immunoprecipitation


IMMUNOPRECIPITATION-RECAPTURE Once an antigen has been isolated by immunoprecipitation, it can be dissociated from the beads and reimmunoprecipitated (“recaptured”) with either the same antibody used in the first immunoprecipitation or with a different antibody (Fig. 7.2.2). Immunoprecipitationrecapture with the same antibody allows identification of a specific antigen in cases where the first immunoprecipitation contains too many bands to allow unambiguous identifica-

BASIC PROTOCOL 2

antigen 2 antigen 1 antibody 1 antibody 2

protein A- agarose bead

1

protein A- agarose bead

denaturation (see Basic Protocol 2)

2

antibody 2 binding to protein A- agarose bead (see Basic Protocol 1, steps 8 to 14)

3 recapture

wash and analysis

Figure 7.2.2 Scheme showing the stages of immunoprecipitation-recapture. (1) Dissociation and denaturation of the antigen: an antigen immunoprecipitated with antibody 1 bound to protein A–agarose beads is dissociated and denatured by heating in the presence of SDS and DTT. (2) Immobilization of the second antibody: antibody 2 is bound to protein A–agarose beads. (3) Recapture: the denatured antigen (striped oval) is recaptured on antibody 2 bound to protein A–agarose beads. Alternatively, antibody 1 can be used again for further purification of the original antigen (square).



tion. By using a different antibody in the second immunoprecipitation, immunoprecipitation-recapture can be used to analyze the subunit composition of multi-protein complexes (Fig. 7.2.3). The feasibility of this approach depends on the ability of the second antibody to recognize denatured antigens. Dissociation of the antigen from the beads is achieved by denaturation of antigen-antibody-bead complexes at high temperature in the presence of SDS and DTT. Prior to recapture, the SDS is diluted in a solution containing Triton X-100, and the DTT is neutralized with excess iodoacetamide. Recapture is then performed as in the first immunoprecipitation (see Basic Protocol 1, step 26). Materials Elution buffer (see recipe) Beads containing bound antigen (see Basic Protocol 1, step 26) 10% (w/v) BSA (APPENDIX 2A) Nondenaturing lysis buffer (see recipe) Heating block set at 95°C (Eppendorf Thermomixer 5436 or equivalent) 1st Ab to:

IP

2nd

IP

BSA

σ3

σ3

µ3

BSA

1

2

3

4

5

200

98 66 46

30 22

15

Immunoprecipitation

Figure 7.2.3 Example of an immunoprecipitation-recapture experiment. Human M1 fibroblasts were labeled overnight with [35S]methionine (UNIT 7.1) and extracted with nondenaturing lysis buffer (see Basic Protocol 1). The cell extract was then subjected to immunoprecipitation with antibodies to BSA (irrelevant antibody control; lane 1) and to the AP-3 adaptor (σ3; lane 2), a protein complex involved in protein sorting. Notice the presence of several specific bands in lane 2. The AP-3 immunoprecipitate was denatured as described in Basic Protocol 2 and individual components of the AP-3 complex were recaptured with antibodies to two of its subunits: σ3 (Mr ∼22,000; lane 3) and µ3 (Mr ∼47,000; lane 4). An immunoprecipitation with an antibody to BSA was also performed as a nonspecific control (lane 5). The amount of immunoprecipitate loaded on lanes 1 and 2 is ∼1⁄10 the amount loaded on lanes 3 to 5. Notice the presence of single bands in lanes 3 and 4. The positions of Mr standards (expressed as 10−3 × Mr) are shown at left. IP, immunoprecipitation.


1. Add 50 µl elution buffer to 15 µl beads containing bound antigen. Mix by vortexing. The DTT in the elution buffer reduces disulfide bonds in the antigen and the antibody, and the SDS contributes to the unfolding of polypeptide chains.

2. Incubate 5 min at room temperature and 5 min at 95°C in a heating block. Cool tubes to room temperature. 3. Add 10 µl of 10% BSA. Mix by gentle vortexing. BSA is added to prevent adsorption of antigen to the tube, and to quench nonspecific binding to antibody-conjugated beads.

4. Add 1 ml nondenaturing lysis buffer. The iodoacetamide in the nondenaturing lysis buffer reacts with the DTT and prevents it from reducing the antibody used in the recapture steps. The presence of PMSF and leupeptin in the buffer is not necessary at this step.

5. Incubate 10 min at room temperature. 6. Clear the lysate and perform second immunoprecipitation (see Basic Protocol 1, steps 6 to 26). REAGENTS AND SOLUTIONS Use deionized or distilled water in all recipes and protocol steps. For common stock solutions, see APPENDIX 2A; for suppliers, see SUPPLIERS APPENDIX.

Denaturing lysis buffer 1% (w/v) SDS 50 mM Tris⋅Cl, pH 7.4 (APPENDIX 2A) 5 mM EDTA (APPENDIX 2A) Store up to 1 week at room temperature (SDS precipitates at 4°C) Add the following fresh before use: 10 mM dithiothreitol (DTT, from powder) 1 mM PMSF (APPENDIX 2A) 2 µg/ml leupeptin (store 10 mg/ml stock in H2O up to 6 months at −20°C) 15 U/ml DNase I (store 15,000 U/ml stock solution up to 2 years at −20°C) 1 mM 4-(2-aminoethyl)benzenesulfonyl fluoride (AEBSF), added fresh from a 0.1 M stock solution in H2O, can be used in place of PMSF. AEBSF stock can be stored up to 1 year at −20°C.

Detergent-free lysis buffer PBS (APPENDIX 2A) containing: 5 mM EDTA (APPENDIX 2A) 0.02% (w/v) sodium azide Store up to 6 months at 4°C Immediately before use add: 10 mM iodoacetamide (from powder) 1 mM PMSF (APPENDIX 2A) 2 µg/ml leupeptin (store 10 mg/ml stock in H2O up to 6 months at −20°C) 1 mM 4-(2-aminoethyl)benzenesulfonyl fluoride (AEBSF), added fresh from a 0.1 M stock solution in H2O, can be used in place of PMSF. AEBSF stock can be stored up to 1 year at −20°C. Protein Labeling and Immunoprecipitation


Elution buffer 1% (w/v) SDS 100 mM Tris⋅Cl, pH 7.4 (APPENDIX 2A) Store up to 1 week at room temperature 10 mM dithiothreitol (DTT, add fresh from powder before use) Nondenaturing lysis buffer 1% (w/v) Triton X-100 50 mM Tris⋅Cl, pH 7.4 (APPENDIX 2A) 300 mM NaCl 5 mM EDTA (APPENDIX 2A) 0.02% (w/v) sodium azide Store up to 6 months at 4°C Immediately before use add: 10 mM iodoacetamide (from powder) 1 mM PMSF (APPENDIX 2A) 2 µg/ml leupeptin (store 10 mg/ml stock in H2O up to 6 months at −20°C) 1 mM 4-(2-aminoethyl)-benzenesulfonyl fluoride (AEBSF), added fresh from a 0.1 M stock solution in H2O, can be used in place of PMSF. AEBSF stock can be stored up to 1 year at −20°C.

Wash buffer 0.1% (w/v) Triton X-100 50 mM Tris⋅Cl, pH 7.4 (APPENDIX 2A) 300 mM NaCl 5 mM EDTA (APPENDIX 2A) 0.02% (w/v) sodium azide Store up to 6 months at 4°C COMMENTARY Background Information

Immunoprecipitation

The use of antibodies for immunoprecipitation has its origin in the precipitin reaction (Nisonoff, 1984). The term precipitin refers to the spontaneous precipitation of antigen-antibody complexes formed by interaction of certain polyclonal antibodies with their antigens. The precipitation arises from formation of large networks of antigen-antibody complexes, due to the bivalent or polyvalent nature of immunoglobulins and to the presence of two or more epitopes in some antigens. This phenomenon was quickly exploited to isolate antigens from protein mixtures; however, its use remained limited to antibodies and antigens that were capable of multivalent interaction. In addition, the efficiency of precipitate formation was highly dependent on the concentrations of antibody and antigen. Thus, the precipitin reaction was not generally applicable as a method for immunoprecipitation. A significant improvement was the use of secondary anti-immunoglobulin reagents (generally anti-immunoglobulin serum) to crosslink the primary antibodies, thus promoting the

formation of a precipitating network. Protocols based on the use of cross-linking secondary antibodies are still used in immunoprecipitation and are reputed to give very low backgrounds (Springer, 1996). In the 1970s, immunoprecipitation became widely applicable to the study of cellular antigens as a result of several technological advances. A critical development was the introduction of methods for the production of monoclonal antibodies (Köhler and Milstein, 1975). The ability to produce unlimited amounts of antibodies with specificity against virtually any cellular antigen had a profound impact in many areas of biology and medicine. The fact that preparation of monoclonal antibodies did not require prior purification of the antigens accelerated the characterization of cellular proteins and organelles, a process in which immunoprecipitation protocols played a major role. To this day, monoclonal antibodies produced in mice or rats continue to be among the most useful tools in cell biology. Another important development was the discovery of bacterial Fc receptors, proteins found


on the surface of bacteria that have the property of binding a wide range of immunoglobulins. Two of the most widely used bacterial Fc receptors are protein A from Staphylococcus aureus and protein G from group G streptococci. Protein A and protein G bind both polyclonal and monoclonal antibodies belonging to different subclasses and from different animal species (Table 7.2.1). Protein A was initially used to adsorb immunoglobulins as part of fixed, killed Staphylococcus aureus particles. Both protein A and protein G are now produced in large quantities by recombinant DNA procedures and are available coupled to solid-phase matrices such as agarose. In most cases, the binding of polyclonal or monoclonal antibodies to immobilized protein A (or G) avoids the need to use a secondary antibody to precipitate antigen-antibody complexes. Because of their broad specificity and ease of use, protein A– agarose and protein G–agarose (and related products) are the state-of-the-art reagents for the isolation of soluble antigen-antibody complexes in immunoprecipitation protocols. Recent progress in the field of antibody engineering (reviewed by Rapley, 1995; Irving et al., 1996) promises to make antibody production a less time-consuming and haphazard process. Antibody fragments with high affinity for specific antigens can now be selected from phage display antibody libraries. Selected recombinant antibodies can then be produced in large quantities in Escherichia coli. Techniques have been developed for producing antibodies in soluble, secreted form. Affinity tags are added to the recombinant antibody molecules to facilitate purification, detection, and use in procedures such as immunoprecipitation. While attractive in principle, the production of recombinant antibodies has been plagued by technical difficulties that so far have limited their widespread use in cell biology. However, as technical problems are overcome, recombinant techniques will progressively replace immunization of animals as a way of producing antibodies for immunoprecipitation and for other applications.

Critical Parameters Extraction of antigens Isolation of cellular antigens by immunoprecipitation requires extraction of the cells so that the antigens are available for binding to specific antibodies, and are in a physical form that allows separation from other cellular components. Extraction with nondenaturing deter-

gents such as Triton X-100 (see Basic Protocol 1 and Alternate Protocol 1) or in the absence of detergent (see Alternate Protocol 3) allows immunoprecipitation with antibodies to epitopes that are exposed on native proteins. Other nondenaturing detergents such as Nonidet P-40, CHAPS, digitonin, or octyl glucoside are also appropriate for extraction of native proteins (UNIT 5.1). Some of these detergents (e.g., digitonin) preserve weak protein-protein interactions better than Triton X-100. If the antigen is part of a complex that is insoluble in nondenaturing detergents (e.g., cytoskeletal structures, chromatin, membrane “rafts”) or if the epitope is hidden within the folded structure of the protein, extraction under denaturing conditions is indicated (see Alternate Protocol 2). The number of cells necessary to detect an immunoprecipitated antigen depends on the cellular abundance of the antigen and on the efficiency of radiolabeling. The protocols for radiolabeling (UNIT 7.1) and immunoprecipitation described in this book are appropriate for detection of antigens that are present at low to moderate levels (10,000 to 100,000 copies per cell), as is the case for most endogenous integral membrane proteins, signal transduction proteins, and transcription factors. For more abundant antigens, such as cytoskeletal and secretory proteins or proteins that are expressed by viral infection or transfection, the quantity of radiolabeled cells used in the immunoprecipitation can be reduced accordingly. Production of antibodies Immunoprecipitation can be carried out using either polyclonal or monoclonal antibodies (see discussion of selection below). Polyclonal antibodies are most often prepared by immunizing rabbits, although polyclonal antibodies produced in mice, guinea pigs, goats, sheep, and other animals, are also suitable for immunoprecipitation. Antigens used for polyclonal antibody production can be whole proteins purified from cells or tissues, or can be whole or partial proteins produced in bacteria or insect cells by recombinant DNA procedures. Another useful procedure is to immunize animals with peptides conjugated to a carrier protein. Production of polyclonal antibodies to recombinant proteins and peptides has become the most commonly used approach to obtain specific probes for immunoprecipitation and other immunochemical techniques, because it does not require purification of protein antigens from their native sources. The only requirement for making these antibodies is knowledge of the



sequence of a protein, information which is now relatively easy to obtain as a result of cDNA and genomic DNA sequencing projects. Polyclonal antibodies can be used for immunoprecipitation as whole serum, ammonium sulfate–precipitated immunoglobulin fractions, or affinity-purified immunoglobulins. Although all of these forms are suitable for immunoprecipitation, affinity-purified antibodies often give lower backgrounds and are more specific. Most monoclonal antibodies are produced in mice or rats. The sources of antigen for monoclonal antibody production are the same as those for production of polyclonal antibodies, namely, proteins isolated from cells or tissues, recombinant proteins or protein fragments, and peptides. A significant advantage of monoclonal antibodies is that antigens do not need to be purified to serve as immunogens, as long as the screening method is specific for the antigen. Another advantage is the unlimited supply of monoclonal antibodies afforded by the ability to grow hybridomas in culture or in ascitic fluid. Many monoclonal antibodies can now be produced from hybridomas deposited in cell banks or are directly available from companies. Ascitic fluid, cell culture supernatant, and purified antibodies are all suitable sources of monoclonal antibodies for immunoprecipitation. Ascitic fluid and purified antibodies should be used when a high antibody titer is important. Cell culture supernatants have lower antibody titers, but tend to give cleaner immunoprecipitations than ascitic fluids due to the lack of contaminating antibodies.

Immunoprecipitation

Selection of antibodies: Polyclonal versus monoclonal What type of antibody is best for immunoprecipitation? There is no simple answer to this question, as the outcome of both polyclonal and monoclonal antibody production protocols is still difficult to predict. Polyclonal antibodies to whole proteins (native or recombinant) have the advantage that they frequently recognize multiple epitopes on the target antigen, enabling them to generate large, multivalent immune complexes. Formation of these antigenantibody networks enhances the avidity of the interactions and increases the efficiency of immunoprecipitation. Because these antibodies recognize several epitopes, there is a better chance that at least one epitope will be exposed on the surface of a solubilized protein and thus be available for interaction with antibodies. Thus, the likelihood of success is higher. These properties can be a disadvantage, though, as

some polyvalent antibodies can cross-react with epitopes on other proteins, resulting in higher backgrounds and possible misidentification of antigens. By being directed to a short peptide sequence, anti-peptide polyclonal antibodies are less likely to cross-react with other proteins. However, their usefulness is dependent on whether the chosen sequence turns out to be a good immunogen in practice, as well as on whether this particular epitope is available for interaction with the antibody under the conditions used for immunoprecipitation. Unfractionated antisera are often suitable for immunoprecipitation. However, there is a risk that serum proteins other than the antibody will bind nonspecifically to the immunoadsorbent, and in turn bind proteins in the lysate that are unrelated to the antigen. For instance, transferrin can bind nonspecifically to immunoadsorbents, potentially leading to the isolation of the transferrin receptor as a contaminant (Harford, 1984). Polyclonal antisera can also contain antibodies to other antigens (e.g., viruses, bacteria) to which the animal may have been exposed, and these antibodies can also crossreact with cellular proteins during immunoprecipitation. Affinity-purified antibodies are a better alternative when antisera do not yield clean immunoprecipitations. Affinity-purification can lead to loss of high-affinity or low-affinity antibodies; however, the higher specificity of affinity-purified antibodies generally makes them “cleaner” reagents for immunoprecipitation. The specificity, high titer, and limitless supply of the best immunoprecipitating monoclonal antibodies are unmatched by those of polyclonal antibodies. However, not all monoclonal antibodies are useful for immunoprecipitation. Low-affinity monoclonal antibodies can perform acceptably in immunofluorescence microscopy protocols (UNIT 4.3) but may not be capable of holding on to the antigen during the repeated washes required in immunoprecipitation protocols. The use of ascitic fluid has the same potential pitfalls as the use of polyclonal antisera, as ascites may also contain endogenous antibodies to other antigens and proteins such as transferrin that can bind to other proteins in the lysate. In conclusion, an informed empirical approach is recommended in order to select the best antibody for immunoprecipitation. In general, it is advisable to generate and/or test several antibodies to a particular antigen in order to find at least one that will perform well in immunoprecipitation protocols.


Antibody titer The importance of using the right amount of antibody for immunoprecipitation cannot be overemphasized. This is especially the case for quantitative immunoprecipitation studies, in which the antibody should be in excess of the specific antigen. For instance, in pulse-chase analyses of protein degradation or secretion (UNIT 7.1), it is critical to use sufficient antibody to deplete the antigen from the cell lysate. This is particularly important for antigens that are expressed at high levels, a common occurrence with the growing use of high-yield protein expression systems such as vaccinia virus or replicating plasmids in COS cells. Consider for example a protein that is expressed at high levels inside the cell, and of which only a small fraction is secreted into the medium. If limiting amounts of antibody are used in a pulse-chase analysis of this protein, the proportion of protein secreted into the medium will be grossly overestimated, because the limiting antibody will bind only a small proportion of the cell-associated protein and a much higher proportion of the secreted protein. The same considerations apply to degradation studies. Thus, it is extremely important in quantitative studies to ensure that the antibody is in excess of the antigen in the cell samples. This can be ascertained by performing sequential immunoprecipitations of the samples (see Basic Protocol 1, annotation to step 21). If the second immunoprecipitation yields only a small amount of the antigen relative to that isolated in the first immunoprecipitation (10%, either more antibody or less antigen should be used. Too much antibody can also be a problem, as nonspecific immunoprecipitation tends to increase with increasing amounts of immunoglobulins bound to the beads. Thus, titration of the antibody used for immunoprecipitation is strongly advised. Immunoadsorbent If cost is not an overriding issue, the use of protein A– or protein G–agarose is recommended for routine immunoprecipitation. Protein A– or protein G–agarose beads (or equivalent products) have a very high capacity for antibody binding (up to 10 to 20 mg of antibody per milliliter of gel). Both protein A and protein G bind a wide range of immunoglobulins (Table 7.2.1). Backgrounds from nonspecifically bound proteins are generally low. Protein A–

and protein G–agarose beads are also stable and easy to sediment by low-speed centrifugation. A potential disadvantage, in addition to their cost, is that some polyclonal or monoclonal antibodies bind weakly or not at all to protein A or protein G (Table 7.2.1). This problem can be solved by using an intermediate rabbit antibody to the immunoglobulin of interest. For example, a goat polyclonal antibody can be indirectly bound to protein A–agarose by first incubating the protein A–agarose beads with a rabbit anti-goat immunoglobulin, and then incubating the beads with the goat polyclonal antibody. Anti-immunoglobulin antibodies (e.g., rabbit anti–goat immunoglobulins) coupled covalently to agarose can also be used for indirect immunoprecipitation in place of protein A– or protein G–agarose. Fixed Staphylococcus aureus particles (Pansorbin) can be used as a less expensive alternative to protein A–agarose. They have a lower capacity, can give higher backgrounds, and take longer to sediment. However, they work quite well in many cases. In order to establish if they are appropriate for a particular experimental setup, conduct a preliminary comparison of the efficiency of protein A–agarose with Staphylococcus aureus particles as immunoadsorbent. Specific antibodies coupled covalently to various affinity matrices can also be used for direct immunoprecipitation of antigens. After binding to protein A–agarose, antibodies can be cross-linked with dimethylpimelimidate (Gersten and Marchalonis, 1978). Purified antibodies can also be coupled directly to derivatized matrices such as CNBr-activated Sepharose (Springer, 1996). This latter approach avoids having to bind the antibody to protein A–agarose. Covalently bound antibodies should be used when elution of immunoglobulins from the beads complicates further analyses of the complexes. This is the case when proteins in immunoprecipitates are analyzed by one- or two-dimensional gel electrophoresis (UNIT 6.1) followed by Coomassie blue or silver staining, or are used for microsequencing. Also, the released immunoglobulins could interfere with detection of some antigens by immunoblotting (UNIT 6.2) following immunoprecipitation. Nonspecific controls For correct interpretation of immunoprecipitation results, it is critical to include appropriate nonspecific controls along with the specific samples. One type of control consists of setting up an incubation with an irrelevant antibody in the same biochemical form as the



experimental antibody (e.g., serum, ascites, affinity-purified immunoglobulin, antibody bound to protein A–agarose or directly conjugated to agarose), and belonging to the same species and immunoglobulin subclass as the experimental antibody (e.g., rabbit antiserum, mouse IgG2a). For an antiserum, the best control is preimmune serum (serum from the same animal obtained before immunization). Nonimmune serum from the same species is an acceptable substitute for preimmune serum in some cases. “No-antibody” controls are not appropriate because they do not account for nonspecific binding of proteins to immunoglobulins. In immunoprecipitation-recapture experiments, control immunoprecipitations with irrelevant antibodies should be performed for both the first and second immunoprecipitation steps (Fig. 7.2.3). Another type of control is to perform an immunoprecipitation from cells that do not express a specific antigen in parallel with immunoprecipitation of the antigen-expressing cells. For instance, untransfected cells are a perfect control for transfected cells. In yeast cells, null mutants that do not express a specific antigen are an ideal control for wild-type cells. Order of stages In the immunoprecipitation protocols described in this unit, the antibody is prebound to protein A–agarose before addition to the cell lysate containing the antigen. This differs from other protocols in which the free antibody is first added to the lysate and the antigen-antibody complexes are then collected by addition of the immunoadsorbent. Although both procedures can give good results, the authors prefer the protocols described here because this method allows better control of the amount of antibody bound to the immunoadsorbent. Prebinding antibodies to the immunoadsorbent beads allows removal of unbound antibodies. The presence of unbound antibodies in the incubation mixture could otherwise result in decreased recovery of the antigen on the immunoadsorbent beads. Another advantage of the prebinding procedure is that most proteins other than the immunoglobulin in the antibody sample (e.g., serum proteins) are removed from the beads and do not come in contact with the cell lysate. This eliminates potential adverse effects of these proteins on isolation of the antigen.

Immunoprecipitation

Washing The five washes described (see Basic Protocol 1; four with wash buffer and one with PBS)

are sufficient for maximal removal of unbound proteins; additional washes are unlikely to decrease the background any further. The last wash with PBS removes the Triton X-100 that can lead to decreased resolution on SDS-PAGE. It also removes other components of the wash buffer that could interfere with enzymatic treatment of immunoprecipitates. It is not advisable to complete all the washes quickly (e.g., in 5 min), because this may not allow enough time for included proteins to diffuse out of the gel matrix. Instead, beads should be washed over ∼30 min, which may require keeping the samples on ice for periods of 3 to 5 min between washes. In order to reduce nonspecific bands, samples can be subjected to an additional wash with wash buffer containing 0.1% (w/v) SDS, or with a mixture of 0.1% (w/v) SDS and 0.1% (w/v) sodium deoxycholate (Fig. 7.2.4). This wash should be done between the fourth wash and the wash with PBS.

Troubleshooting Two of the most common problems encountered in immunoprecipitation of metabolically labeled proteins are failure to detect specific antigens in the immunoprecipitates, and high background of nonspecifically bound proteins for antigens that were radiolabeled in vivo and analyzed by SDS-PAGE (UNIT 6.1) followed by autoradiography or fluorography (UNIT 6.3). When immunoprecipitates are analyzed by immunoblotting (UNIT 6.2), an additional problem may be the detection of immunoprecipitating antibody bands in the blots (Table 7.2.2).

Anticipated Results For antigens that are present at >10,000 copies per cell, the radiolabeling and immunoprecipitation protocols described in this book would be expected to result in the detection of one or more bands corresponding to the specific antigen and associated proteins in the electrophoretograms. Specific bands should not be present in control immunoprecipitations done with irrelevant antibodies. If antigens are labeled with [35S]methionine (UNIT 7.1), specific bands should be visible within 2 hr to 2 months of exposure. Due to the relatively low yield of the immunoprecipitation-recapture procedure (??@A@BC>DEF>GHEF@AH IJKLEMNMOIJKLENPQ

RSEES

RSEES TII

UV

WI

TT

KX K

T

W

U

Figure 7.2.4 Lowering background by washing with SDS and sodium deoxycholate (DOC). In this experiment, BW5147 cells (mouse thymoma) labeled with [35S]methionine for 1 hr were extracted with nondenaturing lysis buffer (see Basic Protocol 1). The extracts were subjected to immunoprecipitation with protein A–agarose beads incubated with either preimmune (PI) or immune (I) serum from a rabbit immunized with the ribosomal protein L17 (doublet at Mr ∼22,000). Lanes 1 and 2 correspond to immunoprecipitates obtained using the protocols described in this unit. Notice the presence of nonspecific bands and/or associated proteins in lane 2. Lanes 3 and 4 correspond to beads that were washed an additional time with 0.1% (w/v) SDS and 0.1% (w/v) DOC. Notice the disappearance of most of the nonspecific bands and/or associated proteins. The positions of Mr standards (expressed as 10−3 × Mr) are shown at left.



Table 7.2.2

Troubleshooting Guide for Immunoprecipitation

Problem

Cause

No specific radiolabeled antigen band Gel is completely blank after Poorly labeled cells: too little prolonged autoradiographic radiolabeled precursor, too few cells exposure labeled, lysis/loss of cells during labeling, too much cold amino acid in labeling mix, wrong labeling temperature Only nonspecific bands present Antigen does not contain the amino acid used for labeling Antigen expressed at very low levels

Solution Check incorporation of label by TCA precipitation (UNIT 7.1); troubleshoot the labeling procedure

Label cells with another radiolabeled amino acid, or for glycoproteins, with tritiated sugar Substitute cells known to express higher levels of antigens as detected by other methods; transfect cells for higher expression Use pulse labeling

Protein has high turnover rate and is not well labeled by long-term labeling Protein has a low turnover rate and is Use long-term labeling not well labeled by short-term labeling Protein is not extracted by lysis buffer Solublize with a different used to solubilize cells nondenaturing detergent or under denaturing conditions Antigen is not extracted with Triton Extract with Triton X-100 at 37°C or X-100 at 4°C use another detergent Antibody is nonprecipitating Identify and use antibody that precipitates antigen Epitope is not exposed in native Extract cells under denaturing antigen conditions Antibody does not recognize Extract cells under nondenaturing denatured antigen conditions Antibody does not bind to Use a different immunoadsorbent immunoadsorbent (Table 7.2.1); use intermediate antibody Antigen is degraded during Ensure that fresh protease inhibitors immunoprecipitation are present High background of nonspecific bands Isolated lanes on gel with high Random carryover of background detergent-insoluble proteins

High background in all lanes

Incomplete washing Poorly radiolabeled protein Incomplete removal of detergent-insoluble proteins Insufficient unlabeled protein to quench nonspecific binding

Remove supernatant immediately after centrifugation, leaving a small amount with pellet; if resuspension occurs, recentrifuge Cap tubes and invert several times during washes Optimize duration of labeling to maximize signal-to-noise Centrifuge lysate 1 hr at 100,000 × g Increase concentration of BSA continued


Table 7.2.2

Troubleshooting Guide for Immunoprecipitation, continued

Problem

Cause

Solution

Antibody contains aggregates

Antibody solution contains nonspecific antibodies

Too much antibody Incomplete preclearing

Nonspecifically immunoprecipitated proteins

Immunoprecipitating antibody detected in immunoblots Complete immunoglobulin or heavy Protein A conjugate or secondary and/or light chains visible in imantibody recognizes munoblot immunoprecipitating antibody

of an entire immunoprecipitation-recapture experiment requires a very long workday. Alternatively, samples can be frozen after the first immunoprecipitation, and the elution and recapture can be carried out another day.

Literature Cited Franzusoff, A., Rothblatt, J., and Schekman, R. 1991. Analysis of polypeptide transit through yeast secretory pathway. Methods Enzymol. 194:662-674. Gersten, D.M. and Marchalonis, J.J. 1978. A rapid, novel method for the solid-phase derivatization of IgG antibodies for immune-affinity chromatography. J. Immunol. Methods 24:305-309. Harford, J. 1984. An artefact explains the apparent association of the transferrin receptor with a ras gene product. Nature 311:673-675. Harlow, E. and Lane, D. 1988. Antibodies: A Laboratory Manual. Cold Spring Harbor Laboratory Press, Cold Spring Harbor, N.Y. Irving, R.A., Hudson, P.J., and Goding, J.W. 1996. Construction, screening and expression of re-

Microcentrifuge antibody 15 min at maximum speed before binding to beads Use affinity-purified antibodies; absorb antibody with acetone extract of cultured cells that do not express antigen; for yeast cells, absorb antibody with null mutant cells Use less antibody Preclear with irrelevant antibody of same species of origin and immunoglobulin subclass bound to immunoadsorbent Fractionate cell lysate (e.g, ammonium sulfate precipitation, lectin absorption, or gel filtration) prior to immunoprecipitation; after washes in wash buffer, wash beads once with 0.1% SDS in wash buffer or 0.1% SDS/0.1% sodium deoxycholate Use antibody coupled covalently to solid-phase matrix for immunoprecipitation; probe blots with primary antibody from a different species and the appropriate secondary antibody specific for immunoblotting primary antibody

combinant antibodies. In Monoclonal Antibodies: Principles and Practice (J.W. Goding, ed.). Academic Press, London. Köhler, G. and Milstein, C. 1975. Continuous cultures of fused cells secreting antibody of predetermined specificity. Nature 256:495-497. Nisonoff, A. 1984. Introduction to Molecular Immunology. Sinauer Associates, Sunderland, Mass. Rapley, R. 1995. The biotechnology and applications of antibody engineering. Mol. Biotech. 3:139-154. Springer, T.A. 1996. Immunoprecipitation. In Current Protocols in Immunology (J.E. Coligan, A.M. Kruisbeck, D.H. Margulies, E.M. Shevach, and W. Strober, eds.) pp. 8.3.1-8.3.11. John Wiley & Sons, New York.

Contributed by Juan S. Bonifacino and Esteban C. Dell’Angelica National Institute of Child Health and Human Development Bethesda, Maryland



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ABTS reagent ABTS buffer: 0.05 M Na2HPO4 0.1 M sodium acetate Adjust pH to 5.0 using concentrated HCl Store up to 3 months at room temperature ABTS reagent: Immediately before assay, dissolve 11 mg of 2,2′-azinobis(3-ethylbenzthiazoline)sulfonic acid (ABTS; Sigma) in 0.5 ml water. Mix 67 µl of 30% H2O2 with 7 ml water. Add 0.5 ml of this ABTS solution to 10 ml ABTS buffer (see above) and 100 µl H2O2 solution. Mix thoroughly. This amount of reagent is sufficient for one full 96-well plate assay.

Binding buffer Mix the following components in the order indicated: 150 mM NaCl 25 mM Tris⋅Cl, pH 7.4 (APPENDIX 2A) 1 mM MnCl2 0.1% (w/v) BSA (fraction V; Sigma, 99% pure) Prepare fresh This is conveniently prepared from Tris/saline [25 mM Tris⋅Cl (pH 7.4)/150 mM NaCl] and a stock solution of 1 M MnCl2. BSA is then added.

Cell Adhesion


Supplement 15

Blocking solution Prepare a solution of 25 mM Tris⋅Cl containing 150 mM NaCl. Add sufficient 20% sodium azide stock for a final concentration of 0.05% (w/v). Add BSA (fraction V; Sigma, 98% pure) for a final concentration of 5% (w/v) and dissolve by vigorous stirring. Centrifuge the solution in 50-ml centrifuge tubes for 5 min at 2800 × g (4000 rpm in a JA-10 rotor), and filter the supernatant through a 20-ml disposable column. Store up to 3 months at 4°C. This is conveniently prepared from Tris-saline [25 mM Tris⋅Cl (pH 7.4)/150 mM NaCl] to which sodium azide is added from a 20% stock solution. BSA is then added and dissolved by vigorous stirring. The solution is then centrifuged and filtered as above. Final concentrations are 150 mM NaCl; 25 mM Tris⋅Cl, pH 7.4, 5% (w/v) BSA, and 0.05% (w/v) sodium azide.

Elution buffer 10 mM sodium acetate, pH 3.25 1 mM CaCl2 1 mM MgCl2 0.1% (w/v) Triton X-100 (Sigma, Ultra grade) Store up to 1 month at 4°C Extraction buffer 150 mM NaCl 25 mM Tris⋅Cl, pH 7.4 (APPENDIX 2A) 2% (w/v) Triton X-100 (Sigma, Ultra grade) 1 mM PMSF (100 mM stock solution prepared in isopropanol; also see APPENDIX 1B) 10 µg/ml leupeptin (1 mg/ml stock solution prepared in water; also see APPENDIX 1B) 2 mg/ml BSA (fraction V; Sigma, 98% pure) Prepare fresh, then cool on ice Homogenization buffer 150 mM NaCl 25 mM Tris⋅Cl, pH 7.4 (APPENDIX 2A) 0.005% (w/v) digitonin Store for up to 3 months at 4°C SDS-PAGE sample buffer, 5× 25% (v/v) glycerol 125 mM Tris⋅Cl, pH 6.8 (APPENDIX 2A) 10% (w/v) SDS 0.1% (w/v) bromophenol blue Store indefinitely at 4°C. Warm in a hot water bath and mix well before use. Wash buffer 150 mM NaCl 25 mM Tris⋅Cl, pH 7.4 (APPENDIX 2A) 1 mM CaCl2 (add from 1 M stock) 1 mM MgCl2 (add from 1 M stock) 0.1% (w/v) Triton X-100 (Sigma, Ultra grade) Store up to 1 month at 4°C Analyzing IntegrinDependent Adhesion

This is conveniently prepared from Tris-saline [25 mM Tris⋅Cl (pH 7.4)/150 mM NaCl] and stock solutions of 1 M CaCl2 and 1 M MgCl2.

9.4.12 Supplement 15


COMMENTARY Background Information Monoclonal antibodies were of crucial importance in the initial identification of cell surface receptors that mediated adhesion of cells to extracellular matrix components (e.g., Wayner and Carter, 1987). A wide range of well characterized anti-integrin mAbs are now commercially available for use in cell attachment and spreading assays. The choice of which anti-integrin mAbs to use is determined in part by the substrate. For example, if collagen type I is the substrate, anti-α1 and anti-α2 mAbs should be tested, as these are the likely integrins involved. The complement of integrins expressed by the cell also helps to determine which mAbs should be tested. If the profile of integrin expression is unknown, it can be determined by flow cytometry or by immunoprecipitation of surface-labeled cells. Conversely, if a mAb blocks cell adhesion, it is important to demonstrate that the corresponding integrin subunit is expressed by the cells. Historically, the first purification of an integrin receptor was a major advance in understanding the molecular basis of cell-matrix interactions (Pytela et al., 1985). This and later methods employed a ligand affinity column as the major purification procedure. In particular, fibronectin fragments have been used to purify α5β1, and RGD peptides have been used to purify αVβ3 and αIIbβ3 (Pytela et al., 1987; Smith and Cheresh, 1988; Yamada and Yamada, 1990). Ligand affinity columns remain the method of choice where mAbs are unavailable. However, the use of mAbs has the advantage that the purification is more specific (e.g., several different integrins bind to fibronectin affinity columns) and higher yields of integrins can be obtained. Multiple β1 integrins can also be obtained from the same source. For example, other β1 integrins from human placenta that are present in the flowthrough from the mAb 16 column can be purified using other specific anti-α mAbs. For some assays, and particularly where one β1 integrin predominates in the tissue or cell extract, it may be sufficient to purify the total β1 integrins and use this partially purified preparation in the solid-phase assay. For example, β1 integrins purified from MOLT-4 cells (Newham et al., 1998) contain ∼75% α4β1, the remainder being α5β1. The protocol described here can be adapted to purify other integrins from other tissues or from pellets of cultured cells.

The major advantage of assays using purified integrins, as compared to cell-based assays, is that integrin-ligand binding can be studied in isolation. Integrin clustering, signaling, and cytoskeletal interactions are all known to affect the strength of adhesion in cell-based assays. Furthermore, adhesion may be modulated by indirect effects (e.g., by signaling from other cell-surface receptors). Although doubts are often expressed as to whether plastic-adsorbed integrin is representative of integrin in its native environment, a number of careful studies have shown no significant differences between the behavior of integrins in solid-phase assays and on cell surfaces. Hence, this approach has been broadly validated. The first solid-phase integrin-ligand binding assay was described by Charo et al. (1991) for studying fibrinogen binding to αIIbβ3. The α5β1-fibronectin assay developed by the author of this unit is both extremely sensitive and highly versatile. For example, the author has described how the assay can be used to investigate the effects on ligand binding of divalent cations, activating and inhibitory mAbs, peptide inhibitors, and mutations (Mould et al., 1995a,b, 1996, 1997). Another important area in which this type of assay is finding use is in the pharmacological screening of inhibitors of integrin-ligand interactions. This assay can give information about the inhibitory potency of a compound and whether it is a direct competitive or allosteric inhibitor of ligand binding. The attenuation of mAb epitopes can also provide data on the location of the binding site of an inhibitor on the integrin (Mould et al., 1997). The author’s preferred method for labeling of integrin ligands is biotinylation, because of its safety and simplicity. One potential drawback is that if one or more lysyl residues in the ligand are crucial for integrin binding, their modification may render the ligand inactive. In this case, a possible solution may be to reduce the amount of biotinylation reagent so that some of the lysyl residues remain unmodified. Other labeling methods such as radioiodination can also be used. Alternatively, if the ligand is a recombinant protein, a “tag” such as an epitope sequence or the Fc region of IgG can be incorporated for use in the detection of bound ligand.

Cell Adhesion


Supplement 15

Table 9.4.2

Troubleshooting Guide for Problems Encountered in Solid-Phase Assays

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Possible cause

Solution

High background binding to BSA- Insufficient blocking of wells coated wells Ligand concentration too high

Block for longer time (e.g., overnight) Test range of ligand concentrations for optimal signal/background

Spuriously high signal in some wells Wide variation in signal in experimental wells

Follow washing protocol carefully Add reagents to the center of wells Mix diluted integrin thoroughly before coating the plate Add blocking reagent before removing coating solution Use fresh, clean plates Use lower dilution of integrin Use higher concentration of ligand Check activity of ligand in cell-based assay

Low signal above background binding to BSA

Insufficient washing Tops of wells contaminated Integrin added to plate insufficiently mixed Wells aspirated before adding blocking reagent Plate contaminated, e.g., by dust Insufficient integrin Ligand concentration too low Inactive ligand

Critical Parameters and Troubleshooting For cell-based assays, as described in UNIT the health of the cells and careful preparation are important for achieving optimal spreading or attachment. In order to optimize the sensitivity of cell attachment or spreading to inhibition, the author recommends that a concentration of substrate be chosen that gives 50% to 70% of maximal adhesion (unless this value is low, in which case a concentration of substrate that gives near maximal adhesion should be used). If the level of adhesion is near maximal, inhibitors are less effective at blocking adhesion. Failure of a mAb to inhibit spreading or attachment can normally be taken to mean that the integrin it recognizes is not involved in adhesion. However, it is important to check that the antibody is effective in a system where inhibition should be observed (e.g., an anti-α5 mAb should block spreading of HT-1080 fibrosarcoma cells on fibronectin). Conversely, if an antibody does inhibit, it is important to test that it does not inhibit adhesion to an inappropriate ligand (e.g., an anti-α5 mAb should not perturb HT-1080 cell spreading on collagen). Most of the antibodies described in Table 9.4.1 are well characterized and should not show any non-specific effects. However, as described earlier, it is essential that the antibodies not contain sodium azide. It is also important to bear in mind that inhibition of adhesion can be caused by “crosstalk”—i.e., where ligation of one integrin (e.g., by peptide) indirectly affects the activity of a

9.1,

Analyzing IntegrinDependent Adhesion

second integrin via intracellular signaling (Diaz-Gonzalez et al., 1996). While it is often difficult to rule out such effects, they normally only cause a partial reduction in adhesion, whereas adhesion is frequently totally ablated by specific antibody or peptide inhibition. Finally, if anti-integrin mAbs or peptides do not completely block cell adhesion, it is possible that non-integrin receptors may play some role. This may be observed particularly in cell attachment assays. For example, cell-surface proteoglycans contribute to melanoma cell attachment to the heparin-binding domain of fibronectin (Mould et al., 1994). For solid-phase assays, the specificity of the assay must be tested carefully. The most important test for specificity is the ability of unlabeled ligand to compete with labeled ligand for binding to the integrin. Hence, in the presence of a large excess of unlabeled ligand, very little binding of labeled ligand should be observed. Nearly all integrin-ligand interactions are divalent-cation dependent. Hence, replacing the Mn2+ in the binding buffer with EDTA should reduce binding to levels similar to that observed for BSA-coated wells. Further tests for specificity can be carried out. For example: (1) mAbs that are inhibitory in cell-based assays should also inhibit ligand binding in solid-phase assays, (2) mutations known to affect integrin binding sites should perturb ligand recognition, and (3) known ligand mimetics (e.g., RGD peptides for α5β1 or CS1 peptide for α4β1) should block ligand binding. All of these tests have been performed for the α5β1-fibronectin



Table 9.4.3

Troubleshooting Guide for Problems Encountered in Integrin Purification

Problem

Possible cause

Solution

Large number of proteins copurify Inadequate preclearing or filtration of with integrins extract, or precipitate forms during purification procedures Small amounts of integrins Insufficient mAb coupled to Sepharose purified Affinity of mAb too low Column has been used many times

Recentrifuge after preclearing and filtering extract on Sepharose 4B, or when any precipitate is visible Couple more mAb to Sepharose Use mAb with higher affinity Replace with fresh mAb-Sepharose

Integrin degraded

Increase levels of protease inhibitors and BSA in extraction buffer Perform all manipulations at 4°C or on ice

Insufficient levels of protease inhibitors in extraction buffer Extraction or other manipulations performed at too high a temperature

assay. Table 9.4.2 is a troubleshooting guide for solid-phase ligand-binding assays. For integrin purification, it is essential to have sufficient tissue or cell pellets for the extraction and enough mAb-Sepharose to perform a successful purification. Table 9.4.3 is a troubleshooting guide for integrin purification.

Anticipated Results For cell-based assays, in most cases, it is possible to obtain levels of spreading or attachment of >50%. It should be feasible using anti-integrin mAbs, either alone or in combination, to reduce the level of spreading or attachment to close to that seen on BSA. For integrin purification, yields of α5β1 are ∼2 mg per placenta (estimated by Coomassie blue staining). Each full 96-well-plate solidphase assay uses 100 proteins involved in cell-cell adhesion. Its members are found in vertebrates, invertebrates, and also in yeast. Most of the molecules of the IgSF are cell surface molecules that are membrane-anchored either by a single transmembrane segment or by a glycosylphosphatidylinositol (GPI) anchor that is posttranslationally attached to the C-terminus. Some of the IgSF-CAMs also occur in soluble form, e.g., in the cerebrospinal fluid or the vitreous fluid of the eye, due to a cleavage of the GPI-anchor or the membrane-proximal peptide segment. In some cases, such as NCAM, various forms may be generated by alternative splicing. This unit provides protocols for the purification of IgSF-CAMs from tissue extracts and tissue culture supernatants and for the analysis of the adhesive functions of IgSF-CAMs with isolated molecules and in the cellular context. Following personal expertise, the authors have added a few frequently used functional assays demonstrating the role of IgSF-CAMs in neural development, such as neurite outgrowth from cultured neurons, and the use of antibodies for the inhibition of IgSF-CAM functions in vitro. The first group of protocols describe affinity purification of IgSF-CAMs (see Basic Protocol 1), preparation of the affinity column (see Support Protocol 1), solubilization of membrane proteins (see Support Protocol 2), transient transfection of HEK 293 cells to express IgSF-CAMs (see Support Protocol 3), and detection of IgSF-CAMs by dot blot analysis (see Support Protocol 4). Assays using fluorescent microspheres with coupled proteins are used for one type of functional analysis based either on interactions between microspheres (see Basic Protocol 2) or on interactions between microspheres and cultured cells (see Basic Protocol 3). There are two protocols for coupling proteins to microspheres: coupling proteins to fluorescent microspheres (see Support Protocol 5) and coupling proteins to glutaraldehyde-activated amino beads (see Support Protocol 6). A second group of protocols analyze the functions in cell-based assays. Trans-interactions are studied using IgSF-CAM-transfected myeloma cells (see Basic Protocol 4). This protocol requires stable transfection of myeloma cells (see Support Protocol 7). Cis-interactions are detected by chemical cross-linking (see Basic Protocol 5) and antibody co-capping (see Basic Protocol 6). IgSF-CAMs and other substrates have the ability to promote neurite outgrowth (see Basic Protocol 7), which requires coating of the growth surface with IgSF-CAM (see Support Protocol 8), nitrocellulose as a binder for the substrate of interest (see Support Protocol 9), poly-D-lysine (see Support Protocol 10), collagen (see Support Protocol 11), or laminin (see Support Protocol 12). Differential fixation protocols are used for fluorescent immunohistochemistry samples (see Support Protocol 13) or for morphological analyses (see Support Protocol 14). Finally, there is a protocol for assessing the effect of inhibiting CAM-CAM interactions in vitro (see Basic Protocol 8).

Cell Adhesion Contributed by Peter Sonderegger, Stefan Kunz, Christoph Rader, Daniel M. Suter, and Esther T. Stoeckli

9.5.1

Current Protocols in Cell Biology (2001) 9.5.1-9.5.52 Copyright © 2001 by John Wiley & Sons, Inc.

Supplement 11

PREPARATION OF IgSF-CAMs BASIC PROTOCOL 1

Purification of IgSF-CAMs by Immunoaffinity Chromatography The best method for the purification of native, functionally intact proteins is certainly the use of standard chromatography, such as ion exchange, hydrophobic interaction, and gel permeation columns. However, the establishment of such a standard purification protocol can be time-consuming and requires expensive equipment. As the specific characteristics exploited for chromatography differ from protein to protein, purification protocols cannot be generalized. A faster and less expensive way to purify IgSF-CAMs is to use affinity chromatography. Tissue homogenates, body fluids in the case of secreted proteins, or cell lines engineered to express a particular protein of interest, either transiently or stably, can be used as a source for IgSF-CAM purification. Affinity chromatography makes use of the specific binding properties of the proteins, e.g., receptors for their ligands, enzymes for their substrates, or antibodies for their antigens. For IgSF-CAMs, the purification by immunoaffinity is most commonly used. A disadvantage of affinity purification is the possibility of the loss of activity, as the protein is sometimes eluted from the column by rather harsh conditions. The protocol described below has been successfully used for the purification of functionally intact IgSF-CAMs (Stoeckli et al., 1991, 1996; Rader et al., 1993). The general principle of immunoaffinity chromatography is the use of a resin-coupled monoclonal antibody directed against the protein to be purified. Generally, activated Sepharose resins are used. The resin is packed into a column connected to a peristaltic pump and to a UV-detector to monitor the elution profile of the column. The purity of the eluted protein is analyzed by SDS-PAGE (UNIT 6.1). Here, we describe a purification protocol for a membrane-bound IgSF from brain membranes (see Support Protocol 2 for membrane preparation and protein solubilization). Materials CNBr-activated Sepharose 4B column (see Support Protocol 1) Loading buffer: 0.5% CHAPS in PBS with Ca2+/Mg2+ Elution buffer: 0.5% CHAPS in 50 mM diethylamine Protein solution (see Support Protocol 2) 1 M Tris⋅Cl, pH 7.0 (APPENDIX 2A) PBS with Ca2+/Mg2+ (see recipe) 0.02% (v/v) merthiolate or equivalent bacteriostatic agent Prepare column 1. Rinse the column extensively with loading buffer, especially for a column that was prepared earlier and has been stored for a while. Add 2 to 3 vol (i.e., 3 times the volume of the column) of elution buffer to the column to test the stability of the column under the elution conditions and to make sure that the column does not contain any contaminations, such as unspecifically bound proteins from previous use of the column. 2. Properly reequilibrate the column to loading conditions before loading the protein solution (see Support Protocol 2 for protein solution preparation).

Cell-Cell Contact by Ig Superfamily Cell Adhesion Molecules

As the binding affinity of antibodies is usually not temperature-sensitive, the authors recommend running the affinity column at 4°C rather than at room temperature. Keeping the column and the protein solution to be loaded at 4°C helps to prevent contamination and slows down degradation of the proteins.

9.5.2 Supplement 11


Load the column 3. During loading of the protein solution onto the column, adjust the flow rate to a sufficiently slow rate (usually 0.1 ml/min is chosen for good yields) to allow good interaction between antigen and antibodies coupled to the sepharose beads. The capacity of the column can be tested by collecting fractions of the flow-through and subsequent immunoblot analysis (see Support Protocol 4). For best yield, the capacity of the column should not be exhausted. Typically, column volumes of 1 ml are used.

Elute column 4. Incubate the column in 0.9 vol elution buffer for 10 min. This will increase the elution efficiency (a higher concentration of the eluted protein in eluate, rather than a broad elution peak with a long tail). Do not expose the column to elution conditions for longer times than necessary, because the high pH of the buffer could be detrimental for the affinity column.

5. Elute the column at a rate of 1 ml/min. Collect the eluate in vials containing enough 1 M Tris⋅Cl, pH 7.0, to buffer the eluate at a neutral pH value. For a 1-ml column, 1-ml fractions are collected in 1.5-ml microcentrifuge tubes. The volume of 1 M Tris⋅Cl, pH 7.0, required for restoring the pH should be determined at 4°C, as the pH value of Tris is extremely temperature sensitive. The flow rate for elution can be much higher than that for loading. However, check the maximal flow rate acceptable for a specific resin. For Sepharose resins a maximal flow rate of 30 ml hr−1cm−2 is recommended. The eluate can be stored a few days at 4°C; for longer storage below −20°C is recommended. However, keep in mind that repeated thawing and freezing is detrimental to the protein. Furthermore, freezing of dilute protein solutions is not recommended.

6. Re-equilibrate the column to loading conditions for a second run, or prepare the column for storage. Regenerate and store the affinity column 7. Immediately after elution re-equilibrate the column to neutral pH values with PBS with Ca2+/Mg2+. For storage, add 0.02% merthiolate or an equivalent bacteriostatic agent to the PBS to prevent bacterial growth. Store the column at 4°C. Prevent drying of the column during storage. The column can be stored for several months at 4°C.

Preparation of the Affinity Column This protocol only describes the preparation of an immunoaffinity column. Generally, because IgSF-CAMs have low binding affinities for their binding partners and have no enzymatic activity that could be used for substrate-based purification, the use of immunoaffinity columns is the method of choice.

SUPPORT PROTOCOL 1

However, a prerequisite is the availability of a monoclonal antibody against the protein to be purified. This antibody is covalently coupled to a Sepharose resin. Affinity columns are versatile, they can be used for tissue homogenates, solubilized membrane proteins, or culture supernatants from cell lines that are engineered to produce and release IgSFCAMs. If stored appropriately, affinity columns can be reused many times over several months. Cell Adhesion


Supplement 11

Materials CNBr-activated Sepharose 4B gel 1 mM HCl Buffer I: 0.5 M NaCl in 0.1 M NaHCO3, pH 8.3 Monoclonal antibody against the protein to be purified 0.2 M glycine, pH 8.0 Buffer II: 0.5 M NaCl in 0.1 M sodium acetate, pH 4.0 Loading buffer (see Basic Protocol 1) Sintered glass filter connected to a vacuum pump Column (e.g., Poly-Prep column, Bio-Rad) U-bottomed polypropylene vial that can be closed tightly 1. Soak 1:2 (w/v) CNBr-activated sepharose 4B in 1 mM HCl for 15 min at room temperature. For a 1-ml column, start with 350 to 400 mg Sepharose.

2. Transfer beads to a sintered glass filter connected to a vacuum pump, and wash with ≥25 vol of 1 mM HCl, followed by buffer I. It is very important to prevent the beads from drying between the additions of buffer (one gram dry resin yields ∼3.5 ml swollen gel).

3. Transfer the slurry to a U-bottomed polypropylene vial containing the antibody solution in buffer I. Carry out the reaction for 2 hr at room temperature. Close the vial tightly. The final concentrations should be: 100 mg Sepharose (dry weight) and 5 mg antibody per ml coupling reaction mix. Ideally, rotating the vial end-over-head is used to maximize the coupling efficiency. Do not use a magnetic stirrer, which will damage the agarose beads.

4. Stop the reaction by gently centrifuging the Sepharose beads for 5 min at 2000 × g, room temperature. 5. Add 3 vol 0.2 M glycine, pH 8.0, to the pellet and continue to rotate the vial end-over-head for an additional 2 hr. Collect the supernatant of the coupling reaction to check the coupling efficiency.

6. Pack the slurry into a column. Typically, column volumes are ∼1 ml with a column diameter of 0.5 cm.

7. Wash the column with 5 vol buffer I followed by 5 vol buffer II. Repeat wash procedure four times to remove excess uncoupled ligand. 8. Before loading the protein solution (see Support Protocol 2), wash the column thoroughly with 25 to 30 vol loading buffer. An affinity column can be used repeatedly, if stored appropriately with an antibacterial agent (e.g., 0.02% merthiolate) at 4°C. Before using the column after storage, rinse the column extensively with loading buffer. Use 2 to 3 vol elution buffer to clean the column, restore loading conditions by rinsing thoroughly with loading buffer (≥10 vol).


9.5.4 Supplement 11


Solubilization of Membrane Proteins Most IgSF-CAMs are either glycosyl-phosphatidylinositol-anchored or transmembrane proteins, therefore, they have to be solubilized from cell membranes with detergents. In the authors’ experience, the use of CHAPS has given the best results with respect to yield and functional integrity of the purified proteins. Keep in mind that for many functional assays, detergent removal from the protein solution is necessary after purification; the presence of detergents can also interfere with binding assays. Especially sensitive assays are those that involve neurons, such as neurite outgrowth assays (see Basic Protocol 7). For removal of detergents from protein solutions, the authors have used SM-2 beads from BioRad or Calbiosorb from Calbiochem. The use of high concentrations of detergents can interfere with the purification by immunoaffinity columns, therefore, dilution of the protein solution after the solubilization step, i.e., before the solution is loaded onto the affinity column, is recommended. The protocol given below has been successfully used for the purification of functionally intact L1/NgCAM from E14 chicken brain membranes (e.g., Stoeckli et al., 1991, 1996). It is adapted from the purification protocol described by Grumet and Edelman (1984). However, the authors have used the same protocol for the solubilization and purification of other IgSF-CAMs (Rader et al., 1993; Fitzli et al., 2000). For storage, membranes and proteins can be frozen at the indicated steps. However, repeated freez-thaw cycles are detrimental for proteins and for high yields of intact proteins and should be minimized.

SUPPORT PROTOCOL 2

NOTE: All protocols using live animals must first be reviewed and approved by an Institutional Animal Care and Use Committee (IACUC) or must conform to governmental regulations regarding the care and use of laboratory animals. Materials 14-day-old chicken embryo brains, freshly frozen in liquid nitrogen Liquid nitrogen Ca2+/Mg2+-free buffer (CMF buffer; see recipe) 0.8 M and 2.25 M sucrose in PBS 1 M and 2 M NaCl in PBS 50 mM triethylamine 0.5% and 1% CHAPS in PBS Mortar and pestle Dounce homogenizer Centrifuge tubes for Sorval SS-34 or equivalent rotor 38-ml polycarbonate tubes for ultra high-speed centrifuge Prepare membranes 1. Remove brains of 14-day-old chicken embryos and immediately freeze in liquid nitrogen. If necessary, brains can be stored at −70°C for extended periods of time.

2. Cool a mortar and pestle of sufficient size with liquid nitrogen and grind frozen brains in batches to a fine powder. Add small volumes of liquid nitrogen to keep brains/brain powder frozen during grinding. Carefully avoid thawing of the tissue at any time.

3. Add ∼3 vol CMF buffer to 1 vol brain powder. Homogenize the brain powder in a Dounce homogenizer. 4. Centrifuge homogenate for 20 min at 45,000 × g (10,000 rpm in a SS-34 rotor), 4°C. Cell Adhesion


Supplement 11

5. Resuspend the pellets in 1.5 to 2 vol of 2.25 M sucrose, use Dounce homogenizer to get homogenous suspension. 6. Transfer ∼24 ml suspension to each polycarbonate ultracentrifuge tube, overlay with 1⁄ vol of 0.8 M sucrose. 4 7. Centrifuge for 60 min at 150,000 × g, 4°C. 8. Transfer the membranes that are accumulated in the interphase to the Dounce homogenizer, resuspend them in 25 vol PBS. 9. Centrifuge for 60 min at 150,000 × g, 4°C. 10. Decant supernatant and resuspend in PBS, centrifuge as in step 9, and resuspend pellets in smallest possible volume of PBS for storage at −20°C, or use directly for stripping. Samples can be stored up to 1 month at −20°C, or at −70°C for longer storage.

Strip packed membranes 11. Add 1 vol of 2 M NaCl in PBS to packed membrane suspension (from step 10), homogenize suspension. 12. Add 1 M NaCl in PBS to a final volume of 4 to 5 times the volume of the packed membranes. 13. Stir suspension 1 hr on a magnetic stirrer at low speed, 4°C, to strip membranes from peripheral membrane proteins. 14. Centrifuge for 60 min at 150,000 × g, 4°C. 15. Resuspend pellets in 20 vol of 50 mM triethylamine, stir for 60 min at 4°C. 16. Repeat centrifugation step 14. Carefully remove the supernatant with a pipet, as the membranes do not form a stable pellet after the high pH extraction step. 17. Resuspend the pellets in PBS. 18. Centrifuge for 60 min at 150,000 × g, 4°C. 19. Repeat washing the membranes with PBS at least one time to restore a pH value between 7.2 and 7.6. Keep an aliquot for measuring the protein concentration. 20. Freeze the stripped membranes at −20°C or use directly for solubilization step. For storage longer than 1 month, store at −70°C.

Solubilize integral membrane proteins 21. If frozen membranes are used, they should be washed once again with PBS (steps 17 and 18). 22. Transfer pellets into a Dounce homogenizer and resuspend in 1 vol of 1% CHAPS buffer. Add 4 vol of 0.5% CHAPS buffer. For good solubilization the protein concentration should be adjusted to ∼1 mg/ml.

23. Extract the membrane proteins by stirring the suspension for 60 min at 4°C. 24. Centrifuge for 60 min at 150,000 × g , 4°C. Cell-Cell Contact by Ig Superfamily Cell Adhesion Molecules

25. Combine supernatants, measure the volume, and remove aliquot to determine the protein concentration. Do not freeze the solubilized proteins, but use directly for affinity purification step.

9.5.6 Supplement 11


Production of Recombinant CAM by Transient Transfection of HEK293 Cells with Calcium Phosphate

SUPPORT PROTOCOL 3

Transient transfection of HEK 293 cells with expression vectors containing cDNAs of IgSF-CAMs represents a fast and convenient method for the production of intermediate amounts (100 µg to 1 mg) of recombinant protein. The human embryonic kidney cell line 293 (HEK 293) is a well established, easily transfectable cell line that is widely used for the expression of recombinant proteins. The transfection method of choice with HEK 293 cells is always calcium phosphate transfection, an inexpensive, convenient technique that results in high efficiencies of transfection. DNA can be introduced into a wide variety of cultured cell lines as a calcium phosphate complex (Graham and van der Eb, 1973; Wigler et al., 1977). The transfected DNA can either integrate into the genome of the recipient cell, resulting in stable transgene expression accompanied by a stably altered phenotype of the cell (stable transfection) or remain episomal resulting in only transient expression of the transgene (transient transfection). The following protocol can be used likewise for the generation of stable cell lines by subsequent selection for stable transfectants or for transient expression only. Efficient transfection requires the formation of a fine precipitate of calcium phosphate in the presence of DNA. The formation of the DNA-containing calcium phosphate particles is initiated under defined chemical conditions, in the absence of cells or serum. The particle size is the most critical parameter regarding efficiency of transfection, that is uptake of DNA-containing calcium phosphate particles by the cells. The main determinants of particle size are calcium and phosphate concentrations, the concentration of DNA, size of DNA fragments involved, pH, temperature, and time of incubation. After initial formation of the DNA-containing calcium phosphate particles, the precipitate is added to the cells. During the incubation of the precipitate with the cells, the formation of DNA-containing calcium phosphate particles continues and preexisting particles grow in size. The particles adhere to the cells and are taken up by endocytosis. After a few hours of exposure, the medium is changed and the cells start to express the recombinant protein. The period of efficient transgene expression varies between different expression vectors but lasts generally for a few days. Materials HEK 293 cells Cell culture medium for HEK 293 cells (see recipe) Purified DNA of interest CaCl2 solution (see recipe) HBS solution (see recipe) 175-cm2 tissue culture flasks Additional reagents and equipment for trypsinizing cells (UNIT 1.1) NOTE: All solutions and equipment coming into contact with living cells must be sterile, and aseptic technique should be used accordingly. NOTE: All cell culture incubations should be carried out in a 37°C, 5% CO2 humidified incubator. 1. For transient transfection, grow HEK 293 cells of low passage number (passage 30 kb), preincubation of no longer than 1 min is recommended.

5. Incubate the DNA-containing calcium phosphate particles with the cells for 4 to 6 hr at 37°C in an incubator. The formation of the calcium phosphate precipitate can be monitored under an inverted light microscope with a 63× objective. The precipitate should become visible as tiny particles (∼300 nm in diameter) especially on the surface between the cells. However, long exposure to the atmosphere outside of the incubator should be avoided.

6. Change medium after the incubation period and continue incubation. Begin testing for expression of the recombinant protein 1 day after transfection. The cells should now express the recombinant protein for a few days. Uptake of the DNA-containing calcium phosphate particles by cells can be checked by examination of free surfaces between cells. The margins surrounding the cells should be cleared of particles, due to their uptake by the cells. SUPPORT PROTOCOL 4


Detection of IgSF-CAMs by the Dot Immunoblot Method A rapid and convenient method for the detection of IgSF-CAMs in eluates from chromatography or affinity columns are dot immunoblots. Similar to immunoblots, dot blots are a semi-quantitative method to detect proteins transferred onto a nitrocellulose membrane via a chromogenic reaction. For this purpose, secondary antibodies coupled to peroxidase or alkaline phosphatase are used to detect binding of the first antibodies directed against the protein of interest. However, in contrast to immunoblots, proteins are not separated on a polyacrylamide gel, therefore dot blots are a much faster way to demonstrate the presence of a specific protein in a given sample. Furthermore, as the proteins are directly applied to the nitrocellulose membrane for dot blots, there is no loss due to difficulties in

9.5.8 Supplement 11


transfer from the gel to the membrane. The fact that only small volumes can be applied onto the membrane for dot blots may, however, limit its use for detection of proteins in dilute solutions. The procedure described here can also be used to stain IgSF-CAMs after immunoblotting. Use manufacturer’s manuals and protocols for sample preparation, SDS-PAGE, and blotting of proteins onto nitrocellulose (also see UNITS 6.1 and 6.2). Materials Protein solution of interest TBS: 0.2 M NaCl in 50 mM Tris, pH 7.4 Blocking solution: 2% (w/v) milk powder in TBS with or without 0.1% (w/v) Tween 20 Antibody against the protein of interest diluted in blocking solution Secondary antibody coupled with horseradish peroxidase (HRP) diluted in blocking solution 4-chloro-1-naphthalene solution (see recipe) 0.2-µm nitrocellulose membrane (e.g., Schleicher and Schuell) 96-well plates Rotary shaker 1. Place matching round disks of 0.2-µm nitrocellulose membrane into wells of a 96-well plate. When large numbers of samples have to be analyzed, the use of a commercially available protein-dispersing device is advised (e.g., Dot Punch IM-96, Inotech AG). These devices allow the application of protein solutions to the membrane before they are put into the wells. This is more convenient and less time-consuming compared to the procedure described below.

2. Apply small volumes (50% is obtained under the conditions described. In the case of the IgSF-CAMs axonin-1 and NgCAM, ∼16,000 molecules were found to bind per Covasphere bead (Kuhn et al., 1991).

Covalent Coupling of Proteins to Glutaraldehyde-Activated Amino Beads As an alternative method for coupling proteins to beads, this protocol describes the glutaraldehyde-activated coupling of proteins to microspheres with amino-functional groups. Purified IgSF-CAMs can be coupled directly (not oriented) to amino beads. Alternatively, Fc-containing recombinant proteins can be bound to protein A–conjugated microspheres in an oriented manner. Additional Materials (also see Basic Protocol 2) Amino-functional microspheres (e.g., silica aminopropyl beads from Bangs Laboratories) 0.1 M NaOH or HCl (optional) 8% (v/v) EM-grade glutaraldehyde, newly opened bottle 400 µg/ml biochemically purified IgSF-CAMs (see Basic Protocol 1) in a phosphate-based buffer system (either PBS or 20 mM sodium phosphate, pH 7.0) Blocking solution (see recipe)

SUPPORT PROTOCOL 6

Cell Adhesion


Supplement 11

1. Prepare an appropriate volume (e.g., 0.5 ml) of 1% amino-functional microsphere stock solution in ultra pure water. Wash beads two times with 1 ml water. Centrifuge for 4 min at 16,000 × g, room temperature, to separate microspheres from wash solution. Remove supernatant and resuspend microspheres in 0.5 ml water. The amino-functional microspheres are available as powder or 10% (w/v) solution. Keep microsphere stock solutions at 4°C and never freeze.

2. Before adding the glutaraldehyde solution, check under the microscope whether the bead solution is monodisperse. If the bead solution contains significant clumps, sonicate in the water bath sonicator for 5 min at room temperature. If clumps persist, use a Branson microtip sonicator at the lowest output power needed to disrupt the clumps. Check pH of bead solution with cut pH strips and, if necessary, adjust pH to 6.5 or 7.0 with 0.1 M NaOH or HCl, respectively. Glutaraldehyde activation works best at pH 6 to 7.

3. If the bead solution is monodisperse and has a pH of 6.5 to 7.0, add an equal volume of a newly opened bottle of 8% EM-grade glutaraldehyde and mix. Incubate beads on rotator for ≥6 hr or overnight at room temperature. 4. Centrifuge beads as in step 1, remove supernatant, and wash activated beads at least three times with 1 ml water, and once with the buffer in which the protein is dissolved (PBS or 20 mM sodium phosphate). 5. Add 0.5 ml of 400 µg/ml purified IgSF protein in PBS, pH 7.3, or in 20 mM sodium phosphate, pH 7.0, to bead pellet and resuspend beads in protein solution (1% beads during coupling reaction). Incubate on rotator for 4 hr at room temperature or overnight at 4°C. If less protein is available, scale down amounts of protein and beads proportionally.

6. Centrifuge beads 4 min at 16,000 × g, room temperature, save supernatant for coupling analysis on SDS-PAGE, and resuspend beads in 0.5 ml blocking solution. Incubate beads in blocking solution for 30 min at room temperature on rotator. 7. Centrifuge beads again. Resuspend beads in 0.5 ml blocking solution (1% bead stock) and store at 4°C for months. BASIC PROTOCOL 3


Binding of Protein-Conjugated Microspheres to Cultured Cells This protocol determines binding specificity of a cell adhesion molecule to a receptor on a particular cell type. Both primary cell cultures (Kuhn et al., 1991; Suter et al., 1995) and cell lines (Buchstaller et al., 1996; Rader et al., 1996) can be incubated with protein-conjugated microspheres under live conditions to determine interactions between specific proteins and cells. Antibody preincubations can be used as a control but also for receptor identification. Furthermore, cells transiently transfected with mutated receptor forms can be used for the identification of extracellular protein domains necessary for binding of the cell adhesion protein (Rader et al., 1996; Kunz et al., 1998; Fitzli et al., 2000). Materials Primary cultures of neuronal and/or glial cells or other cells Complete medium used for cell cultures Serum-free BSA-containing cell culture medium 1011 beads/ml protein-conjugated fluorescent polystyrene microspheres; stock solutions in 0.5% BSA (see Support Protocol 5)



0.5 mg/ml Fab against protein of interest in serum-free medium (optional) Fixation solution (see recipe) PBS (see recipe) Mounting medium (see recipe) Waterbath sonicator 37°C, 10% CO2 humidified incubator Glass microscope slides Fluorescence microscope Prepare and bind microspheres to cells 1. Cultivate primary neurons and/or glial cells in conditions that allow live cell incubations in relatively small volumes. For example, grow cells on substrate-coated coverslips using a removable donut-shaped Teflon ring to limit the incubation volume to 200 to 250 2l (Suter et al., 1995). If culture conditions include the use of serum, prepare also a corresponding serum-free, BSA-containing medium as in Stoeckli et al., 1991; see recipe.

2. Wash cultured cells one time with complete culture medium and one time with serum-free, BSA-containing cell culture medium. Medium exchanges are carried out carefully with a pipet.

3. Prepare 1:1000 dilutions of 1011 beads/ml protein-conjugated fluorescent polystyrene microspheres in serum-free cell culture medium, sonicate dilutions for 2 min, room temperature and immediately add to cultured cells. 4. (Optional) To test whether a specific CAM binds to a particular cell type via a characterized receptor (to which antibodies are available), preincubate cells with 0.5 mg/ml Fab fragments in serum-free medium, before adding the beads, for 2 hr in a 37°C, 10% CO2 humidified incubator. Remove unbound antibodies by washing two times with serum-free medium before proceeding to step 5. To test for the specificity of bead binding to cells, use control protein–conjugated beads as well as beads that were preincubated with the corresponding Fab fragments. 5. Incubate the cells with the bead solution (step 3) for 1 hr in a 37°C, 10% CO2 humidified cell culture incubator. 6. Use a Pasteur pipet to carefully aspirate medium with unbound beads. Immediately add serum-free medium. Repeat this step three times. Do not allow the cells to dry. Beads bound to live cells can now be inspected. However, if more time is needed for analysis, the authors recommend fixing the cells. Fixation also allows processing of the cells for immunostaining of marker proteins or CAMs and, therefore, correlative analysis of CAM binding and CAM expression.

Fix cells 7. Fix the cells by adding 80 µl of 4× fixation solution to the cultures, which are in 240 µl of serum free medium. Gently mix and incubate 1 hr at 37°C. 8. Wash fixed cells three times with PBS. 9. For immunofluorescence staining of the cells, proceed as described in UNIT 4.3. Otherwise mount cells on a glass microscope slide in mounting medium for inspection under a fluorescence microscope. Beads are generally intensely fluorescent. Therefore, relatively short exposure times are sufficient when taking pictures. Cell Adhesion


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CELL-BASED ASSAYS FOR IgSF-CAM FUNCTION BASIC PROTOCOL 4

Trans-Interaction Assay with Myeloma Cells Interactions between IgSF-CAMs that have been determined either by a fluorescent microsphere assay or by binding of fluorescent microspheres to cells do not provide any information on whether the interaction occurs between molecules located on different cells (trans-interaction) or between molecules residing on the membrane of the same cell (cis-interaction). In order to distinguish between a trans- and a cis-interaction, the IgSF-CAMs need to be studied in a natural environment, i.e., as membrane-bound molecules residing in their proper orientation in a biological membrane. Expression in nonadherent myeloma cells (see Support Protocol 7) provides a means of assessing IgSF-CAMs for trans-interactions. Two populations of stably transfected myeloma cell clones are stained with optically distinct intracellular fluorescent dyes. The cells are dissociated, incubated, and examined under a microscope in order to determine whether re-aggregation has occurred. To evaluate whether two populations of myeloma cell clones expressing the IgSF-CAMs of interest on their surface can adhere to each other, they are labeled, mixed, dissociated, and allowed to re-aggregate. Labeling with optically distinct fluorescent dyes is necessary to distinguish the two populations. Materials Two populations of myeloma cell clones expressing the CAMs of interest Selection medium, e.g., 5 mM L-histidinol in DMEM supplemented with 10% (v/v) FCS PBS with Ca2+/Mg2+ (see recipe) Stock solution of green fluorogenic dye, e.g., 1 mM 2′,7′-bis-(2-carboxyethyl)-5-(and-6)-carboxyfluorescein acetoxymethyl ester (Molecular Probes) in DMSO Stock solution of red fluorogenic dye, e.g., 7.5 mM 5-(and-6)-carboxynaphtofluorescein diacetate (Molecular Probes) in DMSO 1% (v/v) FCS in PBS with Ca2+/Mg2+ 5 mg/ml p-phenylenediamine in 1% FCS in PBS with Ca2+/Mg2+ 15-ml conical polypropylene centrifuge tubes Hemacytometer V-shaped 96-well microtiter plate (e.g., Costar, Corning) Centrifuge and rotor for microtiter plates 22-G needle attached to 1-ml syringe Glass microscope slide Fluorescence microscope with appropriate filters for green and red fluorescence, e.g., FITC and Texas Red Additional reagents and equipment for counting cells (UNIT 1.1) Prepare cells 1. Grow transfected myeloma cell clones that express the CAM(s) of interest on their surface in selection medium to a cell density of ∼5 × 105 cells/ml. Use a hemacytometer to monitor the cell density (the aggregation assay requires 5 × 105 cells per population per sample; see UNIT 1.1).


Transfected myeloma cell clones that express high concentrations of a homophilically trans-interacting cell adhesion molecule can grow in very large cell aggregates consisting of thousands of cells that are easily visible by eye. The authors found that cultures of aggregating myeloma cell clones contain more dead cells than nonaggregating myeloma cell clones. This might, at least in part, be due to a limited oxygen and nutrition supply to



the center of the aggregate. In order to limit the number of dead cells, aggregating myeloma cell clones should be split more frequently avoiding large cell aggregates. The cell density can be determined with a hemacytometer after dissociating the cells by repeated pipetting through a 22-G needle.

2. Transfer 5 to 10 ml of cell culture into 15-ml conical polypropylene tubes and centrifuge for 3 min at 500 × g, room temperature. Aspirate the supernatant and resuspend the cells in 1 to 2 ml PBS with Ca2+/Mg2+. Determine the cell density with a hemacytometer (UNIT 1.1). Add PBS with Ca2+/Mg2+ to give a cell density of 5 × 105 cells/150 µl. Pipet 150 µl/well to a V-shaped 96-well plate. The aggregation assay requires two populations in distinct wells per sample. Use wells A1 and A2 for the first sample, B1 and B2 for the next sample, etc. When desired, the cells can be pre-incubated with antibodies at this step by incubating the cells with Fab monoclonal or polyclonal antibodies in a concentration range of 10 to 500 µg/ml in 1% FCS in PBS with Ca2+/Mg2+ for 1 hr at room temperature.

3. Centrifuge the microtiter plate for 2 min at 500 × g, room temperature, remove the supernatant by flicking plate into a sink (cells will remain in wells). Resuspend cells in 90 µl PBS with Ca2+/Mg2+. Label cells 4. Prepare fresh working solutions of both, green and red, fluorogenic dyes by diluting 10 µl of the stock solution in 990 µl PBS with Ca2+/Mg2+. Add 10 µl of the appropriate working solution to the well with the appropriate 90-µl cell suspension. Use one column for one fluorogenic dye (e.g., stain A1, B1, etc. with green and A2, B2, etc. with red). Incubate 30 min at 37°C. The end concentration of the fluorogenic dyes is 1 µM of 2′,7′-bis-(2-carboxyethyl)-5-(and6)-carboxyfluorescein acetoxymethyl ester and 7.5 µM of 5-(and-6)-carboxynaphtofluorescein diacetate. The electrically neutral ester substrates freely diffuse through the cell membrane into the cell, where they are cleaved into fluorescent products by nonspecific intracellular esterases. The charged fluorescent products are retained by cells with intact plasma membranes. Serum of the cell culture medium contains esterases and has to be washed away prior to the incubation and avoided during the incubation.

5. Centrifuge microtiter plate and remove the supernatant as in step 3. Resuspend the cells in 150 µl PBS with Ca2+/Mg2+. Repeat step. 6. Centrifuge microtiter plate 2 min at 500 × g, room temperature and remove the supernatant by flicking plate into sink. Resuspend cells in 75 µl of 1% FCS in PBS with Ca2+/Mg2+. Dissociate cells and allow re-aggregation 7. Combine complementarily stained cells of one sample in one well (e.g., add A2 to A1, B2 to B1, etc.). Dissociate cells by slowly pipetting up and down ten times through a 22-G needle attached to a 1-ml syringe. Avoid foaming. Incubate 45 min at 4°C. During re-association the plate should not be moved.

8. Centrifuge microtiter plate for 2 min at 500 × g, room temperature, and remove the supernatant by flicking plate into sink. Resuspend the cells in 150 µl of 1% FCS in PBS with Ca2+/Mg2+. Repeat step. 9. Centrifuge microtiter plate and remove the supernatant as in step 8. Resuspend the cells in 40 µl of 1% FCS in PBS with Ca2+/Mg2+.

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Analyze cells 10. Immediately prior to microscopic analysis of an individual sample, add 10 µl of 5 mg/ml p-phenylenediamine in 1% FCS in PBS with Ca2+/Mg2+. Pipet the sample several times up and down using a 200-µl pipet tip. Mount 10 µl of the sample on a glass slide. The use of the antifading reagent p-phenylenediamine in an end concentration of 1 mg/ml markedly preserves the fluorescence intensity.

11. Analyze the aggregates with a fluorescence microscope using FITC and Texas Red filters. Filters that allow the simultaneous detection of green and red fluorescence facilitate the analysis. SUPPORT PROTOCOL 7

Stable Transfection of Myeloma Cells by Protoplast Fusion This protocol describes the stable transfection of myeloma cells with a vector that allows the surface expression of IgSF-CAMs. A vector particularly suited for myeloma cell expression was described by Traunecker et al. (1991). Expression by this vector is driven by an Ig κ promoter and enhancer. The 3′ end of the transcript of interest is spliced onto an exon encoding the Ig κ constant domain in order to mimic stable Ig transcripts. The vector contains a histidinol dehydrogenase gene that allows the selection of stable transfectants in the presence of L-histidinol. L-histidinol is a precursor of L-histamine and an inhibitor of protein synthesis. The vector has been stably transfected into the mouse myeloma cell line J558L for the production of soluble lymphocyte-derived cell-surface receptor proteins (Traunecker et al., 1991). The system has also been used for the surface expression of IgSF-CAMs (Rader et al., 1993; Buchstaller et al., 1996; Fitzli et al., 2000). Alternatively, other mammalian expression vectors and myeloma cells can be used. Myeloma cell clones that stably express large amounts of IgSF-CAMs on their surface were generated by a transfection method known as protoplast fusion. Transfection by protoplast fusion is a highly efficient method for the direct transfer of mammalian expression vectors from bacteria to mammalian cells (Schaffner, 1980; Sandri-Goldin et al., 1981; Rassoulzadegan et al., 1982; Gillies et al., 1983). It involves digesting bacterial cell walls with lysozyme to produce protoplasts and then fusing the protoplasts to mammalian cells in the presence of polyethylene glycol. The following protocol is based on the myeloma expression system described by Traunecker et al. (1991) and can easily be adapted to other systems.


Materials Glycerol stock of an E. coli strain 803 clone (ATCC #35581) transformed with a mammalian expression vector containing the cDNA of the IgSF-CAM of interest (store at −80°C) LB agar/ampicillin plates (see recipe; store at 4°C) DMEM supplemented with 10% (v/v) FCS LB medium (see recipe), prewarmed to 37°C 50 mg/ml ampicillin (store at −20°C) 60 mg/ml chloramphenicol in ethanol (store at −20°C) DMEM supplemented with 10% (w/v) sucrose and 10 mM MgCl2, prewarm 20% (w/v) sucrose in 50 mM Tris⋅Cl, pH 8.0, ice cold 1 mg/ml lysozyme (Roche Molecular Systems), freshly dissolved 10 mg in 10 ml of 250 mM Tris⋅Cl, pH 8.0, and filtered through 0.22-µm filter 250 mM EDTA, pH 8.0, ice cold 50 mM Tris⋅Cl, pH 8.0, ice cold



10 mg/ml DNase I (Roche Molecular Systems; store at −20°C) DMEM PEG 1500 in DMEM supplemented with DMSO (see recipe) Mouse BALB/c myeloma cell line J558L (ECACC #88032902) or another myeloma cell line 50 mg/ml kanamycin 50 mM L-histidinol (see recipe) Polyclonal anti-IgSF-CAM antibody Fluorescein-conjugated secondary antibody 25-ml cell culture flasks 12-ml and 50-ml polypropylene tubes 15- and 50-ml conical polypropylene centrifuge tubes 37°C bacterial shaker 500-ml Erlenmeyer flask Refrigerated tabletop centrifuge 37°C water bath Glass microscope slides Microscope with 1000× magnification Multipipet trays 24- and 96-well tissue culture plates Multipipettor and tips Plastic wrap (e.g., Saran) 96-well plates with V-shaped wells Additional reagents and equipment for indirect immunofluorescence (UNIT 4.3) and freezing cells (UNIT 1.1) NOTE: All solutions and equipment coming into contact with living cells must be sterile, and aseptic technique should be used accordingly. NOTE: All cell culture incubations should be carried out in a 37°C, 5% CO2 humidified incubator. NOTE: The following protocol is written for one sample. It is not recommended to handle more than four samples in parallel. Day 1: grow transformed bacterial strain 1. Streak a glycerol stock of an E. coli strain 803 clone containing the mammalian expression vector onto an LB agar/ampicillin plate. Grow overnight at 37°C. E. coli strain 803 (also termed 1106) might be more efficient than other E. coli strains in producing stable protoplasts (Rassoulzadegan et al., 1982). The antibiotic has to be adapted to the prokaryotic selection marker of the mammalian expression vector.

2. Grow myeloma cells in DMEM supplemented with 10% (v/v) FCS in 25-ml cell culture flasks. Aim at a high cell density of ∼1 × 106 cells/ml that is reached on day 3 (protoplast fusion requires 5 × 106 cells per sample). Day 2: grow transformed bacterial cultures 3. Inoculate 2 ml LB medium prewarmed at 37°C in a 12-ml polypropylene tube with a single E. coli colony from the freshly streaked LB agar/ampicillin plate. Add 2 µl of 50 mg/ml ampicillin. Grow for 4 hr at 250 rpm in a 37°C bacterial shaker. Cell Adhesion


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4. Dilute 100 µl of cell culture into a 500-ml Erlenmeyer flask with 100 ml of LB medium and 100 µl of 50 mg/ml ampicillin. Grow to an OD600 of ∼0.6. Start checking the optical density after 3 hr. 5. After OD600 ∼0.6 is reached, add 200 µl of 60 mg/ml chloramphenicol to a final concentration of 120 µg/ml. Grow overnight at 250 rpm in a 37°C bacterial shaker. Plasmids carrying the colE1 origin of replication can be amplified in the presence of chloramphenicol (Hershfield et al., 1974).

Day 3: harvest bacterial cells and form protoplasts 6. Transfer the overnight culture into two 50-ml conical polypropylene centrifuge tubes and centrifuge 10 min at 2500 × g, 4°C. 7. In the meantime, prewarm 20 ml DMEM supplemented with 10% sucrose and 10 mM MgCl2 in a 50-ml polypropylene tube in a 37°C water bath. 8. Pour off the supernatants of the spun culture. Prepare protoplasts 9. From here on proceed in a sterile laminar flow bench. Vortex and combine the two bacterial pellets in 2.5 ml ice-cold 20% sucrose in 50 mM Tris⋅Cl, pH 8.0. 10. Add 500 µl ice-cold 1 mg/ml lysozyme in 250 mM Tris⋅Cl, pH 8.0, mix gently (swirl), and incubate 5 min on ice. 11. Add 1 ml ice-cold 250 mM EDTA, pH 8.0, gently swirl, and store on ice 5 min. 12. Add 1 ml ice-cold 50 mM Tris⋅Cl, pH 8.0, gently swirl, and incubate 10 min at room temperature. During this incubation period, mount 10 µl of the sample on a glass microscope slide and analyze protoplast formation under a microscope. A microscope with 1000× magnification is required to distinguish between spherical protoplasts and rod-shaped bacteria. At the end of incubation, ∼90% protoplasts should be formed.

13. Add 20 ml DMEM supplemented with 10% sucrose and 10 mM MgCl2 very slowly to the protoplast preparation. To do this, swirl protoplasts gently, start adding drops of DMEM supplemented with 10% sucrose and 10 mM MgCl2, and slowly increase the added volume. The prepared protoplasts are fragile and need to be handled with care. Protoplast lysis is indicated by an increasing viscosity of the preparation due to the release of genomic DNA. The preparation can be analyzed using a microscope as above.

14. Add 40 µl of 10 mg/ml DNase I and incubate 15 min at room temperature. Addition of DNase I reduces the viscosity of the protoplast preparation.

Prepare for fusions 15. In the meantime, prewarm the following in separate 50-ml propylene tubes in a 37°C water bath: 15 ml DMEM 10 ml DMEM supplemented with 10% (v/v) FCS 50 ml DMEM supplemented with 10% (v/v) FCS. Cell-Cell Contact by Ig Superfamily Cell Adhesion Molecules

Thaw at room temperature: 2 ml PEG 1500 in DMEM supplemented with DMSO.



16. Centrifuge protoplast preparation in the 50-ml polypropylene tube 30 min at 2500 × g, room temperature. 17. In the meantime, transfer 5 × 106 mouse BALB/c myeloma cells into 15-ml conical polypropylene centrifuge tubes and centrifuge 10 min at 500 × g, room temperature. Aspirate the supernatant and resuspend the cells in 5 ml prewarmed DMEM. The myeloma cell preparation should be serum-free. Other myeloma cell lines that have been transfected by protoplast fusion or electroporation include mouse P3-X63Ag8.653, mouse Sp2/0-Ag14, mouse NSO, and rat YB2/0 (Gillies et al., 1989; Nakatani et al., 1989; Bebbington et al., 1992; Shitara et al., 1994).

18. Pour off the supernatant of the centrifuged protoplast pellet. The protoplast pellet should have a smooth surface.

Carry out fusions 19. Slowly layer the myeloma cell preparation on top of the protoplast pellet in the 50-ml conical polypropylene centrifuge tube. Centrifuge 10 min at 500 × g, room temperature. 20. Aspirate the supernatant. Mix cell and protoplast pellet by hand-flicking the tube and tapping it on the benchtop. 21. Add 2 ml PEG 1500 in DMEM supplemented with DMSO. Resuspend the pellet by pipetting up and down several times. 22. After addition of the PEG solution (∼1 to 2 min), very slowly add 10 ml prewarmed DMEM. To do this, swirl protoplasts gently, start adding drops of DMEM, and slowly increase the added volume. 23. Add 10 ml prewarmed DMEM supplemented with 10% FCS, swirl gently, and centrifuge 10 min at 500 × g, room temperature. 24. Aspirate the supernatant, resuspend the pellet in 50 ml prewarmed DMEM supplemented with 10% FCS, and add 100 µl of 50 mg/ml kanamycin. 25. Pour into a multipipet tray and distribute among five 96-well tissue culture plates by adding 100 µl/well using a multipipettor. Wrap tissue culture plates in plastic wrap and incubate for 48 hr in a 37°C, 10% CO2 humidified incubator. Day 5: select transfected cells 26. After 48 hr, prepare selection medium by adding 10 ml prewarmed 50 mM L-histidinol to 40 ml prewarmed DMEM supplemented with 10% FCS. Add 100 µl of 50 mg/ml kanamycin. Pour into a multipipet tray and add 100 µl/well using a multipipet. Rewrap tissue culture plates in plastic wrap and continue incubation in a 37°C, 10% CO2 humidified incubator. The antibiotic, here L-histidinol, has to be adapted to the eukaryotic selection marker of the mammalian expression vector. Only transfected myeloma cells will survive the treatment with L-histidinol.

27. Examine plates visually for clones ∼10 days after selection medium is added. No medium change or any other treatment is necessary during this time

Identify IgSF-CAM expressing clones 28. Once a clone becomes clearly visible by eye, analyze it for the expression of IgSF-CAM by indirect immunofluorescence staining (UNIT 4.3). Transfer ≤50% of the cloned cells to a well of a 96-well plate with V-shaped wells and perform indirect Cell Adhesion


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immunofluorescence analysis with a polyclonal anti-IgSF-CAM and a fluoresceinconjugated secondary antibody. Identify clones that are expressing IgSF-CAM. 29. Expand positive clones into 24-well tissue culture plates. 30. Subclone positive clones by limiting dilution in 96-well tissue culture plates. 31. Maintain positive subclones in selection medium, e.g., 5 mM L-histidinol in DMEM supplemented with 10% FCS. Store backup cells frozen in liquid nitrogen using standard procedures (UNIT 1.1). Cells from positive subclones are used in myeloma cell aggregation assays. BASIC PROTOCOL 5

Detecting Cis-Interactions between IgSF-CAMs by Chemical Cross-Linking The following basic protocols describe methods for the detection of cis-interactions, i.e., interactions between proteins that reside in the same membrane. In the chemical crosslinking methods described in this protocol, bifunctional reagents are used to establish covalent cross-bridges between associated proteins. The covalently linked protein complexes are then analyzed by SDS-PAGE (UNIT 6.1) and immunoblots (UNIT 6.2), using specific antibodies. In the antibody-induced co-capping methods (see Basic Protocol 6), a hypothesized cis-interaction between two cell surface proteins is evaluated by inducing a redistribution of one molecule and testing whether the putative binding protein follows. For both methods, the cells analyzed should be cultured at low density, in order to prevent contact between cells. Under these conditions, close associations of proteins are only possible between proteins residing in the membrane of the same cell. Chemical cross-linking joins two molecules by means of a cross-linking reagent. The method critically depends on the cultivation of the cells of interest as single cells at low density to avoid the formation of cell-cell contacts. Moreover, the structural integrity of the cells should be maintained throughout the procedure. The use of hydrophilic, membrane-impermeable bifunctional cross-linking reagents restricts the cross-linking to extracellular domains of membrane proteins. Considering the close spatial association between proteins interacting within the same membrane, a high degree of specificity of chemical cross-linking is mandatory.


The specificity of chemical cross-linking is mainly determined by the chemical reactivity of the functional groups of the cross-linking reagent and the length of the spacer separating the reactive groups. The N-succinimidyl group combines efficient reactivity with a high selectivity for primary amino groups, thereby limiting the cross-linking to lysine side chains at the surface of proteins engaged in interactions. Since most IgSF-CAMs contain multiple lysine residues in their extracellular domains, the amino-group-specific homobifunctional N-succinimide-derivatives are suitable reagents for the detection of cis-complexes formed between such molecules. The following protocol will primarily focus on the application of hydrophilic homobifunctional di-N-succinimidyl derivatives with relatively short (0.6 to 1.2 nm) spacer sequences separating the reactive groups. In order to enhance the specificity of the chemical cross-linking reaction to stably associated proteins, lateral movement in the cell membrane is reduced by performing the cross-linking reaction on ice. Ideal spacer length of the cross-linking reagent, optimal concentrations of cross-linkers, and reaction times strongly depend on the specific system that is analyzed and must be evaluated empirically. The parameters described below turned out to be optimal in many experimental situations and represent a good starting point for further optimizations. After quenching the cross-linking reaction, cells are lysed and the crosslinked complexes ideally isolated by immunoprecipitation (UNIT 7.2) using specific antibodies against the molecule of interest. Immunoprecipitates are separated by SDS-PAGE and cross-linked complexes can be detected by immunoblot analysis. The presence of



known binding partners within the cross-linked complexes can be addressed by immunochemical techniques, which require only low amounts of proteins. However, immunochemical analysis clearly restricts the detection of cross-linked binding partners to known molecules against which antibodies are available. Scaling-up of the procedure may result in the isolation of sufficient amounts of cross-linked material for subsequent microsequencing allowing the detection of novel binding partners. NOTE: All solutions and equipment coming into contact with living cells must be sterile, and aseptic technique should be used accordingly. NOTE: All cell culture incubations should be carried out in a 37°C, 5% CO2 humidified incubator. Materials Cells of interest growing in tissue culture at low density PBS with Ca2+/Mg2+ (see recipe) 100 mM cross-linking reagent (see recipe): Bis(sulfosuccinimidyl)suberate (BS3) in water Disuccinimidyl tartrate (DST) in water-free DMSO Disulfo disuccinimidyl tartrate (Sulfo-DST) in water 3, 3′-Dithiobis(sulfosuccinimidyl propionate) (DTSSP) in water 5 mM EDTA in Ca2+/Mg2+-free PBS 1 M glycine solution in water, pH 8.0 Lysis buffer (see recipe) Primary antibody: serum, purified immunoglobulin, or purified immunoglobulin immobilized on agarose or Sepharose matrix Protein A or protein G coupled to agarose or Sepharose matrix (optional) Wash buffer (see recipe) Sample buffer for SDS-PAGE (APPENDIX 2A) 10-cm tissue culture dishes precoated with poly-D-lysine combined with additional substrates, such as laminin (see Support Protocols 10 and 12) Horizontal shaker Cell scraper 2-ml microcentrifuge tubes End-over-end rotator (model 750) 100- or 200-µl and 500-µl Hamilton syringe and 22-G needle Additional reagents and equipment for SDS-PAGE (UNIT 6.1), immunoblot analysis (UNIT 6.2), and immunoprecipitation (UNIT 7.2) Prepare cells 1. Prior to the experiment, seed cells onto 10-cm tissue culture dishes at a low density. Plate slowly dividing (division rate of 30 kb) frequently result in the formation of much larger precipitates, reducing the efficiency of transfection considerably. A reaction at pH 7.0, room temperature (20° to 25°C), and 1 min reaction time are good starting conditions for optimizations. The formation of precipitate after mixing CaCl2 solution with HBS solution can be monitored by measuring the absorbance at 320 nm (e.g., for the testing of new batches of solutions). After addition to the cells, the stability of the DNA-containing calcium phosphate particles is the most critical factor for the efficiency of transfection. One source of instability is a reduction in medium pH due to the metabolic activity of cells. It is therefore important to include 10 mM HEPES (final concentration) in the 293 cell culture medium. The reduction of the CO2 partial pressure to 3% is another option to raise the pH value, if necessary. Increased time of exposure to the precipitate can further enhance the efficiency of transfection. An incubation time of 4 to 6 hr given in the protocol is a good starting point and can be extended up

to 24 hr in cases where no cytotoxicity is observed. However, the combination of exposure to the precipitate with an osmotic shock, e.g., by adding 10% (w/v) glycerol to the cell culture medium, as suggested by some authors for the calcium phosphate transfection of Chinese hamster ovary (CHO) cells, is not recommended for 293 cells. Immunoblotting No positive dots. The concentration of the protein of interest may be below the detection limit. Use repeated applications of protein solution to the nitrocellulose membrane. Apply the purified protein onto the membrane as a positive control. If the total protein concentration of the solution applied to the membrane is high, even repeated applications of protein solution to the membrane may not give a satisfactory result. High background. Make sure that the volumes of protein solution applied to the membrane are small. Let the dot dry before the next step. Wash membranes more thoroughly. Change the blocking solution and/or blocking time. For instance, try blocking with serum from the same species that was used to raise the secondary antibody. Microsphere assays False negative results in the bead aggregation assay. Coupling reaction results in protein orientation on the bead surface in a way that binding sites are not accessible. Alternatively, proteins could not be physiologically active after purification (e.g., elution after immunoaffinity column). False positive results in the bead aggregation assay. Protein preparations that are not pure and contain another CAM could result in false positive binding results. Furthermore, if each of the two test proteins has homophilic binding properties, weak unspecific aggregation of preformed homophilic aggregates could result in false positive results with respect to heterophilic interactions. Therefore, check for unevenly distributed beads in mixed aggregates under the microscope (bead distribution in aggregates is not detectable in a flow cytometer). Discrepancies between results of bead and cell aggregation assays. The orientation of proteins on the beads is likely to be random and not uniform as in a biological membrane. This has to be considered when interpreting and comparing the results with the ones of the cell aggregation assay. Furthermore, the orientation problem has to be taken into account when

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analyzing cis- and trans-interactions (Kuhn et al., 1991; Buchstaller et al., 1996; Rader et al., 1996; Sonderegger et al., 1998). Orientation of proteins on beads can be better controlled when using proteins with domains or tags that tightly bind to a linker protein that is coupled first to the microsphere. For example, recombinant Fc-fusion proteins can be coated on protein A-conjugated beads in an oriented manner. One has to consider that this linkage includes a noncovalent protein interaction. Trans-interactions The stably transfected myeloma cell clones may differ from the parental myeloma cell line in more than only the expression of the IgSFCAM. The upregulation of endogenous cell adhesion molecules, such as integrins and IgSF-CAMs, can result in myeloma cell aggregation (Kawano et al., 1991). It is therefore essential to run a series of control experiments that address the question whether any cell aggregation is caused by a trans-interaction of the expressed IgSF-CAMs or by other factors. For this, follow these guidelines: 1. Confirm the functional surface expression of the IgSF-CAM using, for example, monoclonal antibodies directed against conformational epitopes. 2. Each aggregation assay should be performed in duplicate with cross-wise exchange of the fluorescent dyes. 3. Myeloma cell clones that express an IgSF-CAM should not form mixed aggregates with the parental myeloma cell line. 4. Independent myeloma cell clones that express the same IgSF-CAM should give the same aggregation pattern. 5. Independent myeloma cell clones that express different quantities of the same IgSFCAM should be analyzed for a correlation between expression and aggregation. 6. Pre-incubation with polyclonal Fab directed to the IgSF-CAM should prevent aggregation. 7. Pre-incubation with an enzyme that removes the IgSF-CAM selectively from the surface should prevent aggregation. A very useful enzyme for this is phosphatidylinositol-specific phospholipase C, which selectively cleaves glycosyl-phosphatidylinositol-anchored proteins.


Chemical cross-linking The application of cross-linking reagents on intact, live cells may result in possible artifacts due to the perturbation of the architecture of the

cell membrane by the chemicals. Very often, the reagents are used at millimolar or higher concentrations (0.1 to 10 mM) and must be dissolved in organic solvents prior to use. It is therefore very important to assure that the reagents by themselves and especially organic solvents, if used, do not affect the structural integrity and viability of the cells tested. Watersoluble reagents like BS3, Sulfo-DST, and DTSSP that carry sulfonyl groups can be used to circumvent the solvent problem. Successful detection of cis-interactions between IgSF-CAMs requires a high specificity of chemical cross-linking on the one hand, and sufficient yields of cross-linked materials on the other hand, for subsequent biochemical characterization of cross-linked partners. As described above, the specificity and efficiency of the cross-linking reaction is determined by the following parameters: length of the spacer separating the reactive groups; chemical reactivity of the functional groups of the cross-linking reagent; concentration of cross-linking reagent; and reaction time. Cross-linking reagents with short spacers between the reactive groups such as DST and Sulfo-DST restrict chemical coupling to closely associated molecules and are therefore preferable for the detection of cis-complexes between membrane proteins. However, due to the spatial proximity of their functional groups, these reagents exhibit an enhanced tendency for intramolecular cross-linking, i.e., coupling lysine side chains of the same molecule with each other, which may result in very low yields of cross-linked material. The application of reagents with longer spacers between the reactive groups such as BS3 and DTSSP generally results in higher yields, but bears the risk of unspecific reactions. As previously discussed, the chemical reactivity of the functional groups present in commercially available cross-linking reagents range from highly selective, like the N-succinimidyl group, to rather unselective photoactivated groups, like aryl azides, which generate highly reactive, unstable intermediates that undergo reactions with a wide variety of chemical structures within a protein. Although the N-succinimidyl group is most frequently used for modifications and cross-linking of cell surface proteins, its selective reactivity with primary amino groups (mainly lysine side chains) excludes this class of crosslinkers from extended hydrophobic interfaces through which IgSF-CAMs may interact. Upon photoactivation, reagents containing aryl azide groups exhibit a high reactivity towards ali-



phatic hydrocarbon groups, like RCH2R and R2CHR, that allows cross-linking within a strongly hydrophobic environment. However, the high reactivity and low selectivity of photoactivated aryl azides frequently results in a high degree of unspecific cross-linking, not only among proteins, but also extensive protein-lipid cross-linking. For the detection of specific interactions among the closely associated proteins on the surface of intact cells, these reagents are, therefore, not recommended as first choice, but represent an option in cases where no cross-linking products are obtained using more specific reagents. Optimal concentrations of cross-linking reagents must be evaluated empirically for every experimental system. A more detailed discussion of the chemical background can be found in Lomant and Fairbanks (1976), Lewis et al. (1977), and Smith et al. (1978). For homobifunctional N-succinimidyl derivatives, like DST and DSSP, optimal concentrations for cross-linking on intact cells are consistently in the range of 0.1 to 10 mM in published protocols. This range of concentrations represents a good starting point for optimizations. Crosslinking reagents with higher reactivity, like e.g., aryl azides, are generally applied in much lower concentrations, usually between 10 µM and 1 mM, for cross-linking on intact cells. A further critical parameter is reaction time. Longer reaction times generally result in better yields of cross-linked products, but also in more unspecific reactions. Using homobifunctional N-succinimidyl reagents, quenching of the cross-linking reaction after 0, 5, 15, and 45 min results frequently in a “kinetic profile” of the process. Specifically, a different pattern of cross-linked products is observed during the time course of the cross-linking reaction. This reflects the tendency of many membrane molecules, such as IgSF-CAMs, to form oligomeric or even multimeric aggregates in the membranes of living cells. The appearance of initial cross-linked molecules, after a few minutes, is often followed by the appearance of further cross-linked complexes with higher molecular masses, generated from the coupling of the initial complexes with additional molecules. Initial cross-linked products that are generated within the first minutes of the reaction may correspond to the first assembly units from which larger oligomeric or multimeric complexes are formed. The “kinetic profile” of the cross-linking reaction may, therefore, give some information about the nature of the com-

plexes or aggregates formed by molecules like IgSF-CAMs. Co-capping The demonstration of a cis-interaction by antibody-induced co-capping of cell surface molecules critically depends on many factors. To allow the induction of caps by antibody cross-linking and co-capping mediated by a cis-interaction, the molecules of interest must have a minimal lateral mobility within the plane of the cell membrane. In addition to mobility, the relative stoichiometry of the molecules of interest is critical for the detection of co-capping after antibody-induced capping. Ideally, comparable levels of expression at the cell surface allow reciprocal co-capping of two molecules that interact with each other in cis with sufficient affinity. However, large stoichiometric excess of one molecule due to different expression levels results in asymmetric results in reciprocal co-capping experiments; capping of the more abundant component is followed by clearly detectable co-capping of the less abundant molecule. In contrast, capping of the molecule expressed at lower level results in only partial co-capping of the more abundant component. For the detection of partial co-capping, confocal laser scanning microscopy is a powerful technique that allows the reliable detection of locally enhanced fluorescence signals (co-capping) versus a relatively high homogeneous background signal (uncapped monomers of the molecule present in stoichiometric excess). In addition to the factors described above, the availability of specific primary antibodies against the molecules of interest, raised in different species, is an essential prerequisite for co-capping experiments. For antibody-induced capping, polyclonal or monoclonal antibodies can be used. Polyclonal antibodies are able to cross-link the cell surface antigen to some extent in the absence of secondary antibody, whereas in the case of monoclonal primary antibodies, cross-linking of bound primary antibody with the secondary antibody is required for the induction of caps. The most critical factors of every co-capping experiment is the specificity of the primary antibodies used. A major source of experimental artifacts is the potential of cross-reactivity of primary antibodies, either between the molecules of interest or with other, unidentified molecules expressed on the cell. It is therefore of pivotal importance to exclude any cross-reactivity under the conditions (antibody concentrations, temperature,

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incubation times, etc.) used for co-capping and counter-staining. It is not recommended to test for cross-reactivity with other immunochemical techniques like immuno-blot or ELISA, because the antigens are presented in a different form on nitrocellulose and on plastic surfaces than on live cells. The absence of cross-reactivity of the partially or totally denatured proteins present in such immunochemical assays does, therefore, not necessarily exclude cross-reactivity of the native proteins present on the live cells under the conditions used for co-capping experiments. In situations where the proteins of interest are co-expressed in recombinant form in a cell type that does not normally express them, the test for cross-reactivity is straightforward. Specificity of detection can be checked by immunostaining of single transfectants and mock-transfected cells (as a negative control) with both antibodies (see, e.g., Buchstaller et al., 1996; Kunz et al., 1998). It should be noted that cross-reactivity of antibodies is a phenomenon that depends on the antibody concentrations used. Even in cases where no crossreactivity is reported in the literature (e.g., based on experience with standard protocols for immunofluorescence), cross-reactivity may occur at the higher antibody concentrations that are normally applied in co-capping experiments. Cross-reactivity of the secondary antibodies represents only a minor problem, since highly specific preparations of fluorochromelabeled secondary antibodies against a wide variety of species are commercially available. The use of phylogenetically more distant species is desirable since the potential of cross-reactivity between the secondary antibodies is lower. Combinations that are frequently documented in the literature are mouse/rabbit, mouse/goat (or sheep), rabbit/goat (or sheep). A considerable risk of cross-reactivity exists especially for the combination mouse/rat. In this case the choice of secondary antibodies has to be made with care. Several controls must be included to ensure specificity of antibody-induced capping. The use of preimmune serum, or purified IgG from preimmune serum, is an essential negative control in cases where complete sera or total IgG fractions, respectively, are used as a source of primary antibodies. Additional controls should include the detection of unrelated molecules expressed by the cell of interest that are not expected to co-distribute with one of the molecules tested for cis-interaction. Apart from false-positive results due to cross-reactivity of antibodies, false-negative

results can be due to potential interference of antibody binding with the interaction between the molecules of interest. Polyclonal antibodies directed against a variety of epitopes can perturb molecular interactions. It is therefore worthwhile to test several different antibodies, polyclonal as well as monoclonal, if available. The potential interference of antibody binding with the interaction between the molecules of interest can be prevented by using heterologously expressed recombinant proteins. Specifically, different N-terminal peptide tags, like the myc-tag or the influenza hemagglutinin-tag can be introduced by molecular cloning into the polypeptide sequences of the studied proteins. Subsequent capping with antibodies specific for the N-terminally localized tag sequences will reduce the risk of interference with a binding site on the surface of the molecule. It should always be kept in mind, that the detection of antibody-induced co-capping between two molecules is no proof of a direct molecular interaction between the two components. In order to demonstrate such a direct binding, additional experimental techniques, like chemical cross-linking (see Basic Protocol 5) or biochemical binding assays using isolated, purified proteins (see Basic Protocol 2) are required. Neurite outgrowth Many IgSF-CAMs have a neurite outgrowth-promoting activity, therefore, neurite growth assays are widely used for functional analysis. The preparation of neuronal cultures takes some practice and should best be learned in a laboratory where culture techniques are established. The reproducibility of the cultures is very important to get results. Keep in mind that the growth and the morphology of neurites depend on the substrate and the medium. As the quality and the components of serum differ considerably from batch to batch, the lot of serum used should not be changed during analysis. For best reproducibility, the use of a chemically defined, serum-free medium is recommended. All the solutions and the purified proteins to be tested for their neurite outgrowthpromoting activity have to be of high quality. Especially, the absence of endotoxins is extremely important for the survival of neurons. Make sure that detergents are removed carefully from protein solutions before using them in tissue culture. The growth characteristics of neurites are age-dependent. In particular, the dependence on specific growth factors may change dramati-



cally. Therefore, it is essential not to mix different ages of tissue. Similarly, the response of neurons to different substrates may be speciesspecific. Remember that neurite length is not the only criterion that can be assessed in neurite growth assays. Closely monitor neurite and growth cone morphologies and branching patterns.

moderate amounts of the IgSF-CAM, a small percentage typically reveals very high expression. Thus, the higher the transfection efficiency, the higher becomes the likelihood to obtain a myeloma cell clone with very high expression. This makes protoplast fusion the method of choice for the transfection of myeloma cells.

Anticipated Results

Chemical cross-linking During optimization of the cross-linking protocol, the emphasis should be placed on the specificity of the reaction. As described, there is generally an inverse relationship between specificity of a cross-linking reaction and its efficiency. A protocol that ensures a high degree of specificity often has the drawback of low yields of cross-linked material. Based on published results and experience in the authors’ laboratory, yields of cross-linking protocols, like the one described, range from 0.1% to 1%, corresponding to a few hundred nanograms of cross-linked material from a reaction performed on 106 cells. This amount of protein is normally sufficient for immunochemical characterization of the cross-linked molecules, e.g., by immunoblot analysis or re-immunoprecipitation (Buchstaller et al., 1996; Kunz et al., 1998). The appearance of only one or a few complexes of a molecule of interest indicates some degree of specificity of the reaction. The crosslinked complexes isolated by immunoprecipitation can be separated by SDS-PAGE (ideally two-dimensional; UNIT 6.4). The detection of presumed binding partners in the cross-linked complexes by immunoblot analysis should always include controls, that is the detection of a membrane protein present in the cell used for cross-linking that is not expected to associate with the molecule of interest.

Transfection For transient transfections with the protocol described, reporter constructs can be used to assess the efficiency of transfection and calculate the percentage of transfected cells. Using reporter constructs that express green fluorescent protein (GFP) as a reporter under the control of the human cytomegalovirus (CMV) immediate early promoter (plasmid size of 5 to 10 kb, DNA amounts for transfection between 40 and 80 µg), the authors repeatedly observed transfection efficiencies of 30% to 40% based on the detection of GFP expression 24 to 48 hr after transfection. Expression of recombinant protein under the control of strong viral promoters lasts, generally, for 48 to 96 hr after transfection. Microsphere assays With the appropriate controls, all the protocols described here that involve microsphere and cell aggregation techniques are powerful methods to study binding properties of IgSFCAMs. Furthermore, these assays are not very time-consuming and enable processing of several tests in parallel. Trans-interactions Myeloma cell clones that were stably transfected to express IgSF-CAMs form aggregates when the IgSF-CAMs interact in trans. Homophilic trans-interactions are indicated by aggregates of cells that express the same IgSF-CAM. Heterophilic trans-interactions are indicated when two populations of cells that express distinct IgSF-CAMs form mixed aggregates. Transfection-protoplast fusion On average, 100 clones are obtained per 96-well tissue culture plate, i.e., 500 clones per 5 × 106 transfected myeloma cells. Thus, the transfection efficiency is in the range of 1 × 10−4. Even with an optimized electroporation procedure, the transfection efficiency was ∼20 times lower (Rader et al., 1993). While the majority of myeloma cell clones expresses

Co-capping Antibody-induced co-capping can be observed between molecules that undergo direct or indirect cis-interactions. Very clear results can be obtained in cases where the two components are expressed at comparable levels and directly interact with each other with relative high affinity, as demonstrated in the example shown in Figure 9.5.3. Co-capping of molecule A (NgCAM) with molecule B (axonin-1) was studied on stably double-transfected CV-1 cells. Capping of molecule A (NgCAM), induced by the subsequent incubation with a mouse monoclonal primary and a rabbit-anti mouse secondary antibody (Fig. 9.5.3A), re-

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Figure 9.5.3 Antibody-induced co-capping of axonin-1 with NgCAM on stably double-transfected CV-1 cells. In doubletransfected CV-1 cells, NgCAM (A) and axonin-1 (C) were randomly distributed. Capping of NgCAM was induced by a mouse monoclonal antibody and a rabbit anti-mouse IgG. NgCAM caps were detected by a Texas Red–labeled donkey-anti-rabbit IgG (B). The distribution of axonin-1 was detected by counter-staining with goat anti-axonin-1 Fab fragments and a FITC-labeled donkey anti-goat IgG (D). For examination of the cells, a confocal laser-scanning microscope equipped with an argon/krypton laser was used. Texas Red was detected using the 568-nm band-pass excitation filter (A,B) and FITC with the 488-nm band-pass excitation filter (C,D), minimizing cross-talk between the two fluorochromes.

sults in extensive co-capping of molecule B (axonin-1), as shown by counter-staining with goat anti-B (axonin-1) Fab fragments on the same cell (Fig. 9.5.3C). As expected, no capping or co-capping is detected in the absence of primary antibody against molecule A (NgCAM).

Time Considerations Preparation of affinity column The preparation of an affinity column takes 1 working day. Because affinity columns have to be loaded slowly, it is convenient to load large volumes of protein solution overnight to have the column ready for elution the following day. However, ensure that the column never runs dry. Transfection The entire transfection procedure can be performed in - %$ # % $ 718 # $ # % & !!& 5 $ %&&#C$ %%. $33(1',; "&5 '!" !' −71°4 (2 & 7KHSURWHLQFRQFHQWUDWLRQVKRXOGEHWRPJPO %$6,& 35272&2/

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Matrix Metalloproteinases

UNIT 10.8

This unit describes a set of methods that are relatively unique to studies of matrix metalloproteinases (MMPs) and their inhibitors (TIMPs, α2M), including cell-mediated dissolution of type I collagen fibrils (see Basic Protocol 1), direct and reverse zymography (see Basic Protocols 2 and 3), enzyme capture techniques based on α2-macroglobulin (α2M) and TIMP-1, and -2 (see Basic Protocol 4 and Alternate Protocol), and detection and demonstration of cryptic thiol groups in MMP precursors (see Basic Protocol 5). Support Protocols are included for preparation (see Support Protocol 1) and labeling of collagen (see Support Protocol 2). DISSOLUTION AND DEGRADATION OF COLLAGEN FIBRILS BY LIVE CELLS

BASIC PROTOCOL 1

Comparatively few methods allow detailed analysis of how live cells orchestrate MMP and inhibitor functions in the degradation and remodeling of extracellular matrices. The methods described in this protocol were developed to study the function of matrix metalloproteinases (MMPs) in the degradation of type I collagen fibrils by live cells under controlled but readily variable conditions. In its simplest form, cells are seeded on a few-micron-thick film of reconstituted collagen fibrils, then incubated for a period of 1 to 7 days. The progressive dissolution of the film under the cell layer—in response, e.g., to changing environmental conditions, inducing agents, or inhibitors—may be monitored directly and related to the level of expression of key components of the requisite proteolytic machinery. The system is readily manipulated in a number of ways: by induction/repression of transcription of components of the signaling and effector systems; by transfection of new genes of potential importance to the process; or by selective or specific blocking strategies using antisense-, MMP-specific inhibitor–, or antibody-based approaches. The limited susceptibility of type I collagen fibrils to cleavage and dissolution by MMPs permits one to narrow the scope of the investigation to a small number of (“collagenolytic”) enzymes. This characteristic also makes it a realistic objective to dissect the entire sequence or set of reactions involved in cell-mediated dissolution of collagen fibrils, starting from the initial engagement of cell surface receptors by cytokines, growth factors, and other catabolic reagents, through the final enzymatic cleavage, dissolution, and disposal of the substrate. Important questions that may be addressed using this approach include the following: a. What enzymes are actually involved in the cleavage reaction itself and in the precursor activation steps? b. How do cells regulate the activity of the enzymes? c. What role is played by TIMPs in modulating, containing, and blocking the response? d. What is the ultimate fate of the collagen chains and peptides generated as a result of proteolysis? Recent studies have shown that type I collagen (in solution or in reconstituted fibrillar form) may be cleaved by a larger number of enzymes than previously anticipated, including the three classical “collagenases,” MMP-1, MMP-8, and MMP-13 (BirkedalHansen et al., 1993; Knäuper et al., 1996). In addition, reports suggest that MMP-14 (Ohuchi et al., 1997) and TIMP-free MMP-2 may also dissolve collagen fibrils at meaningful rates under physiologic conditions (Aimes and Quigley, 1995). It is of note that although the three classical collagenases (MMP-1, MMP-8, and MMP-13) were Extracellular Matrix Contributed by Jack Windsor, Anne Havemose Poulsen, Susan Yamada, Guy Lyons, Bente Birkedal-Hansen, William Stetler-Stevenson, and Henning Birkedal-Hansen

10.8.1

Current Protocols in Cell Biology (2002) 10.8.1-10.8.23 Copyright © 2002 by John Wiley & Sons, Inc.

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discovered because of their ability to dissolve reconstituted fibrils of type I collagen, no definitive proof has yet been rendered that cleavage of collagen fibrils is indeed the exclusive or even prevailing biologic function of any of these enzymes. Admittedly, the evidence seems compelling based on a large number of in vitro studies. Earlier versions of this method have been published (Birkedal-Hansen, 1987; BirkedalHansen et al., 1989, 1993; Lin et al., 1987). The isolation and purification techniques of type I collagen and the methods for formation of reconstituted hydrated gels of type I collagen have been described elsewhere in detail (Birkedal-Hansen, 1987). The method relies on the ability of neutral solutions of type I collagen in an appropriate concentration range (0.1 to 5 mg/ml) to form hydrated gels of reconstituted fibrils by heating to 37°C. The method also takes advantage of the observation that such loose hydrated gels may be collapsed by gentle air-drying into a thin film of uniform, densely packed, randomly oriented fibrils which remain as highly resistant to proteolysis by enzymes such as trypsin, chymotrypsin, and plasmin as hydrated gels or natural fibrils (Fig. 10.8.1). Trypsin, which is often used as a standard for testing the resistance of collagen fibrils to “unspecific” proteolytic cleavage, is unable to dissolve the collagen fibril films prepared as described. The same is true for a large number of proteinases of all four classes, and it is this unique resistance to proteolysis which renders this assay system particularly valuable as it greatly reduces the number of proteinases that are involved in the cleavage/dissolution reaction. Several variants of the method may be used. While the authors often prefer (for ease of presentation and interpretation) to seed the cells in a small button in the middle of a much larger dish (35 mm; Fig. 10.8.1A, middle) in order to maintain medium excess, it is also possible to seed the cells over the entire collagen-coated surface, although a confluent monolayer rapidly exhausts the medium. The collagen may be used in its natural state or labeled either with radioactive or fluorescent tags to facilitate monitoring (see Support Protocol 2), retrieval, and quantification of dissolved collagen chains and fragments. Depending on the casting conditions, collagen films may be generated with a thickness down to 1 to 2 µm, which is approximately the thickness of a single layer of well-spread cells. Most cell types seeded on this film spread within minutes to hours, although often more slowly than on plastic. Cells that express an appropriate complement of MMPs either constitutively or after exposure to cytokines and growth factors (or phorbolester) progressively dissolve the underlying fibril coating, and, within 24 to 96 hr, clear a path to the

A

B

1 µm

C

0.5 µm

Collagen Fibrils

Figure 10.8.1 Reconstituted collagen fibril film. (A) Rat tail tendon type I collagen is polymerized by heat gelation. The gel is air dried and reduced in thickness to a few microns. Cells are seeded in the middle of the plate and incubated with culture medium. After incubation, the cells are removed and a clearing beneath the cell layer is exposed by staining with Coomasie blue. (B) The air-dried collagen fibril film consists of uniform, randomly oriented reconstituted fibrils (C). Detail of cell attached to the collagen fibril film.



plastic surface (Fig. 10.8.1A, lower; Fig. 10.8.2). Coomassie blue staining of the residual collagen fibril film after removal of the cells is usually sufficient to visualize the dissolution of the underlying film (Fig. 10.8.2). Materials 3 mg/ml rat tail tendon type I collagen in 13 mM HCl (see Support Protocol 1) 13 mM HCl, 4°C Neutralizing buffer (see recipe), 4°C Phosphate-buffered saline (PBS) without Ca2+ and Mg2+ (CMF-PBS; APPENDIX 2A) supplemented with 100 U/ml penicillin G and 100 µg/ml streptomycin sulfate Cells of interest (e.g., fibroblasts, keratinocytes, or tumor cells) DMEM (APPENDIX 2A) supplemented with 100 U/ml penicillin G and 100 µg/ml streptomycin sulfate with and without 10% (v/v) FBS (or other medium appropriate for cell type) Growth factors/cytokines: e.g., IL-1β, TNF-α, TGF-α, or TPA; or phorbol ester (12-O-tetradecanoylphorbol-13-acetate, TPA, or phorbol myristate acetate, PMA) 1% (v/v) Triton X-100 0.5% (w/v) trypsin/0.53 mM EDTA (Invitrogen) Coomassie blue stain (see recipe) 6-well cell culture plates Additional reagents and equipment for trypsinizing and counting cells (UNIT 1.1) Prepare collagen-coated plates 1. To cast one 6-well plate, dilute 1 ml of 3 mg/ml type I collagen stock solution with 7 ml of 13 mM HCl at 4°C. Mix the collagen solution with 2 ml of cold neutralizing buffer in a precooled test tube either by gently pipetting up and down while avoiding formation of air bubbles (which will form defects in the gel) or by gently inverting the tube several times.

ANNE HAVEMOSE-POULSEN, Thesis, Copenhagen, 1995

Figure 10.8.2 Dissolution of collagen fibrils by live adherent cells. Fibroblasts seeded in the center of the well dissolve the underlying collagen fibril film. Upper left panel shows scanning electron micrograph of fibroblast attached on collagen fibril film. Recreated from Havemose-Poulsen et al. (1998).

Extracellular Matrix


Supplement 13

The neutralizing buffer is designed to bring the pH of the solution to 7.4 (check with pH paper). The concentration of this buffer is 0.2 M inorganic phosphate (as Na2HPO4 /NaH2PO4) and 0.47 M NaCl. The final collagen concentration is 300 ìg/ml in 40 mM Pi /∼0.10 M NaCl. Since pH dramatically influences the gelling properties of the collagen solution it is often advantageous to first test the efficacy of the neutralizing buffer by mixing 4 volumes of 13 mM HCl with one volume of neutralizing buffer and checking the final pH (7.4). The final thickness of the collagen film depends on the concentration of the collagen solution. A 300 ìg/ml solution dispensed at a volume of 1.5 ml (35 mm) dish yields a film of 1.5 to 2.0 ìm in thickness after drying. Higher concentrations yield thicker films. The lower concentration limit for proper gelling is around 100 ìg/ml using rat tail tendon collagen prepared as described (see Support Protocol 1) but somewhat higher (500 ìg/ml) with commercial type I collagen preparations.

2. Immediately after mixing, add a 1.5-ml aliquot of neutralized collagen solution to each well of 6-well culture plate. Rotate the plate to permit the collagen to cover the entire well bottom evenly. Incubate in humidified incubator for 2 hr at 37°C. Avoid movement of gel and plate during gelling. 3. Remove plate from incubator, remove lid, and place at room temperature in an air stream (laminar flow hood) overnight (during this process the gel dries down to a thin film). Wash three times with distilled water, each time for 30 min at room temperature or 37°C, to remove salt crystals formed during the drying (check efficacy of washing step using a phase-contrast microscope). Dry again overnight in laminar flow hood and check for absence of residual salt crystals. It is important that all salt crystals are removed by washing before the plates are used.

4. Add 2 ml CMF-PBS or DMEM supplemented with penicillin/streptomycin. Store in this solution in incubator at 37°C or in refrigerator at 4°C in closed plastic bag to prevent evaporation. The plate can be stored in this manner for up to 2 weeks as long as evaporation is avoided.

5. Immediately before seeding cells, remove medium from wells by aspiration and wash with 2 ml distilled water for 30 min. Remove water and leave plate to air dry in hood. Plate cells 6. Trypsinize and count cells (see UNIT 1.1), then dilute cell suspension to the appropriate concentration in DMEM/10% FBS, or in medium appropriate for the cell type being used. Best results are obtained with 10,000 to 50,000 cells in a 25-ìl aliquot, using a cell suspension of 4 × 105 to 2 × 106 cells/ml, somewhat depending on cell size. The intent is to form a coherent monolayer in a small central button (Fig. 10.8.1A, middle).

7. Deliver a 25-µl aliquot to the center of the well without touching the fragile collagen film. Fill plate volume between wells with distilled water to avoid evaporation during seeding and attachment. Place plate in plastic box on wet paper towels to avoid evaporation, and then place in incubator for 5 hr or overnight at 37°C to allow cells to attach. 8. Add to each well 2 ml DMEM/10% FBS or appropriate medium and incubate overnight at 37°C to allow cell spreading. Some cell types can be transferred immediately to serum-free medium while others require overnight incubation in serum-supplemented medium. Matrix Metalloproteinases

Once the cells are spread, incubation may be performed either with or without serum. The result depends somewhat on cell type. Some cells tend to detach in the absence of serum



while others can be maintained for 2 to 3 days in complete absence of serum while degrading the collagen fibril matrix.

9. If the experiment is to be performed in the absence of serum, thoroughly and repeatedly wash with CMF-PBS or serum-free DMEM for 10 min at 37°C, to remove remnants of serum. Some cells may require special media formulations, i.e., keratinocytes. Most fibroblast strains do well under serum-free conditions either in DMEM or DMEM/F12 (1:l).

Induce expression of MMPs 10. Induce cells for expression of MMPs at this stage by including in the medium cytokines such as IL-1β (10−9M), TNF-α (10−8M), TGF-α (10−8M), or TPA (1 to 2× 10−7 M). Alternatively, cytokine or TPA induction may be achieved during the last 24 hr before trypsinization and seeding. If incubated under serum-free conditions, plasminogen may be added to the medium. Some cells respond to exposure to plasminogen by greatly accelerating the rate of dissolution, while others do not. If desired, plasminogen is added from a stock solution in CMF-PBS to give a final concentration of 4 ìg/ml. Human plasminogen is either purchased from one of several commercial sources (i.e., Pharmacia Hepar or Sigma-Aldrich) or prepared as described (Deutsch and Mertz, 1970) from outdated human plasma by lysine-sepharose chromatography.

11. Incubate the plates at 37°C for 1 to 4 days (or up to 7 days) depending on the experimental design. Follow the progress of the process with a phase-contrast microscope. To avoid evaporation it may be advantageous to fill the volume between the wells with sterile distilled water.

Stain plate and quantitate results 12. In order to visualize the dissolution of the film beneath the cell layer, remove the cells either by dissolution in 1% (v/v) Triton X-100, by 0.5% trypsin/0.53 mM EDTA (10 min, 37°C), or by a combination thereof. Avoid use of SDS, which dissolves the collagen fibril film as well as the cells.

13. Rinse the wells with distilled water. 14. Stain with Coomassie blue stain for 5 to 15 min to visualize residual collagen film, then wash three times with distilled water. 15. Destain in distilled water for 30 min (or perform three quick washes with water) and finally allow plates to air dry. After drying the plates, they can be stored indefinitely (Fig. 10.8.2). In order to follow the progressive dissolution of the collagen fibril film it is advantageous to terminate sample wells on consecutive days and to contrast the dissolution after 1, 2, 3 ... days. If desired the plates can be scanned directly into Adobe Photoshop using a scanner capable of scanning transparent originals.

16. Determine the extent or rate of dissolution of the substrate The degree of dissolution at the conclusion of the experiment may be measured photometrically in Coomassie blue–stained plates by measuring the absorption of light in a conventional light microscope equipped with a exposure (photo)meter as described in Havemose-Poulsen et al. (1998). The relationship between amount of collagen present on



Supplement 13

the plate and exposure time is strictly linear at least up to three times the collagen layer thickness used in this protocol. Alternatively, if the cells are seeded evenly as a confluent monolayer over the entire collagen-coated well bottom (see below), progression may be monitored daily by removal of aliquots of medium and measuring the release of collagen chains and peptides. To this end the collagen may be labeled either with 3H (Birkedal-Hansen, 1987; Birkedal-Hansen and Danø, 1981) or with fluorescent tags (Ghersi et al., 2001). This approach is less useful if the cells are seeded in a small 2- to 4-mm button at the center of the well, because the background release of radioactivity and fluorescent label from the entire film compromises the sensitivity of the analysis (typically only 10% to 20% of the collagen fibril film is covered by cells in this variation). SUPPORT PROTOCOL 1

PREPARING RAT TAIL TENDON COLLAGEN TYPE I Methods for isolation and preparation of rat tail tendon type I collagen have been described in detail elsewhere (Birkedal-Hansen, 1987; Birkedal-Hansen and Danø, 1981). Alternatively, rat, bovine or human type I collagen may be purchased from Becton Dickinson Biosciences Discovery Labware. Briefly, tendons teased from rat tails are washed with distilled water and with 0.5 M NaCl. The acid-soluble collagen fraction is then extracted in 0.5 M acetic acid, and type I collagen is purified by sequential salt precipitation at neutral to slightly alkaline pH, first with 5% NaCl, then (after redissolution in acetic acid) with 0.02 M Na2HPO4. NOTE: All protocols using live animals must first be reviewed and approved by an Institutional Animal Care and Use Committee (IACUC) or must conform to governmental regulations regarding the care and use of laboratory animals. Materials Tails of ∼400 g rats (freshly removed or stored frozen at −80°C) 0.5 M NaCl in 50 mM Tris⋅Cl, pH 7.4 (see APPENDIX 2A for Tris⋅Cl) 5 mM, 50 mM, and 0.5 M acetic acid NaCl (solid) 0.02 M Na2HPO4 13 mM HCl Neutralizing buffer (0.2 M NaPi) Glass wool or cheesecloth 500-ml centrifuge bottles High-speed centrifuge (Sorvall with SS-34 and GSA rotors, or equivalent centrifuge and rotors) 10,000 to 14,000 MWCO dialysis membrane One large (25-liter) or several smaller (4-liter) dialysis tanks Sterile scissors 125-ml glass Wheaton bottles Additional reagents and equipment for dialysis (APPENDIX 3C) Extract collagen 1. Skin 10 to 20 rat tails and place tails on ice. Break tails at joints and tease out individual collagen fibers. Wash in large volume distilled water (2 to 3 liter) for 1 hr with agitation. Change wash water three to four times. The yield is 200 to 400 mg collagen per rat.


2. Extract overnight at 4°C with agitation in 2 liters of 0.5 M NaCl/50 mM Tris⋅Cl, pH 7.4. Discard extract and repeat step.



3. Discard second salt extract and wash collagen fibers extensively (over a 3-hr period with change two to three times per hr) in distilled water to remove salt. 4. Extract overnight at 4°C with slow agitation in 2 liters of 0.5 M acetic acid. 5. Remove insoluble remnants by filtration through glass wool or cheesecloth, then centrifuge in 500-ml bottles for 30 min at 11,000 × g (8500 rpm in a GSA rotor), 4°C. Add solid NaCl little by little to a final concentration of 5% w/v (50 g/liter) under constant vigorous stirring. When the salt is completely dissolved, turn off stirrer, cover beaker, leave in cold room overnight and let precipitate gather at bottom of vessel. The collagen immediately starts to precipitate upon addition of the salt.

6. Collect precipitate by centrifugation for 30 min at 11,000 × g, 4°C. Discard supernatant. 7. Redissolve collagen by adding 450 ml of 0.5 M acetic acid to first centrifuge bottle, transfer liquid to the second bottle, and so on, until collagen is redissolved/redispersed into ∼900 to 1000 ml in 0.5 M acetic acid. 8. Stir vigorously overnight at 4°C until collagen is completely dissolved. If not dissolved overnight, add more acetic acid and bring volume up to 1600 to 1800 ml.

Dialyze collagen solution 9. Place collagen solution, 300 to 400 ml at a time, in dialysis bags. Dialyze in tank against 25 liters of 0.5 M acetic acid, then for 3 to 4 days against 50 mM acetic acid. Change daily and mix content of bags. See APPENDIX 3C for additional details on dialysis.

10. Dialyze against several changes of 0.02 M Na2HPO4 in 25-liter tank over the next 72 hr. Precipitation should happen as fast as possible, so change solution frequently in the beginning and massage bags frequently to facilitate even distribution of reagents. The collagen precipitates as a thick white gel.

11. Harvest precipitate by centrifugation in 500-ml bottles for 30 min at 11,000 × g, 40°C. Redissolve collagen in 0.5 M acetic acid by vigorous stirring overnight at 4°C. 12. Dialyze 3 to 4 hr against 0.5 M acetic acid, then overnight against 50 mM acetic acid, and, finally, overnight against several changes of 5 mM acetic acid. 13. Centrifuge 1 hr at 11,000 × g, 4°C. Lyophilize supernatant and store in dessicator at 4°C, −20°C, or −80°C. 14. Redissolve as follows. a. Weigh out no more than 150 mg collagen. b. Cut into 1-cm pieces with sterile scissors. c. Place collagen pieces into a 125-ml glass Wheaton bottle that has been autoclaved with a stir bar inside. d. Add cold 13 mM HCl to make a 3 mg/ml solution and stir briskly at 4°C with occasional shaking for ∼24 hr. The collagen solution should be slightly opalescent.

15. Centrifuge solution for 20 min at 50,000 × g (20,000 rpm in an SS-34 rotor), 4°C, to remove any insoluble material, if necessary. Note that the solution remains somewhat opalescent even after centrifugation. This solution may be stored for months at 4°C. Freezing should be avoided.



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SUPPORT PROTOCOL 2

LABELING OF COLLAGEN Rat tail tendon type I collagen may be labeled using 3H-acetic anhydride as described in detail in Birkedal-Hansen and Danø (1981) and Birkedal-Hansen (1987), or with fluorescent reagents. The following fluorescent labeling method was adapted from a technique devised by the Chen laboratory (G. Ghersi and W.T. Chen, unpub. observ.). Materials 3 to 5 mg/ml rat tail tendon type I collagen originally dissolved in or dialyzed into 20 mM acetic acid (see Support Protocol 1) DMEM Borate buffer: 0.05 M NaB4O7⋅10H2O, pH 9.3, containing 0.04 M NaCl, sterile 2 to 3 mg/ml tetramethylrhodamine-5-(and 6)-isothiocyanate (TRITC) or fluorescein isothiocyanate (FITC) in DMSO Phosphate-buffered saline (PBS; APPENDIX 2A) 0.2 M acetic acid 10-cm culture dishes 1. Mix 8 ml of 3 to 5 mg/ml rat tail tendon type 1 collagen with an equal volume of DMEM, transfer to a 10-cm diameter dish, and incubate at 37°C overnight to form a 3-mm thick gel. Rat tail tendon type I collagen may be prepared as described in this unit or purchased from Becton Dickinson Biosciences Discovery Labware; bovine skin and human placental type I collagen is also available from the same supplier.

2. Wash for 1 hr with sterile borate buffer at room temperature, with shaking. 3. Remove buffer; replace with 10 ml borate buffer plus TRITC or FITC at 2 to 3 mg/ml in a small volume of DMSO. Incubate at room temperature 15 to 20 min or until the dye diffuses through the gel. Protect from light from this point onward. 4. Wash with multiple changes of PBS for several days to remove free dye; wash salt out with water and redissolve in 0.2 M acetic acid at 4°C Collagen is labeled in the fibrillar state so that sites important for subsequent alignment and gelling are not being blocked by the labeling procedure. Consequently, collagen labeled in this fashion readily dissolves in dilute acid and gels again upon neutralization and mild heating. Depending on the need, the fluorescently labeled collagen may be diluted up to 10-fold with unlabeled rat tail tendon collagen and still yield a strong enough signal for quantification. BASIC PROTOCOL 2


GELATIN/CASEIN ZYMOGRAPHY Zymographic methods are designed to analyze the proteolytic capacity of latent and active MMPs (Heussen and Dowdle, 1980; Birkedal-Hansen and Taylor, 1982; Birkedal-Hansen, 1987). This set of techniques is based on a number of unique properties of MMPs: (1) MMPs retain (or refold to display) catalytic activity after electrophoresis in SDS-containing buffers as long as heating and reduction are avoided (Birkedal-Hansen and Taylor, 1982); (2) brief exposure to SDS opens the “cysteine switch” (Springman et al., 1990; Van Wart and Birkedal-Hansen, 1990) so that both precursor and proteolytically truncated (“activated”) forms of the enzyme display catalytic activity; and (3) MMP catalytic activity is reversibly inhibited by SDS and readily restored when SDS is removed by washing with Triton X-100 (Birkedal-Hansen and Taylor, 1982). It is therefore possible to resolve a heterogenous group of MMPs and non-MMPs in SDS-containing gels copolymerized with a suitable substrate (gelatin, casein), remove the SDS, and develop (without distinction) the spontaneous or latent catalytic activity associated with each electrophoretic band. After appropriate incubation (to allow for proteolysis), the discrete



bands of substrate lysis are made visible by Coomassie blue staining of the gel (Fig. 10.8.3). SDS opens the “cysteine switch” but instantly inhibits the switch-open enzyme and blocks autolytic truncation normally associated with activation. The proenzyme bands therefore migrate at their expected high-molecular weight, but display proteolytic activity because the switch is unable to again “close” after removal of the SDS with Triton X-100. Zymography using gels containing 0.1 to 1.0 mg/ml gelatin are by far the most sensitive. Gels may either be purchased (Invitrogen) or prepared as described below. Gelatin works particularly well for MMP-2 and MMP-9, whereas MMP-1, MMP-3, MMP-7, MMP-8, and MMP-10 are better identified in casein-containing gels. Materials Gelatin (bovine skin, Sigma-Aldrich type B6-6269) or casein (Sigma-Aldrich, technical, C-0376) 2.0 M Tris⋅Cl, pH 8.8 (APPENDIX 2A) 30/0.8 acrylamide/bisacrylamide (UNIT 6.1) Glycerol 10% (w/v) SDS (APPENDIX 2A) TEMED 10% (w/v) ammonium persulfate MMP preparation of interest (for standards, use 1 to 5 ng purified MMP) 5× electrophoretic sample buffer (see recipe) Electrophoretic running buffer (see recipe) Gel washing buffers 1 to 4 (see recipe) Coomassie blue stain (see recipe) Gel destaining solution (see recipe) 50-ml centrifuge tubes 57°C water bath Whatman no. 1 filter paper or 0.5-µm syringe filter Gel washing tray of appropriate size Additional reagents and equipment for preparing SDS-PAGE gels according to Laemmli (UNIT 6.1) NOTE: The following procedure is based on a standard 10% SDS-PAGE according to Laemmli (Laemmli, 1970; UNIT 6.1) using a 4% stacking gel and a pH 8.3 running buffer. It is important to avoid heating and/or reduction during sample preparation and running of the gel. 1. Weigh out appropriate amount of gelatin (for 0.1 to 1.0 mg/ml final concentration) or casein (for 1.0 mg/ml final concentration) and place in a 50-ml centrifuge tube. 2. For every 10 ml of solution to be prepared, add 4 ml of 2.0 M Tris⋅Cl, pH 8.8, and 6 ml water. Dissolve by heating in 57°C water bath. Filter through Whatman no. 1 filter paper or syringe filter. 3. Prepare the 10% resolving gel (also see UNIT 6.1) by adding the following to 10 ml filtered gelatin or casein solution (0.2 to 13 mg/ml in 0.8 M Tris⋅Cl, pH 8.8; see step 2): 6.6 ml 30/0.8 acrylamide/bisacrylamide 2 g glycerol 0.2 ml 10% (w/v) SDS 13.3 µl TEMED 67 µl 10% (w/v) ammonium persulfate Pour gel as described in UNIT 6.1.



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4. Prepare 4% stacking gel by combining the following (also see UNIT 6.1): 1 ml 30/0.8 acrylamide/bis acrylamide 0.36 ml 2 M Tris⋅Cl, pH 6.8 75 µl 10% (w/v) SDS 6 ml H2O 8 µl TEMED 60 µl ammonium persulfate Pour gel as described in UNIT 6.1. 5. Mix 1 part MMP solution (partially or fully purified MMP, culture medium, concentrated culture medium, or other preparation containing MMP) with 4 parts of 5× sample buffer (final concentration, 1% SDS). Incubate at room temperature for 10 min, then load 20 to 30 µl into each well of the 15-ml gel prepared in steps 3 and 4. Alternatively, load 20 to 30 ìl per well of an Invitrogen minigel.

6. Run gel at 200 V for 35 to 45 min or until dye front reaches bottom of gel using electrophoretic running buffer, pH 8.3.

casein zymography

gelatin zymography culture medium

A

B

D + 0

+ 5

+ 15

+ 30

– APMA 30 min

coomassie blue

proMMP-9 proMMP-2

MMP-10 MMP-3

S

Q 00 E2

94

-TY

H1

LD

ut

E

WI

C

m

w

ild

ty

an

t

pe

PE

MMP-2

lung extract coomassie blue


Figure 10.8.3 Zymography. (A) Zymography using gelatin-containing polyacrylamide gel. Culture medium containing proMMP-2 (left) or MMP-2/proMMP-2 and proMMP-9 (right). The proenzymes display catalytic activity because exposure to SDS during sample preparation opens the cysteine switch. (B) Detail showing conversion of proMMP-2 to MMP-2 by exposure to aminophenylmercuric acetate. From Caterina et al. (2000). (C) MMP-2 and MMP-9 activity in extracts of lungs of wild-type mice (left) or mice in which the TIMP-2 gene has been mutated to inactive form (modified from Caterina et al., 2000). (D) Zymography using casein-containing polyacrylamide gel. (pro)MMP-3 and MMP-10 cleave casein embedded in the gel (modified from Windsor et al., 1993). (E) Casein zymogram of mutant and wild-ype MMP-1. Inactivation of catalytic activity by mutation of catalytic site glutamic acid (E) to glutamine (Q) that abolishes casein cleavage. A histidine to serine replacement outside the active site does not. Modified from Windsor et al. (1994).



7. Remove gel from electrophoretic apparatus and place in an appropriately sized container. Wash four times, 20 min each, successively, in washing buffers 1, 2, 3, and 4 at room temperature. Shake gently throughout. 8. Replace the last wash buffer with fresh washing buffer 4 and incubate 1 to 24 hr at 37°C. A few hours of incubation is usually sufficient to reveal MMP-2 and MMP-9 by gelatin zymography. Overnight incubation is required to visualize MMP-1, MMP-3, MMP-13, MMP-7, and MMP-10 by casein zymography.

9. Stain gel with Coomassie blue stain for 30 min and destain with gel destaining solution for several hours until bands are clear. Typical results are shown in Figure 10.8.3.

REVERSE ZYMOGRAPHY Reverse zymography is specifically designed to identify electrophoretic bands which display MMP-inhibitory activity. The method is based on incorporation of both MMP activity and gelatin into the running gel. During the ensuing incubation, the SDS-activated MMP-2 (gelatinase A) cleaves the substrate everywhere in the gel except in and immediately around bands with inhibitory activity such as TIMPs. This method yields well resolved bands of TIMP-1, TIMP-2, TIMP-3, and TIMP-4, as well as mutant forms of these inhibitors (Fig. 10.8.4). The following protocol is developed by the Stetler-Stevenson laboratory and used in the authors’ laboratory as well. Quantities are for a 15-ml gel, but can be scaled down as necessary.

BASIC PROTOCOL 3

Materials 8.7 mg/ml gelatin solution (see recipe) MMP-2 (Gelatinase A) 5× electrophoretic sample buffer (see recipe) 2.5% (w/v) Triton X-100 Incubation solution (see recipe) Additional reagents and equipment for “forward” zymography (see Basic Protocol 2) 1. Prepare separating gel (17%), copolymerizing gel with gelatin (2.5 mg/ml) and purified gelatinase A (MMP-2), by mixing the following components (also see UNIT 6.1): 4.2 ml 8.7 mg/ml gelatin solution 0.16 µg/ml (final concentration) gelatinase A (MMP-2) 8.25 ml 30/0.8 acrylamide/bisacrylamide 2.1 ml H2O 0.29 ml 10% (w/v) SDS 7.3 µl TEMED 73 µl 10% (w/v) ammonium persulfate Pour gel as described in UNIT 6.1. Purified MMP-2 may be replaced with culture medium of cells which secrete this enzyme. The appropriate amount should be determined by trial and error.



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+/+

+/–

–/– TIMP-1 TIMP-? TIMP-2

Mutant TIMP-2

Figure 10.8.4 Reverse zymography. Inhibition of MMP-2 by TIMPs. Skin fibroblast culture medium obtained from wild-type, hemizygous, or homozygous TIMP-2-deficient mice was resolved by SDS-PAGE in a gel also containing MMP-2 and gelatin. During incubation, MMP-2 cleaves gelatin unless inhibited by electrophoretic bands of TIMPs. The TIMP-2-deficient cells still express TIMP-1 and unidentified component below TIMP-1, possibly TIMP-3 and a weakly inhibitory truncated mutant of TIMP-2. Modified from Caterina et al. (2000).

2. Prepare 5% stacking gel by combining the following (also see UNIT 6.1): 1.66 ml 30/0.8 acrylamide/bis acrylamide 1.55 ml 2 M Tris⋅Cl, pH 6.8 125 µl 10% (w/v) SDS 8.2 ml H2O 10 µl TEMED 200 µl ammonium persulfate Pour gel as described in UNIT 6.1. 3. Mix samples with 5× sample buffer for reverse zymography. Incubate at room temperature for ≥10 min, then load 20 to 30 µl into each well of the gel. 4. Run gel at 150 V until buffer front reaches bottom of gel. 5. Remove gel and wash in three changes of 2.5% Triton X-100, for 2 hr with gentle shaking. 6. Incubate overnight at 37°C in incubation solution. 7. Stain gel with Coomassie blue stain for 20 min and destain in gel destaining solution for several hours until background is clear. Typical results are shown in Figure 10.8.4.




á2-MACROGLOBULIN (á2M) CAPTURE α2M capture is particularly valuable because it permits assessment of the proteolytic competence and activity of single bands of MMPs in a mixture of many partially or fully processed forms. The method was originally devised (Birkedal-Hansen et al., 1976) for separation of complexes from unreacted forms by molecular sieve chromatography (Fig. 10.8.5), but it is even more valuable when combined with electrophoretic analysis. The protocol is based on the observation that α2M forms complexes only with catalytically competent forms of MMPs. Unactivated MMP precursors or forms devoid of catalytic activity are not captured. The ensuing separation by SDS-PAGE permits easy identifica-

BASIC PROTOCOL 4

MMP-1 proMMP-1

α 2-macroglobulin

AP M A AP M A TR + α 2M YP SI TR N YP S E2 IN 00 + α Q 2M E2 00 Q + H α 19 4S 2M H 19 4S + α2 M

α2 M

Vo

α2M-MMP-1 COMPLEX

52K 45K 42K

MMP-1 1

2

3

4

5

6

7

8

9

10

Figure 10.8.5 α2-macroglobulin (α2M) capture. The capture technique is based on the property that proteolytic cleavage of the α2M bait region results in conformational and eventually covalent capture of the attacking proteinase. Because of the large disparity in molecular weight, captured and free froms of the proteinase may be separated either by molecular sieve chromatography (upper panel Birkedal-Hansen et al., 1976) or by SDS gel electrophoresis (lower panel; Windsor et al., 1994). Covalently bound proteinase is not released and is readily identified by appropriate antibody staining. Latent or inactive proteinases are not captured. The method therefore discriminates between enzyme forms with and without catalytic activity at the moment of testing. The panel shows wild-type and mutant forms of human MMP-1. Samples in lanes 3, 4, and 7 to 10 are pretreated with p-aminophenylmercury acetate (APMA). Samples in lanes 5, and 6 were preactivated by trypsin. Modified from Windsor et al. (1994).



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tion of bands which have been captured and moved to the top of the gel because of the large molecular mass of the α2M (Fig. 10.8.5). Bands that escape capture continue to migrate at their usual position. Complexes formed with α2M are covalent and therefore not easily dissociated. The ability of α2M to discriminate between latent and overtly active forms of the enzyme is a result of the α2M inhibition mechanism. α2M is inert until the attacking proteinase cleaves a peptide bond in the bait region. This cleavage results in rapid conformational change and liberates a thiol ester which covalently bonds to and immobilizes the attacking proteinase. Materials MMP solution to be tested 2 to 3 mg/ml purified α2M in 50 mM Tris⋅Cl standard buffer (see recipe for buffer) 100 µg/ml TPCK-treated trypsin (e.g., Sigma) in 50 mM Tris⋅Cl standard buffer (see recipe), pH 7.4 1.0 mg/ml soybean trypsin inhibitor in 50 mM Tris⋅Cl standard buffer (see recipe), pH 7.4 5× electrophoretic sample buffer (see recipe) Antibodies to MMPs of interest Nitrocellulose paper Additional reagents and equipment for SDS-PAGE according to Laemmli ( UNIT 6.1) immunoblotting (UNIT 6.2) 1. Mix one half of the test solution with a sufficient volume of 1.5 mg/ml α2M to achieve a ≥10× molar ratio of inhibitor to MMP. Incubate 15 min at room temperature. 2. To compare “activated” and “unactivated” samples, preincubate the other half of the test sample with 10 µg/ml trypsin (added from 100 µg/ml stock) for 10 min at room temperature, then add 100 µg/ml soybean trypsin inhibitor (added from 1 mg/ml stock). Incubate separately with α2M as described in step 1. Commercial sources of α2M are available but should always be checked for activity by titration with trypsin using a suitable substrate (Sottrup-Jensen and Birkedal-Hansen, 1989). Alternatively, the inhibitor may be prepared by standard techniques as described by Sottrup-Jensen and Birkedal-Hansen (1989) and Sottrup-Jensen et al. (1983). Activation with trypsin prior to addition of α2M often yields more complete capture than with organomercurials—e.g., NH2PheHgAc (APMA)—which seem to gradually inactivate α2M. Samples preincubated with organomercurials, however, still show partial capture.

3. Mix with 5× electrophoretic sample buffer (final concentration, 1% w/v SDS, 2.5% v/v 2-ME) without heating, resolve by by SDS-PAGE using a 10% gel according to Laemmli (Laemmli, 1970; UNIT 6.1). 4. Transfer to nitrocellulose paper and stain with appropriate MMP antibody using conventional immunoblotting techniques (UNIT 6.2). Typical results are shown in Figure 10.8.5. ALTERNATE PROTOCOL


TIMP CAPTURE Complexes formed with TIMPs are not covalent, although several, but not all, withstand exposure to low concentrations of SDS, as originally observed by DeClerck et al. (1991), who first pioneered this technique. This method detects many but not all activated MMPs that bind TIMPs, including MMP-1 (collagenase-1), MMP-3 (stromelysin-1), MMP-7 (matrilysin), MMP-10 (stromelysin-2), and MMP-13. Detection is most conveniently done by immunoblotting using specific antibodies to the two complex components (MMP and TIMP; Fig. 10.8.6). The method described below is the authors’ adaptation of the



method of DeClerck (DeClerck et al., 1991). It is based on capture with TIMP-1, but TIMP-2 capture works just as well. Materials 0.1 to 1.0 mg/ml TIMP-1 (Oncogene Research Products, Chemicon International; also see Bodden et al., 1994) in 50 mM Tris⋅Cl standard buffer (see recipe), pH 7.4 10.0 mM NH2PheHgAc (APMA; Sigma) in Tris⋅Cl standard buffer (see recipe), pH 7.4 Electrophoretic sample buffer (see recipe, but use only 0.5% w/v SDS) Antibodies to MMPs and TIMP-1 of interest (Calbiochem, Chemicon International) Additional reagents and equipment for SDS-PAGE (UNIT 6.1) and immunoblotting (UNIT 6.2) 1. Incubate control and activated samples with 40 to 100 µg/ml TIMP-1 (added from 0.1 to 1.0 mg/ml stock) with and without 1.0 mM NH2PheHgAc (added from 10.0 mM stock) for 90 min at 37°C. Molecules which are activated by NH2PheHgAc are captured almost instantly by TIMP-1. TIMP-1 may be prepared from cultures of fibroblasts or similar cell lines that express fairly high levels of TIMP-1 activity (Bodden et al., 1994). Concentrations of this compound in the range of 0.1 to 1.0 mg/liter may be recovered from the culture medium. The purification scheme is somewhat cumbersome but greatly facilitated by use of antibody-based affinity chromatography techniques.

2. Mix with 5× electrophoretic sample buffer containing 0.5% SDS. Resolve by SDSPAGE using a 10% gel on ice at 100 V (UNIT 6.1). Note that the SDS concentration of the sample buffer is reduced to 0.1% (final concentration) in order to avoid dissociation of these entirely noncovalent complexes. This change is crucial to the success of the technique.

3. Transfer to nitrocellulose and stain adjacent lanes with antibodies to TIMP-1 and to MMP using standard immunoblotting techniques (UNIT 6.2). Typical results are shown in Figure 10.8.6.

α-MMP-1

A

α-TIMP-1

Complex

B

α-MMP-10

Complex

C

proMMP-1

proMMP-10

MMP-1

MMP-10

α-TIMP-1

D

TIMP-1

TIMP-1

APMA MMP-1 TIMP-1

– + –

+ + –

+ + +

– – +

– + –

+ + –

+ + +

– – +

APMA – MMP-10 + TIMP-1 –

+ + –

+ + +

– – +

Figure 10.8.6 TIMP capture. (A, B) are identical panels stained with antibodies to either human MMP-1 or TIMP-1. Capture of activated human MMP-1 gives rise to a new band in the 70-kDa range containing both MMP-1 and TIMP-1 (arrow). (C) proMMP-10 and activated MMP-10 stained with antibody to human MMP-10. (D) Addition of TIMP-1 to activated MMP-10 results in capture of the enzyme now migrating in a complex with TIMP-1 in the Mr 70-kDa range. Modified from Windsor et al. (1993).



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BASIC PROTOCOL 5

FLUORESCENT LABELING OF CRYPTIC CYS-RESIDUE IN MMPs Most MMP (and ADAM) precursors contain a cryptic thiol group derived from a single, unpaired cysteine residue in the propeptide. This group is coordinately bonded directly to the active site Zn (“cysteine switch”) and in this manner plays a significant role in maintaining the catalytic latency of the proteinase precursors. The protocol below permits unmasking and detection of this cryptic thiol group (Fig. 10.8.7). The “switch” opens upon addition of SDS, which allows reaction of the liberated thiol group with a fluorescent maleimide compound (Yamamoto et al., 1977; Lyons et al., 1991). Materials MMP-containing samples 20 µM fluorescent maleimide N-(7-(di-methylamino-4-methyl-3-coumarinyl) maleimide (DACM) in Tris⋅Cl standard buffer (see recipe for buffer; prepare from 1 mM DACM stock in DMSO or ethanol) 2-mercaptoethanol stock in electrophoretic sample buffer (see recipe for buffer): concentration appropriate to obtain 5% final concentration in reaction mixture Fluorescent lamp Photographic equipment Additional reagents and equipment for SDS-PAGE (UNIT 6.1) 1. Expose companion samples of 50 to 200 µg/ml MMP for 1 hr at room temperature to 20 µM DACM (final concentration) either in the presence or absence of 1% (w/v) SDS.

S +S D

_

S +S D

_

+S D

_

S

2. Stop reaction by adding 2-mercaptoethanol (as stock solution of appropriate concentration in electrophoretic sample buffer) to a final concentration of 5% (v/v).

52K 54K

MMP-1



MMP-3

MMP-10

Figure 10.8.7 Fluorescent labeling of propeptide cryptic thiol residue by fluorescent maleimide. The cysteine switch is “closed” in the nascent proenzyme and therefore not reactive with a fluorescent maleimide compound (DACM). Exposure to SDS “opens” the switch and renders the cryptic thiol group reactive with the maleimide resulting in covalent modification of the proenzyme and generation of a readily detectable fluorescent band. Left panel: MMP-1. Right panel, MMP-3 and MMP-10. Lower edge of each panel shows Coomassie blue staining of the same bands. Modified from Windsor et al. (1993). Current Protocols in Cell Biology

3. Resolve proteins by SDS-PAGE (UNIT 6.1). 4. Photograph under long-wavelength UV illumination. Typical results are shown in Figure 10.8.7.


Coomassie blue stain 0.5% (w/v) Coomassie blue R-250 30% (v/v) methanol 10% (v/v) acetic acid Store up to 6 months at room temperature Electrophoretic running buffer, pH 8.3 0.025 M Tris base 0.192 M glycine 0.1% (w/v) SDS Store up to 1 year at room temperature Electrophoretic sample buffer, 5× 0.2 M Tris⋅Cl, pH 6.8 (APPENDIX 2A) 5% (w/v) SDS 20% (w/v) glycerol 0.1% (w/v) bromphenol blue Store up to 1 year at room temperature This is the sample buffer used in Basic Protocol 2.

Gelatin solution, 8.7 mg/ml Add gelatin (bovine skin, Sigma-Aldrich type B6-6269) to 1 M Tris⋅Cl, pH 8.8 at 8.7 mg/ml. Dissolve by heating to 57°C, then filter through Whatman no. 1 filter paper. Gel destaining solution 30% (v/v) methanol 10% (v/v) acetic acid 60% (v/v) H2O Store up to 1 year at room temperature Gel washing buffers 1 to 4 Buffer 1: 2.5% (v/v) Triton X-100 3 mM NaN3 Buffer 2: 2.5% (v/v) Triton X-100 50 mM Tris⋅Cl, pH 7.5 (APPENDIX 2A) 3 mM NaN3 Buffer 3: 2.5% (v/v) Triton X-100 50 mM Tris⋅Cl, pH 7.5 (APPENDIX 2A) 3 mM NaN3 5 mM CaCl2 1 µM ZnCl2 continued



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Buffer 4: 50 mM Tris⋅Cl, pH 7.5 (APPENDIX 2A) 3 mM NaN3 5 mM CaCl2 1 µM ZnCl2 Buffers may be stored up to 1 year at room temperature.

Incubation solution 50 mM Tris⋅Cl, pH 7.4 (APPENDIX 2A) 0.2 M NaCl 5 mM CaCl2 0.02% (w/v) Brij-35 Store up to 1 year at 4°C Neutralizing buffer (0.2 M NaPi) Prepare the following stock solutions: Solution A: 2.78 g NaH2PO4 in 100 ml H2O Solution B: 5.365 g Na2HPO4⋅7H2O in 100 ml H2O Prepare working solutions as follows: 15.2 ml Solution A 64.8 ml Solution B 16.6 ml 5 M NaCl Add 80 ml 0.1 N NaOH Store up to 1 year at 4°C Tris⋅Cl standard buffer, pH 7.4 50 mM Tris⋅Cl, pH 7.4 (APPENDIX 2A) 0.2 M NaCl 5 mM CaCl2 Store up to 1 year at 4°C COMMENTARY Background Information


Dissolution of collagen type I Substrate. Although collapsing the gel by air drying is advantageous for most purposes, and the resulting collagen film is more similar to the density of collagen in interstitial connective tissues (Fig. 10.8.1), it is possible to seed the cells on top of (or inside) fully hydrated gels and to monitor the process as the cells dissolve their way through the collagen gel. Electron microscopy confirms that hydrated gels are very loose, with the individual fibrils spaced far apart. The collagen content is quite low compared to the liquid phase and accounts for only 0.03% of the mass and for a similarly small volume fraction of the gel. While use of reconstituted type I collagen fibrils as a substrate offers particular advantages because of its resistance to general proteolysis, it is possible to replace this substrate with other extracellular matrix components.

Type II collagen does not form fibrils as readily as does type I but might prove useful after additional refinement of the system. Type III collagen appears to gel adequately for this purpose and may also be used as a substrate. Films and gels of type IV collagen may also be used, as may Matrigel (predominantly composed of laminin), fibrin, and fibronectin. An important variation using fluorescently labeled fibronectin was devised by Chen and coworkers (Chen et al., 1984; Chen and Chen, 1987). Serum. Serum contains a number of factors expected to either promote or inhibit the proteolytic dissolution of the extracellular matrix including collagen fibrils. The high concentration of α2M (2 to 3 mg/ml or 3 to 4× 10−6 M), which effectively blocks most MMPs in test tube experiments, however, does not inhibit cell-mediated dissolution of the collagen fibril film. Serum also contains plasminogen at a concentration of ∼200 µg/ml (2 × 10−6 M). Addition of even low concentrations of plasmi-



nogen (4 µg/ml; 4 × 10−8 M) to serum-free cultures greatly accelerates the rate of dissolution of the collagen fibril film by human foreskin keratinocytes (or other cells) which express urokinase-type plasminogen activator (uPA) or tissue-type plasminogen activator (t-PA). The mechanism is not quite well understood but may involve a role for plasminogen in the extracellular activation of certain proMMP precursors as an essential step in the dissolution of the substrate. Cytokines, transcriptional activation. Addition of cytokines, growth factors, and agents such as TPA, which upregulate or induce expression of MMPs, generally accelerates dissolution of the fibril coating dramatically, but since these reagents upregulate a wide range of MMPs, it is not yet possible to determine whether a single MMP or group of MMPs is responsible for this effect. Inhibition. That dissolution of the collagen fibril coating is mediated by metalloproteinasedependent mechanisms is readily made evident by synthetic inhibitors. Inclusion of the Znchelating agent 1,10-phenanthroline completely blocks dissolution, as do synthetic MMP inhibitors such as BB94, BB2516 (British Biotech), and Galardin. A number of synthetic inhibitors currently exist; some of these may be obtained by directly contacting the pharmaceutical companies in question (British Biotech, Roche Diagnostics, Celltech). Serine proteinase inhibitors such as α1-antitrypsin (α1AT) and soybean trypsin inhibitor, as well as cysteine proteinase inhibitors such as E-64, have no effect on the rate of dissolution. These findings suggest that the process(es) that result in dissolution of the collagen fibrils are absolutely dependent on MMP activity. Zymography Gelatin zymography is a fairly straightforward yet very highly sensitive technique as long as heating and reduction are avoided during sample preparation. The method yields discrete, well-resolved, and distinct unstained bands on a blue background, which are clearly visible and easy to photograph and document with transillumination (Fig. 10.8.3). The activity may be quantified by comparison with standard curves of specific purified MMPs (Kleiner and Stetler-Stevenson, 1994), but the rate of lysis varies considerably from MMP to MMP, and the technique is primarily intended to provide qualitative information. A variation described by Lyons et al. (1991) permits moni-

toring of real-time progress of the reaction under UV-light by use of gelatin labeled by a fluorophore. Although gelatin zymography is highly sensitive, capable of detecting low picogram quantities of MMPs, the assay does not reflect the activity of these proteases present in the sample analyzed. This is because the addition of SDS to the sample prior to electrophoretic separation results in dissociation of many enzyme inhibitor complexes. Therefore, zymography represents an excellent technique for identification of MMP species present in a given sample, but overinterpretation of the results—e.g., assessment of specific activity—is a common pitfall. Casein zymograms develop more slowly, almost invariably require overnight incubation, and tend to produce less sharp bands. (Latent) proenzyme forms also show up because of the “switch”-opening effect of SDS, but these forms do not necessarily acquire full catalytic activity. “Activation” by organomercurials (0.5 to 1.0 aminophenylmercuric acetate in 50 mM Tris⋅HCl buffer, pH 7.5, for 20 min to 20 hr) before sample preparation often results in higher levels of proteolytic activity but also shifts the Mr of the individual bands because of autolytic cleavage and removal of the propeptide. α2M capture Capture techniques permit direct assessment of the ability of various forms of MMPs to bind to natural inhibitors in a manner that resists dissolution by exposure to low concentrations of SDS. Zymographic techniques are not capable of discriminating between latent and catalytically active forms of the enzymes. That, however, can readily be achieved by α2M capture. TIMP capture (see Alternate Protocol) on the other hand does not depend on proteolytic activity and merely requires a correctly folded, but not necessarily catalytically competent, active site (Windsor et al., 1994). TIMP capture The method is particularly useful for analysis of the binding capacity of mutants in TIMPs and in MMPs (Windsor et al., 1994; Caterina et al., 1997). It is important to recognize that TIMP binding is not necessarily synonymous with catalytic competence. Mutants of MMP which are correctly folded but devoid of catalytic activity, such as the E200Q mutant of MMP-1 in which the active site glutamate is replaced with glutamine (and therefore catalytically inactive) still forms complexes with



Supplement 13

TIMP-1 fully as well as the native enzyme (Windsor et al., 1994). TIMP-1 captures both truncated and full-length forms, as long as the “switch” is open (by APMA). Fluorescent labeling of cryptic Cys residue The nascent closure of the “cysteine switch” by bonding of the single unpaired propeptide Cys residue to the active site Zn2+ converts a catalytic Zn-binding site to a structural Znbinding site. In order to monitor the (re)opening of the switch as a preamble to zymogen activation, the authors of this unit reasoned that covalent linkage of a fluorophore to the free thiol group might render this process easily visible and potentially quantifiable. That is indeed the case. The method shows, for instance, that the phenomenon of “switch opening” can be readily visualized in the absence of propeptide cleavage by exposure either to SDS or to EDTA.

Critical Parameters and Troubleshooting


Dissolution of collagen type I Even for the experienced operator, collagen is not an easy protein to work with. Its preparation and use require meticulous and stringent adherence to the rules and conditions that “work,” often with very little leeway for shortcuts and modifications. The most important checkpoint comes after the initial gelling. Unless there is clear and unequivocal evidence of gelling after 2 hr, efforts should be made to identify and correct the problem. Since there is no simple way to measure collagen concentration, the authors have utilized initial dry powder weight from materials stored in refrigerated dessicator jars as a guide. The concentrations mentioned in this unit refer to powder weight under these conditions. It is absolutely necessary that the solution from which the collagen is lyophilized be completely salt-free following extensive dialysis against dilute acetic acid. This problem may be avoided by purchase of commercial preparations of rat or bovine type I collagen, but it is necessary to test the gelling properties of the particular brand in question at the desired concentration and under the desired conditions. After 2 hr of gelation, the gel should be reasonably firm, i.e., it should not disintegrate upon gentle flicking of the plate. If the gel disintegrates during this test, the problem must first be solved before proceeding. The homogeneity of the gel is also very important. This is best checked following the

first air drying and washing step by staining a newly prepared film with Coomassie blue. This will instantly reveal whether the gel is uniform and homogenous and if it contains particulate matter (which can be removed by centrifugation) or air bubbles. Both must be avoided, and the technique must be improved until each gel is completely uniform and homogenous after staining. It is also important to ascertain after the first washing of the first-time dried gel that salt-crystal deposits (formed during the initial drying phase) have been completely removed by washing. This is most easily checked using the phase-contrast microscope. The gel should look granular but uniform; any trace of crystal patterns is a certain indication of inadequate washing. Zymography Gelatin zymography presents few, if any, technical challenges, hence the popularity and universal application of this technique. Because of the longer incubation time required and the lesser sensitivity, casein zymograms often give less distinct and more diffuse bands. Although it has not been widely explored, it is highly likely that a large number of other substrates could be substituted for either gelatin or casein. Reverse zymography, on the other hand, is technically challenging and requires great care and skill as well as considerable practice and experience. The latter method is, however, a uniquely powerful technique to identify discrete MMP-inhibitory bands. Inhibitor capture While commercial preparations of α2M are available the method is critically dependent on the native configuration of the inhibitor. Consequently, the authors rely only on freshly isolated inhibitor. Occasionally methods which are employed to activate MMPs, such as exposure to organomercurials, adversely affect the inhibitor and render the capture reaction partial rather than complete. In some cases trypsin activation (stopped by soy bean trypsin inhibitor) is preferable, but many mutants are highly sensitive to trypsin and rapidly degrade during activation attempts. Fluorescent labeling of cryptic Cys residue The method is fairly straightforward, although care must be taken to exclude any chemicals from the solutions that interfere with the Cys-maleimide reaction (e.g., heavy metals, N-ethylmaleimide, or iodoacetate). Photo-



graphic documentation can be tricky, but usually works well when using reflected UV light.

Anticipated Results Dissolution of collagen type I Use of 1- to 2-µm films results in complete dissolution within 1 to 4 days. Initially the cells penetrate the collagen fibril coating in discrete spots, which eventually coalesce to form contiguous zones devoid of collagen fibrils (Fig. 10.8.2). Dissolution of the fibril coating is strictly limited to the area immediately beneath the cell layer and does not extend beyond the boundaries of the cell colony. A similar pattern is observed in the presence of serum or purified plasminogen. Zymography When performed correctly, the reverse zymography–stained gel shows discrete, well-resolved bands of TIMPs on a virtually unstained background, indicating that all of the gelatin has been degraded except in and around the TIMP bands (Fig. 10.8.4). While this method yields important information when used in qualitative or semiquantitative fashion, the read-out may be quantified as described by Kleiner and colleagues (Oliver et al., 1997). As with direct zymography, reverse zymography is a highly sensitive technique that can detect as little as 50 to 100 pg of TIMPs in a given sample (Oliver et al., 1997). However, as with direct zymography, careful interpretation of results is essential. Again, use of SDScontaining sample buffers and electrophoretic separation of the sample results in dissociation of some protease-inhibitor complexes. Thus, the levels of TIMPs present may not accurately reflect the actual free TIMP levels present in the samples analyzed. Alternatively, as described for the TIMP capture assays, not all TIMP-MMP complexes may be dissociated by SDS, and TIMP-binding to an MMP active site does not necessarily reflect proteolytic competency of the enzyme. α2M capture Incubation of native α2M with activated proteinases that cleave the bait region result in full or partial capture of the attacking proteinase. Complete capture requires a significant molar excess of inhibitor (with the amount varying from proteinase to proteinase) which may be determined by titration in preliminary experiments. Because of the size difference, captured and uncaptured bands are readily re-

solved and identified on Western blots by staining with anti-MMP antibodies. Latent or catalytically inactive forms are not captured and remain at their usual migration position in the gel. TIMP capture Remarkably, most TIMP-MMP complexes survive dilute SDS solutions at room temperature and permit electrophoretic separation of free and complexed forms. The Mr difference (20 to 30 kDa) is sufficient to fully resolve the bands. As with α2M capture, latent forms of MMPs (“switch closed”) are not captured, and this method is therefore valuable in distinguishing “switch-open” and “switch-closed” forms before proteolytic excision of the propeptide during activation. The active site, however, does not have to possess catalytic activity, and inactive mutants (if correctly folded) readily form complexes with TIMPs. Fluorescent labeling of cryptic Cys residue Removal of Zn2+ with EDTA, as expected, also unmasks the cryptic thiol group. Fully converted (“activated”) forms of the enzyme which have lost the entire propeptide no longer react. Note, however, that the free thiol group is only a few residues upstream of the ultimate proteolytic processing site. Partially processed forms of the proenzymes therefore may still react with DACM.

Time Considerations Analysis of the degradation of collagen gels takes ∼2 days to prepare the gels and 1 to 4 days for the assay itself. It requires ∼2 weeks to prepare rat tail tendon collagen type I and 3 to 4 to label the collagen with fluorophore. Direct zymography takes 2 days to complete, while reverse zymography takes 2 days. α2M and TIMP capture take 1 to 2 days depending on the duration of antibody incubation in immunoblotting. Fluorescent labeling of the cryptic Cys residue can be completed in a single day.

Literature Cited Aimes, R.T. and Quigley, J.P. 1995. Matrix metalloproteinase-2 is an interstitial collagenase. Inhibitor-free enzyme catalyzes the cleavage of collagen fibrils and soluble native type I collagen generating the specific 3/4-length and 1/4-length fragments. J. Biol. Chem. 270:5872-5876. Birkedal-Hansen, H. 1987. Catabolism and turnover of collagens: Collagenases. Methods Enzymol. 144:140-171.



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Birkedal-Hansen, H. and Danø, K. 1981. A sensitive collagenase assay using [3H]collagen labeled by reaction with pyridoxal phosphate and [3H]borohydride. Anal. Biochem. 115:18-26. Birkedal-Hansen, H. and Taylor, R.E. 1982. Detergent-activation of latent collagenase and resolution of its component molecules. Biochem. Biophys. Res. Commun. 107:1173-1178.

Heussen, C. and Dowdle, E.B. 1980. Electrophoretic analysis of plasminogen activators in polyacrylamide gels containing sodium dodecyl sulfate and copolymerized substrates. Anal. Biochem. 102:196-202.

Birkedal-Hansen, H., Cobb, C.M., Taylor, R.E., and Fullmer, H.M. 1976. Synthesis and release of procollagenase by cultured fibroblasts. J. Biol. Chem. 251:3162-3168.

Kleiner, D.E. and Stetler-Stevenson, W.G. 1994. Quantitative zymography: Detection of picogram quantities of gelatinases. Anal. Biochem. 218:325-329.

Birkedal-Hansen, H., Birkedal-Hansen, B., Windsor, L.J., Lin, H.Y., Taylor, R.E., and Moore, W.G.I. 1989. Use of inhibitory (anti-catalytic) antibodies to study extracellular proteolysis. Immunol. Invest. 18:211-224.

Knäuper, V., Lopez-Otin, C., Smith, B., Knight, G., and Murphy, G. 1996. Biochemical characterization of human collagenase-3. J. Biol. Chem. 271:1544-1550.

Birkedal-Hansen, H., Moore, W.G. I., Bodden, M.K., Windsor, L.J., Birkedal-Hansen, B., DeCarlo, A., and Engler, J.A. 1993. Matrix metalloproteinases: A review. Crit. Rev. Oral Biol. Med. 4:197-250. Bodden, M.K., Harber, G.J., Birkedal-Hansen, B., Windsor, L.J., Caterina, N.C.M., Engler, J.A., and Birkedal-Hansen, H. 1994. Functional domains of human TIMP-1 (tissue inhibitor of metalloproteinases). J. Biol. Chem. 269:1894318952. Caterina, N.C.M., Windsor, L.J., Yermovsky, A.E., Bodden, M.K., Taylor, K.B., Birkedal-Hansen, H., and Engler, J.A. 1997. Replacement of conserved cysteines in human tissue inhibitor of metalloproteinases-1. J. Biol. Chem. 272:3214132149. Caterina, J.J., Yamada, S., Caterina, N.C.M., Longenecker, G., Holmback, K., Shi, J., Yermovsky, A.E., Engler, J.A., and Birkedal-Hansen, H. 2000. Inactivating mutation of the mouse tissue inhibitor of metalloproteinnases-2 (TIMP2) gene alters proMMP-2 activation. J. Biol. Chem. 275:26416-26422. Chen, J.M. and Chen, W.T. 1987. Fibronectin-degrading proteases from the membranes of transformed cells. Cell 48:193-203. Chen, W.T., Olden, K., Bernard, B.A., and Chu, F.-F. 1984. Expression of transformation-associated protease(s) that degrade fibronectin at cell contact sites. J. Cell Biol. 98:1546-1555. DeClerck, Y.A., Yean, T.D., Lu, H.S., Ting, J., and Langley, K.E. 1991. Inhibition of autoproteolytic activation of interstitial procollagenase by recombinant metalloproteinase inhibitor MI/TIMP-2. J. Biol. Chem. 266:3893-3899. Deutsch, D.G. and Mertz, E.T. 1970. Plasminogen: Purification from human plasma by affinity chromatography. Science 170:1095-1096.


collagen fibrils by gingival fibroblasts isolated from patients of various periodontitis categories. J. Periodontal Res. 33:280-291.

Laemmli, U.K. 1970. Cleavage of structural proteins during the assembly of the head of bacteriophage T4. Nature 227:680-685. Lin, H.Y., Wells, B.R., Taylor, R.E., and BirkedalHansen, H. 1987. Degradation of type I collagen by rat mucosal keratinocytes. J. Biol. Chem. 262:6823-6831. Lyons, J.G., Birkedal-Hansen, B., Moore, W.G.I., O’Grady, R.L., and Birkedal-Hansen, H. 1991. Characteristics of a 95-kDa matrix metalloproteinase produced by mammary carcinoma cells. Biochemistry 30:1450-1456. Ohuchi, E., Imai, K., Fuji, Y., Sato, H., Seiki, M., and Okada, Y. 1997. Membrane type 1 matrix metalloproteinase digests interstitial collagens and other extracellular matrix macromolecules. J. Biol. Chem. 272:2446-2451. Oliver, G.W., Leferson, J.D., Stetler-Stevenson, W.G. and Kleiner, D.E. 1997. Quantitative reverse zymography: Analysis of picogram amounts of metallopooteinase inhibitors using gelatinase A and B reverse zymograms. Anal. Biochem. 244:161-166. Sottrup-Jensen, L. and Birkedal-Hansen, H. 1989. Human fibroblast collagenase-α-macroglobulin interactions. J. Biol. Chem. 264:393-401. Sottrup-Jensen, L., Stepanik, T.M., Wierzbicki, D.M., Jones, C.M., Lonblad, P.B., Kristensen, T., Mortensen, S.B., Petersen, T.E., and Magnusson, S. 1983. The primary structure of α-macroglobulin and localization of a factor XIIIa cross-linking site. Ann. N.Y. Acad. Sci. 421:41-60. Springman, E.B., Angleton, E.L., Birkedal-Hansen, H., and Van Wart, H.E. 1990. Multiple modes of activation of latent human fibroblast collagenase: Evidence for the role of a Cys73 activesite zinc complex in latency and a “cysteine switch” mechanism for activation. Proc. Natl. Acad. Sci. U.S.A. 87:364-368.

Ghersi, G., Goldstein, L.A., Wang, J.-Y., Yeh, Y., Hakkinen, L., Larjava, H. and Chen, W.-T. 2001. Regulation of fibroblast migration on collagenous matrix by novel cell surface protease complex. J. Biol. Chem. In press.

Van Wart, H.E. and Birkedal-Hansen, H. 1990. The cysteine switch: A principle of regulation of metalloproteinase activity with potential applicability to the entire matrix metalloproteinase gene family. Proc. Natl. Acad. Sci. U.S.A. 87:55785582.

Havemose-Poulsen, A.P.H., Stolze, K., and Birkedal-Hansen, H. 1998. Dissolution of type I

Windsor, L.J., Grenett, H., Birkedal-Hansen, B., Bodden, M.K., Engler, J.A., and Birkedal-Han-



sen, H. 1993. Cell-type-specific regulation of SL-1 and SL-2 genes. Induction of SL-2, but not SL-1, in human keratinocytes in response to cytokines and phorbolesters. J. Biol. Chem. 268:17341-17347. Windsor, L.J., Bodden, M.K., Birkedal-Hansen, B., Engler, J.A., and Birkedal-Hansen, H. 1994. Mutational analysis of residues in and around the active site of human fibroblast-type collagenase. J. Biol. Chem. 269:26201-26207. Yamamoto, K., Sekine, T., and Kanaoka, Y. 1977. Fluorescent thiol reagents. XII. Fluorescent tracer method for protein SH groups using N-(7dimethylamino-4-methyl coumarinyl) maleimide. Anal. Biochem. 79:83-94.

Contributed by Henning Birkedal-Hansen and Susan Yamada National Institute of Dental Research National Institutes of Health Bethesda, Maryland

Jack Windsor University of Indiana School of Dentistry Indianapolis, Indiana Anne Havemose Poulsen University of Copenhagen School of Dentistry Copenhagen, Denmark Guy Lyons Kanematsu Laboratories Royal Prince Alfred Hospital Sydney, Australia William Stetler-Stevenson and Bente Birkedal-Hansen Center for Cancer Research National Cancer Institute National Institutes of Health Bethesda, Maryland



Supplement 13

Preparation of Extracellular Matrices Produced by Cultured Fibroblasts

UNIT 10.9

Culturing fibroblasts on traditional two-dimensional substrates induces an artificial polarity between lower and upper surfaces of these normally nonpolar cells. Not surprisingly, fibroblast morphology and migration differ once the cells are suspended in threedimensional collagen gels (Friedl and Brocker, 2000). However, the molecular composition of collagen gels does not mimic the natural fibroblast microenvironment. Fibroblasts secrete and organize extracellular matrix (ECM), which provides structural support for their adhesion, migration, and tissue organization, besides regulating cellular functions such as growth and survival (Buck and Horwitz, 1987; Hay, 1991; Hynes, 1999; Geiger et al., 2001). Cell-to-matrix interactions are vital for vertebrate development. Disorders in these processes have been associated with fibrosis, developmental malformations, cancer, and other diseases. In this unit, a method is described for generating tissue culture surfaces coated with a fibroblast-derived three-dimensional ECM produced and deposited by mouse NIH-3T3 cells (see Basic Protocol). This matrix closely resembles in vivo mesenchymal matrices and is composed mainly of fibronectin fibrillar lattices. Utilizing in vivo–like three-dimensional matrices as substrates allows the acquisition of information that is physiologically relevant to cell-matrix interactions, structure, function, and signaling, and which differs from data obtained by culturing cells on conventional two-dimensional substrates in vitro (Cukierman et al., 2001). These protocols were initially derived from methods described in UNIT 10.4. These methods were modified to obtain fibroblast-derived three-dimensional matrices and characterize cellular responses to them. The basic approach is to allow the cultured fibroblasts to produce their own three-dimensional matrix. For this purpose, fibroblasts are plated and maintained in culture in a confluent state. After 5 to 9 days, matrices are denuded of cells, and cellular remnants are removed. Such extraction results in an intact fibroblast-derived three-dimensional matrix, free of cellular debris, that remains attached to the culture surface (see Fig. 10.9.1). The fibroblast-derived three-dimensional matrices are then washed with PBS and can be stored for periods of up to 2 to 3 weeks at 4°C. In order to evaluate the quality of the fibroblast-derived three-dimensional matrix, support protocols present procedures for assessing the induction of rapid cell attachment (see Support Protocol 1) and the acquisition of in vivo–like spindle-shaped morphology (see Support Protocol 2) by plating new fibroblasts into the three-dimensional matrix. Additional support protocols describe how to mechanically compress the fibroblast-derived threedimensional matrices in order to obtain two-dimensional substrate controls (see Support Protocol 3) and how to solubilize the fibroblast-derived three-dimensional matrices (see Support Protocol 4) to produce a matrix-derived protein mixture for additional two-dimensional coating controls and for subsequent biochemical analysis of the matrices (also see Commentary). NOTE: All solutions and equipment coming into contact with living cells must be sterile, and proper aseptic technique should be used accordingly. NOTE: All culture incubations are performed in a humidified 37°C,10% CO2 incubator unless otherwise specified.

Extracellular Matrix Contributed by Edna Cukierman Current Protocols in Cell Biology (2002) 10.9.1-10.9.15 Copyright © 2002 by John Wiley & Sons, Inc.


A

B

C

D

Figure 10.9.1 Fibroblast-derived three-dimensional matrices before and after extraction process. (A) Culture at day 5 prior to matrix extraction. (B) The resulting fibroblast-derived three-dimensional matrix. (C) Magnified inset from panel A. (D) Magnified inset from panel B. Bars represent 50 µm.

BASIC PROTOCOL

PREPARATION OF EXTRACELLULAR MATRICES PRODUCED BY CULTURED FIBROBLASTS In this protocol, cultured fibroblasts produce their own three-dimensional matrix. The fibroblasts are plated and maintained in culture in a confluent state. After 5 to 9 days, matrices are denuded of cells, and cellular remnants are removed.

Preparation of Extracellular Matrices Produced by Fibroblasts

Depending on the laboratory equipment available and the anticipated uses of the fibroblast-derived three-dimensional matrices, one must select a suitable surface on which the matrices will be produced (e.g., glass-bottom dishes, coverslips, or tissue culture dishes). (1) Disposable glass-bottom dishes (MatTek) can be utilized for real-time fluorescent experiments or for quality assessment assays (e.g., cell attachment) using an inverted fluorescent microscope (see Support Protocol 1). (2) Coverslips can be used for immunofluorescence experiments in which samples are to be fixed and mounted on microscope slides (see Support Protocol 2), or for mechanical compression of the fibroblast-derived three-dimensional matrices to be used as control two-dimensional surfaces (see Support Protocol 3). (3) Regular tissue culture dishes (e.g., 35-mm diameter) can be used for in vivo observations with an inverted microscope, or if the matrix is to be collected and solubilized for additional characterization and/or for biochemistry analysis (see Support Protocol 4).



Materials NIH-3T3 cells growing in tissue culture (see recipe) Trypsin/EDTA solution (see recipe) Confluent medium (see recipe) Absolute (anhydrous) ethanol Phosphate-buffered saline (PBS; APPENDIX 2A) 0.2% (w/v) gelatin solution (see recipe) Matrix medium (see recipe) Extraction buffer (see recipe), 37°C 10 U/ml DNase I (Roche) in PBS (APPENDIX 2A) containing 1 mM CaCl2 and 1 mM MgSO4 (optional) PBS (APPENDIX 2A) supplemented with 100 U/ml penicillin, 100 µg/ml streptomycin, and 0.25 µg/ml Fungizone (additives available from Life Technologies) 15-cm tissue culture dishes Inverted phase-contrast microscope Small tweezers (e.g., Dumont no. 4), sterilized Appropriate surface for producing matrix (one of the following; see discussion above): Glass-bottom no. 1.5 plates (MatTek) 22-mm circular high-quality coverslips (Carolina Biological Supply) with 35-mm tissue culture dishes as containers 6-well tissue culture plates or 35-mm dishes Additional reagents and equipment for tissue culture (UNIT 1.1) Prepare cell cultures 1. Aspirate and discard the culture medium from a semiconfluent (80% confluent) culture of NIH-3T3 cells growing on a 15-cm culture tissue culture dish. 2. Rinse the cell layer briefly with trypsin/EDTA solution. This rinse will remove traces of serum that contains trypsin inhibitors.

3. Add enough trypsin–EDTA solution to cover the cell layer, quickly aspirate excess liquid, and observe under an inverted microscope at room temperature until the cells have detached from the culture dish (1 to 3 min). 4. Collect the cells in 10 ml of confluent medium by adding the medium to the dish and swirling to suspend the trypsinized cells. 5. Add 2 ml of the suspended cells to a 15-cm tissue culture dish and culture for 2 to 3 days, until semiconfluent (80% confluent). As many as five 15-cm culture dishes may be used.

Prepare surfaces for matrix deposition 6. If coverslips are to be used as the surface for fibroblast-derived three-dimensionalmatrix deposition, presterilize by flaming the coverslips after dipping in absolute (anhydrous) ethanol, place in 35-mm dishes, and rinse with PBS. Use small sterilized tweezers (e.g., Dumont no. 4) to handle the coverslips. 7. Add 2 ml of 0.2% gelatin solution to the culture surfaces to be used for fibroblastderived three-dimensional matrix deposition and incubate 1 hr at 37°C. 8. Aspirate liquid and add 2 ml PBS. At this point, the surfaces are ready to be seeded with matrix-producing fibroblasts. Extracellular Matrix


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Allow matrix deposition This protocol was developed for NIH-3T3 cells; nevertheless, other fibroblast cell lines may be used. For example, the same protocol can be followed from this point on using human or other fibroblasts. 9. Trypsinize growing cells as in steps 1 to 3. 10. Collect cells from each dish in 10 ml of matrix medium (see step 4, but substitute matrix medium for confluent medium). Count cells (UNIT 1.1), and dilute with matrix medium to a final concentration of 1 × 105 cells per ml. 11. Aspirate PBS from gelatin-coated dishes (see step 8). 12. Seed 2 × 105 cells (2 ml) per 35-mm dish and culture for 24 hr. Use as many dishes as needed; each 15-cm semiconfluent dish should provide enough cells for about 100 35-mm dishes.

13. After 24 hr, carefully aspirate the medium and replace with fresh matrix medium. 14. Replace medium with freshly made matrix medium every 48 hr for a total of 5 to 9 days after step 12, until the matrix is ready to be denuded of cells (see Fig. 10.9.1A). Extract cells from fibroblast-derived three-dimensional matrices 15. Carefully aspirate the medium and rinse gently with 2 ml PBS by touching the pipet against the dish wall rather than at the bottom of the dish where the cells are located. 16. Gently add 1 ml of prewarmed (37°C) extraction buffer. If coverslips are being used, gently lift the coverslip with the fine-pointed (e.g., Dumont no. 4) tweezers (or a syringe needle) so that extraction buffer reaches underneath. This step will ensure that the matrix deposited on the coverslip will be separated successfully from the remainder of the matrix deposited on the bottom of the culture dish, thus facilitating subsequent handling of the coverslips without tearing the delicate matrix.

17. Observe the process of cell lysis using an inverted microscope. Incubate until no intact cells are seen (about 3 to 5 min; see Fig. 10.9.1B). 18. Dilute the cellular debris by adding 2 to 3 ml PBS. The above dilution process should be carried out gently in order to prevent turbulence that may cause the freshly denuded matrix layer to detach from the surface.

19. As cautiously as possible (using a pipet), aspirate the diluted cellular debris, but do not completely aspirate the liquid layer. To avoid removing the matrix layer, do not attempt to aspirate the whole volume.

20. Gently repeat steps 18 and 19. 21. Optional: To minimize DNA debris, incubate the matrices with 2 ml of 10 U/ml DNase 30 min at 37°C. At the end of the incubation, aspirate the enzyme solution and wash twice with PBS. 22. Cover the matrix-coated plates (or coverslips) with at least 3 ml PBS supplemented with 100 U/ml penicillin, 100 µg/ml streptomycin, and 0.25 µg/ml Fungizone. Seal with Parafilm. Preparation of Extracellular Matrices Produced by Fibroblasts

These matrices can be stored for up to 2 or 3 weeks at 4 °C. However, for signal-transduction assays in serum-free medium, see Commentary.



23. Confirm the integrity of the matrices directly before use by examining for matrix integrity using an inverted phase-contrast microscope; the matrices should be attached to the culture surface and appear similar to the example in Fig. 10.9.1B. ASSESING THE QUALITY OF FIBROBLAST-DERIVED THREE-DIMENSIONAL MATRICES The quality of fibroblast-derived three-dimensional matrices can be tested by one of two assays: induction of rapid cell attachment (see Support Protocol 1) and rapid acquisition of a spindle-shape morphology (see Support Protocol 2). These assays are based on examination of fluorescently labeled cells plated on three-dimensional matrices; the prelabeling with fluorescent dye is required in order to enhance observation of cells within fibroblast-derived three-dimensional matrices. Cell Attachment Assay Human or mouse fibroblasts can be used to evaluate the cell adhesion–promoting activity of the fibroblast-derived three-dimensional matrices. It has been reported that these in vivo–like three-dimensional matrices are about 6-fold more effective than two-dimensional substrates in mediating cell adhesion as quantified by a 10-min cell attachment assay (Cukierman et al., 2001). Briefly, cell nuclei are prelabeled to avoid any background staining from DNA debris on the three-dimensional matrix. The live prelabeled cells are rinsed free of excess dye, trypsinized, and plated on the fibroblast-derived three dimensional matrix to be assessed, or onto control fibronectin-coated surfaces. After 10 min, nonattached cells are washed away, and attached cells are quantified by counting labeled nuclei.

SUPPORT PROTOCOL 1

Materials Semiconfluent fibroblasts (human or mouse) in 15-cm dish (see Basic Protocol, steps 1 to 5) Confluent medium (see recipe) Hoechst 33342 stock solution (see recipe) Phosphate-buffered saline (PBS; APPENDIX 2A), room temperature and 4°C Trypsin/EDTA solution (see recipe) Fixing solution (see recipe) 15-ml conical polypropylene centrifuge tubes Tissue culture centrifuge with rotor suitable for 15-ml conical tubes End-over-end rotator 3 glass-bottom no. 1.5 plates containing fibroblast-derived three-dimensional matrix (see Basic Protocol) 3 glass-bottom no. 1.5 plates with precoated two-dimensional fibronectin (see recipe; also see Support Protocols 3 and 4) Inverted fluorescence microscope with appropriate camera and filter set to visualize Hoechst 33342 (see APPENDIX 1E) Image analysis software capable of counting objects (optional; e.g., MetaMorph from Universal Imaging) Additional reagents and equipment for tissue culture (UNIT 1.1) Prepare cells 1. Aspirate and discard the culture medium from a semiconfluent (80% confluent) 15-cm culture dish containing fibroblasts (mouse or human). Extracellular Matrix


Supplement 16

2. Add 20 ml of confluent medium containing 40 µl of Hoechst 33342 stock solution to the cells. 3. Incubate 15 min at 37°C. 4. Rinse with PBS four times. 5. Add enough trypsin/EDTA solution to cover the cell layer, quickly aspirate excess liquid, and observe under an inverted microscope until the cells have detached from the culture dish (1 to 3 min). 6. Collect the cells in 10 ml of confluent medium by adding the confluent medium to the dish and swirling to suspend the trypsinized cells. Transfer into a 15-ml conical polypropylene centrifuge tube and count an aliquot of the cells (UNIT 1.1). 7. Centrifuge 5 min at 100 × g, room temperature. 8. Discard the supernatant and gently resuspend the cells with confluent medium to a final concentration of 3.5 × 105 cells/ml. 9. Rotate cells in suspension for 20 min at 37°C on an end-over-end rotator. Perform adhesion assay 10. Carefully place a 150-µl drop of cell suspension onto the glass bottom part of each of the dishes coated with three-dimensional matrices or two-dimensional fibronectin controls. 11. Incubate 10 min at 37°C. 12. Remove dishes from incubator, tilt dishes slightly to dislodge the medium droplet containing unattached cells from the glass portion onto the plastic portion of the dish, then aspirate the droplet. 13. Rinse dishes by slowly adding (to the plastic portion of each dish) 3 ml PBS precooled to 4°C. 14. Aspirate the PBS carefully and add 2 ml fixing solution. 15. Incubate 20 min at room temperature. 16. Aspirate and add 2 ml PBS at room temperature. Visualize and analyze cell attachment 17. Using an inverted fluorescence microscope, acquire five random images of the nuclei from each one of the six dishes utilizing a 10× or 20× objective and count the Hoechst-stained nuclei. Counting of the nuclei can be done automatically utilizing commercially available imageanalysis software capable of counting objects (e.g., MetaMorph from Universal Imaging Corporation). If the counting is done automatically, then images should be acquired with a 10× objective. However, if the nuclei are to be counted manually, then a 20× objective is recommended. The mean number of cells attached to the fibroblast-derived three-dimensional matrix should be up to 6-fold higher than the number attached to the fibronectin control. This result will confirm the quality of the fibroblast-derived three-dimensional matrix (Cukierman et al., 2001).




Determination of Cell Shape Human or mouse fibroblasts can be used to evaluate induction of spindle-shaped cell morphology promoted by good-quality in vivo–like three-dimensional matrix. A recent report has established that fibroblasts will acquire an in vivo–like spindle-shaped morphology in cell-derived three-dimensional matrices 5 hr after plating (Cukierman et al., 2001). The protocol consists of prelabeling live fibroblast membranes with a fluorescent dye and incubating the cells on fibroblast-derived three-dimensional matrices or controls for a period of 5 hr. After this period of time, the fibroblast-derived three-dimensional matrix promotes a spindle-shaped morphology resembling in vivo fibroblast morphology, thereby confirming the quality of the three-dimensional matrices.

SUPPORT PROTOCOL 2

Materials 2% (w/v) BSA, heat denatured (see recipe) Phosphate-buffered saline (PBS; APPENDIX 2A) Semiconfluent fibroblasts (human or mouse) in 15-cm dish (see Basic Protocol, steps 1 to 5) Trypsin/EDTA solution (see recipe) Confluent medium (see recipe) 4 µg/ml DiI working solution (prepared in confluent medium; see recipe) Fixing solution (see recipe) Gel Mount mounting medium (Biomeda) 3 coverslips coated with fibroblast-derived three-dimensional matrix (see Basic Protocol) 3 coverslips with precoated two-dimensional matrix (see recipe; also see Support Protocols 3 and 4) 35-mm tissue culture dishes or 6-well plates Small tweezers (e.g., Dumont no. 4, sterilized) 15-ml conical polypropylene centrifuge tubes End-over-end rotator Tissue culture centrifuge with rotor suitable for 15-ml conical tubes Glass microscope slides Fluorescent microscope equipped with digital camera Image analysis software capable of measuring elliptical Fourier parameters (e.g., MetaMorph from Universal Imaging) Additional reagents and equipment for tissue culture (UNIT 1.1) Block nonspecific cell binding with BSA 1. Cautiously place fibroblast-derived three-dimensional matrix and two-dimensional control–coated coverslips into 35-mm tissue culture dishes (or 6-well plates) with the matrix face up. Use small sterilized tweezers (e.g., Dumont no. 4) to handle the coverslips. 2. Block nonspecific cell binding by adding 2 ml of heat-denatured 2% BSA and incubating for 1 hr at 37°C. 3. Rinse all blocked coverslips with 2 ml PBS. At this point coverslips are ready to be seeded with the prelabeled cells.

Label cell membrane with DiI 4. Aspirate and discard the culture medium from a semiconfluent (80% confluent) 15-cm dish of fibroblasts. 5. Rinse the cell layer briefly with trypsin/EDTA solution. This rinse will remove traces of serum that contains trypsin inhibitors.



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6. Add enough trypsin–EDTA solution to cover the cell layer, quickly aspirate excess liquid, and observe under an inverted microscope at room temperature until the cells have detached from the culture dish (1 to 3 min). 7. Collect the cells in 10 ml of 4 µg/ml DiI working solution (in confluent medium) by adding the solution to the dish and swirling to resuspend the trypsinized cells. Transfer the suspended cells into a 15-ml conical polypropylene tube. 8. Incubate the cells with the dye in suspension by rotating gently for 30 min at 37°C. 9. Centrifuge 5 min at 100 × g, room temperature. 10. Aspirate and discard the supernatant, then gently resuspend the cells with confluent medium to a final volume of 10 ml. 11. Repeat steps 9 and 10 four additional times to remove any remaining free dye. 12. Count cells (UNIT 1.1) and dilute with confluent medium to a final concentration of 1 × 104 cells per ml. 13. Carefully aspirate PBS from the coverslips (see step 3). 14. Add 2 ml of the diluted cell suspension to each of the dishes containing the coverslips and incubate 5 hr at 37°C. For fast qualitative analysis, cells can be observed and photographed at the end of 5 hr with an inverted microscope (see APPENDIX 1E for wavelength information).

15. Aspirate medium and rinse with PBS. 16. Aspirate PBS and fix for 20 min at room temperature with 1 ml of fixing solution. 17. Aspirate fixing solution and rinse with PBS. 18. Rinse with water to eliminate residual salt. 19. Carefully lift coverslip and gently discard excess liquid by touching the edge of the coverslip onto a paper towel. 20. Mount coverslips (with cells face-down) on a droplet (∼20 µl) of Gel Mount placed onto a glass microscope slide. 21. Allow mounted samples to dry in the dark at room temperature for ∼1 hr. At this point samples are ready for morphometry analysis, or they can be stored overnight in the dark at 4°C.

Perform morphometry analysis 22. Acquire fluorescent digital images, slightly overexposing to visualize the contour of the cells (for wavelength information, see APPENDIX 1E). Use a magnification that will allow visualization of the entire cell in each image. Randomly capture images of at least 12 cells per sample and a minimum of 36 cells per substrate.

23. Perform the measurements for both the length (span of the longest cord) and the breadth (caliper width) of each cell using image-analysis software. 24. Calculate the inverse axial ratio by dividing length by breadth. Preparation of Extracellular Matrices Produced by Fibroblasts

The mean inverse axial ratio induced by a high-quality fibroblast-derived three-dimensional matrix should be about 3-fold greater than that induced by the two-dimensional fibronectin control (Cukierman et al., 2001).



The inverse axial ratio corresponds to the elliptical form factor (EFF) morphometric parameter found in the integrated morphometry analysis (IMA) function of MetaMorph software.

PREPARING TWO-DIMENSIONAL EXTRACELLULAR MATRIX CONTROLS Any given cell response induced by in vivo–like fibroblast-derived three-dimensional matrices could be due to the three-dimensionality of the matrix, its molecular composition, or a combination of both. The following two support protocols provide methods for obtaining suitable two-dimensional controls with the same molecular composition as the three-dimensional matrices. Mechanical Compression of the Fibroblast-Derived Three-Dimensional Matrix This protocol describes how to apply pressure to the fibroblast-derived three-dimensional matrix in order to collapse the matrix to a flat substrate. Mechanical compression of the three-dimensional matrix ensures that all natural components of the three-dimensional matrix are present, with only the element of three-dimensionality lacking. Briefly, the three-dimensional sample is compressed using a known weight applied to a given area. The surface that comes into contact with the matrix is covered with a Teflon film to prevent sticking and to avoid tearing the flattened matrix as the weight is retracted.

SUPPORT PROTOCOL 3

Materials Superglue Phosphate-buffered saline (PBS; APPENDIX 2A) Fibroblast-derived matrix on 22-mm coverslip Ring stand equipped with a horizontal ring Flat platform large enough to rest on the ring (see Fig. 10.9.2) Suitable spacer smaller in width than the diameter of the ring but longer in height than the ring’s depth (see Fig. 10.9.2) 12-mm round coverslips (Carolina Biological Supply) Small tweezers (e.g., Dumont no. 4, sterilized) Cork borer Teflon film: protective overlay composed of, 0.001-in. FEP film on 0.008-in. vinyl film, with adhesive back (used to cover laboratory bench-tops; Cole-Parmer) Biological hood equipped with UV light (optional) Lifting laboratory jack (Fisher) Weight (∼158 g) Fibroblast-derived three-dimensional matrix on 22-mm circular coverslips (see Basic Protocol) 35-mm tissue culture dishes Inverted phase-contrast microscope NOTE: Any other materials fulfilling the same purpose can be substituted for the hardware listed above. Construct the weight holder for matrix compression 1. Using Superglue, glue the flat platform to the spacer in such a way that the spacer will protrude slightly beyond the bottom of the ring when the platform is placed on the ring (Fig. 10.9.2). Extracellular Matrix


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a

b

c d e f g h

Figure 10.9.2 Diagram showing the components of the mechanical compression device. (a) weight; (b) flat platform; (c) spacer; (d) 12-mm coverslips; (e) Teflon film; (f) ring stand; (g) fibroblastderived three-dimensional matrix to be mechanically compressed; (h) lifting laboratory jack.

2. Glue four coverslips to the end of the spacer (one on top of the other) as an extension of the spacer, and allow enough time for the Superglue to completely dry. Use small sterilized tweezers (e.g., Dumont no. 4) to handle the coverslips. This will facilitate penetration of the coverslip portion into the matrix while avoiding contact between the matrix and the rest of the spacer, and it defines the area of compression.

3. Cut a circle (12-mm diameter) from the Teflon film with the cork borer. 4. Cover the last coverslip with the circle of Teflon film. 5. Sterilize materials by exposing them to a UV light in a biological hood for several hours with the Teflon film facing the light. If the compressed matrices are to be in contact with cells for only short periods of time (e.g., for the 10-min cell attachment assay; see Support Protocol 1), rinsing the Teflon film with ethanol and air-drying should be sufficient to prevent contamination.

6. Place the glued platform with spacer on the ring portion of the stand with the Teflon facing down. 7. Cover the jack’s flat upper surface with Parafilm and position the jack under the ring. Preparation of Extracellular Matrices Produced by Fibroblasts

8. Set the weight on the platform and level the ring so that the Teflon film is situated parallel to the jack’s surface (see Fig. 10.9.2).



Mechanically compress the fibroblast-derived three-dimensional matrix 9. Position the fibroblast-derived three-dimensional matrix-coated coverslip, with the matrix face-up, onto the jack directly underneath the Teflon film. 10. Slowly raise the laboratory jack until the matrix contacts the Teflon film and the platform rises above the ring. 11. Wait for 2 min. At this point the entire weight should be resting on the matrix, compressing it at a specific weight per unit area.

12. Slowly lower the jack until the platform rests once again on the ring and the compressed matrix is separated from the Teflon film. 13. Place the coverslip with the compressed matrix into a 35-mm dish. 14. Carefully add 2 ml PBS and examine by phase-contrast microscopy to confirm continued integrity of the compressed matrix. Solubilization of Fibroblast-Derived Three-Dimensional Matrix This protocol describes how to solubilize fibroblast-derived three-dimensional matrix to generate a protein mixture that can be used for subsequent coating of surfaces or biochemical analysis. Briefly, the matrices are treated with a guanidine solution to denature and solubilize the matrix components, thereby producing a liquid mixture that can be stored and used for coating surfaces.

SUPPORT PROTOCOL 4

Materials Fibroblast-derived three-dimensional matrices on 35-mm dishes (see Basic Protocol) Solubilization reagent: 5 M guanidine containing 10 mM dithiothreitol (store indefinitely at 4°C) Rubber policeman End-over-end rotator Solubilize matrix 1. Aspirate PBS from matrix-covered dishes. 2. Tip dishes ∼30° with respect to the benchtop and hold in that position for 1 min to allow the excess PBS to accumulate on one side of the dish. 3. Aspirate the excess PBS carefully to avoid detaching the matrix layer. 4. Place the dishes on ice and add 300 µl of solubilization reagent. 5. Incubate on ice for 5 min. 6. Scrape the dish with the rubber policeman toward one side of the dish and pipet the mixture into a 1.5-ml microcentrifuge tube. 7. Add an additional 200 µl of solubilization reagent. 8. Rotate at 4°C for 1 hr. 9. Microcentrifuge 15 min at maximum speed, 4°C. 10. Transfer the supernatant into a fresh microcentrifuge tube and store at 4°C. The average protein concentration is 1 to 3 mg/ml. Extracellular Matrix


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BSA, heat-denatured, 2% (w/v) Stock solution (undenatured): Dissolve 2 g bovine serum albumin (BSA) fraction V (Sigma) in 100 ml water and filter sterilize using a low-protein-binding 0.22-µm filter. Store indefinitely at 4°C Heat-denatured working solution: Just prior to use, heat the amount needed for 5 min at 65°C or until the solution starts to appear slightly translucent (not milky). Cool to room temperature before using for blocking procedures. Do not store the heat-denatured BSA. Confluent medium High-glucose Dulbecco’s modified Eagle medium supplemented with: 10% (v/v) fetal bovine serum (FBS; APPENDIX 2A) 100 U/ml penicillin 100 µg/ml streptomycin Store up to 1 month at 4°C Culture medium with calf serum High-glucose Dulbecco’s modified Eagle medium supplemented with: 10% (v/v) calf serum 100 U/ml penicillin 100 µg/ml streptomycin Store up to 1 month at 4°C DiI stock and working solutions Stock solution: Prepare 2.5 mg/ml DiI (1,1′-dioctadecyl-3,3,3′,3′-tetramethylindocarbocyanine perchlorate; Molecular Probes) in ethanol. Store up to 6 months at −20°C. 4 ìg/ml working solution: Dilute DiI stock solution to 4 µg/ml with confluent medium (see recipe) and sterilize by filtration using a 0.22-µm filter. Use promptly; do not store. Extraction buffer PBS (APPENDIX 2A) containing: 0.5% (v/v) Triton X-100 20 mM NH4OH Store up to 1 month at 4°C Fibronectin-coated surfaces, precoated, two-dimensional Prepare a 10 mg/ml solution of human plasma fibronectin (see UNIT 10.5, or purchase from Sigma) in phosphate buffered saline (PBS; APPENDIX 2A). Immediately add 1 ml of this solution per 35-mm tissue culture dish (or other surface to be coated) and incubate 1 hr at 37°C. Remove remaining fibronectin solution and rinse once with PBS. The above procedure can be used with any desired protein for coating dishes or coverslips. If solubilized matrix mixture (see Support Protocol 4) is to be used, the coating protein concentration is 30 ìg/ml.


Fixing solution In a 50-ml conical polypropylene conical tube, combine the following: 2 g sucrose 10 ml 16% (w/v) paraformaldehyde (EM grade; Electron Microscopy Sciences) PBS (APPENDIX 2A) to 40 ml final Prepare fresh before use



Gelatin solution, 0.2% (w/v) Prepare a 0.2% (w/v) gelatin solution in PBS (APPENDIX 2A). Autoclave, cool, and filter through a 0.2-µm filter. Store up to 1 year at 4°C. Hoechst 33342 stock solution Prepare a 2 mM solution of Hoechst 33342 (bisbenzimide H33342 fluorochrome, trihydrochloride; Calbiochem; mol. wt. 615.9 g) in water. Store up to 6 months at 4°C, protected from light. Matrix medium To confluent medium (see recipe) add L-ascorbic acid, sodium salt (Sigma) to a final concentration of 50 µg/ml, from a freshly prepared 50 mg/ml stock solution. Sterilize by filtration with a 0.2-µm filter. NIH 3T3 cells, stock cultures NIH-3T3 cells (ATCC #CRL-1658) must be routinely cultured in DMEM supplemented with 10% calf serum (see recipe for culture medium with calf serum, above) although medium using FBS is used in the protocol steps. Never allow cultured NIH-3T3 cells to become completely confluent while maintaining stock cultures. When cells reach 80% confluence (about once per week), subculture at 1:20 dilution. Trypsin/EDTA solution 2.5 g trypsin 0.2 g tetrasodium EDTA 8 g NaCl 0.4 g KCl 1 g glucose 0.35 g NaHCO3 0.01 g phenol red H2O to 1 liter Sterilize by filtration with a 0.2-µm filter Store up to 3 months at −20°C This solution is also commercially available from Life Technologies.

COMMENTARY Background Information Extracellular matrix (ECM) was historically regarded as a passive scaffold that stabilizes the physical structure of tissues. With time, it became evident that the ECM is much more than a simple physical scaffold. The ECM is a dynamic structure capable of inducing (and responding to) a large variety of physiological cell responses regulating the growth, migration, differentiation, survival, and tissue organization of cells (Buck and Horwitz, 1987; Hay, 1991; Hynes, 1999). Integrins are receptors for matrix molecules and can mediate these cell responses by inducing the formation of membrane-associated multimolecular complexes. These integrin-based structures (cell-matrix adhesions) mediate strong cell-substrate adhesion and transmit information in a bidirectional manner between ECM and the cytoplasm.

There are three main cell-to-matrix adhesions. The “focal adhesion” mediates firm linkage to relatively rigid substrates (Burridge and Chrzanowska-Wodnicka, 1996). Focal adhesions cooperate with “fibrillar adhesions” that generate fibrils from pliable fibronectin (Katz et al., 2000; Pankov et al., 2000). Fibroblasts require culture for several days at high cell density to generate three-dimensional matrices and evolve “three dimensional matrix adhesions.” The requirements for producing threedimensional matrix adhesions include three dimensionality of the ECM, integrin α5β1, fibronectin, other matrix component(s), and pliability of the matrix (Cukierman et al., 2001). The fibroblast-derived matrix provides an in vivo–like three-dimensional environment for cultured fibroblasts, thereby restoring their normally nonpolar surroundings. The fi-



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broblast-derived three-dimensional matrix can be used as a suitable in vitro system to investigate in vivo–like fibroblast-to-matrix interactions, such as three-dimensional matrix–adhesion signaling.

Critical Parameters


The phenotype of cultured 3T3 fibroblasts as monitored by cell morphology is extremely important for the successful preparation of three-dimensional matrix–coated dishes. The fibroblasts should be well spread and flat under sparse culture conditions. If elongated cells are commonly observed in the cell population, recloning of the cell line may be necessary to achieve greater phenotypic homogeneity. The NIH-3T3 line obtained from ATCC (catalog no. CRL-1658) has this morphology and produces excellent matrix. The NIH-3T3 cells must be maintained routinely as subconfluent cultures in medium containing calf serum to retain the correct phenotype. However, if the matrix deposition at confluence is performed in the presence of calf serum, the resultant matrices are thicker but less stable and more likely to detach from the surface than matrices obtained after culture in fetal bovine serum. Therefore NIH-3T3 cells should be changed to medium containing fetal bovine serum prior to matrix deposition. A preadaptation of the cells in fetal bovine serum–containing medium after replating for 2 to 3 days is recommended. The Basic Protocol could potentially be modified for other fibroblastic cell lines capable of secreting and assembling fibronectinbased matrices. In some cases, the resulting matrix may be too thick or dense to obtain efficient extraction. In such cases, more prolonged cell extraction may be needed, with extensive DNase treatment, until no cell debris is detected. The lack of contaminating cellular debris (in the case of NIH-3T3 cells) in the matrices has been confirmed by Western blotting and immunofluorescence staining for cellular proteins like actin. Precoating surfaces with gelatin promotes fibronectin binding and results in smooth layers of relatively homogenous matrices that will not detach from the surface. The thickness of NIH-3T3-derived three-dimensional matrices is measured using a confocal microscope without dehydration of the matrix (no mounting or fixing). The resultant thickness observed varies between 8 and 20 µm. Basic molecular characterization of the matrices revealed the presence (among other molecules) of fibronectin organized in a fibril-

lar mesh, collagen I and III, but not IV, and small traces of nonorganized laminin and perlecan. The integrity of these three-dimensional matrices must be confirmed prior to every use. This can be accomplished by using phase-contrast microscopy and discarding any matrices that are torn or detached (see Fig. 10.8.1B). Moreover, if matrices are to be used for short-term signal transduction assays under serum-depleted conditions, freshly made matrices must be utilized. Matrices stored at 4°C (up to 2 to 3 weeks) should be used only after such assessment of integrity. Freshly prepared or stored matrices can be used to test the induction of cell responses in the presence of serum (e.g., attachment, morphology, motility, or proliferation), biochemical analysis (e.g., western blotting), and immunofluorescence staining.

Anticipated Results The Basic Protocol is based on the ability of densely cultured fibroblasts to coat any available tissue culture surface by deposition of their natural matrix, which gradually forms a threedimensional matrix. This intact, naturally produced ECM is similar in its molecular organization to mesenchymal fibronectin-based extracellular matrices in vivo (Cukierman et al., 2001). The basic approach is to allow cells to deposit their own ECM followed by removal of cells, while avoiding procedures that may alter or denature the native ECM constituents and supramolecular organization. One NIH-3T3 semi-confluent (80%) cultured 15-cm dish can yield enough cells to coat 100 35-mm tissue culture dishes.

Time Considerations The adaptation step after switching NIH3T3 cell medium to fetal bovine serum for future matrix deposition requires 2 to 3 days. Matrix production will require between 5 and 9 days.

Literature Cited Buck, C.A. and Horwitz, A.F. 1987. Cell surface receptors for extracellular matrix molecules. Annu. Rev. Cell Biol. 3:179-205. Burridge, K. and Chrzanowska-Wodnicka, M. 1996. Focal adhesions, contractility, and signaling. Annu. Rev. Cell Dev. Biol. 12:463-518. Cukierman, E., Pankov, R., Stevens, D.R., and Yamada, K.M. 2001. Taking cell-matrix adhesions to the third dimension. Science 294:17081712. Friedl, P. and Brocker, E.B. 2000. The biology of cell locomotion within three-dimensional extracellular matrix. Cell Mol. Life Sci. 57:41-64.



Geiger, B., Bershadsky, A., Pankov, R., and Yamada, K.M. 2001. Transmembrane crosstalk between the extracellular matrix and the cytoskeleton. Nat. Rev. Mol. Cell Biol. 2:793-805. Hay, E.D. 1991. Cell Biology of Extracellular Matrix, 2nd ed. Plenum, New York. Hynes, R.O. 1999. Cell adhesion: Old and new questions. Trends Cell Biol. 9:M33-M77. Katz, B.Z., Zamir, E., Bershadsky, A., Kam, Z., Yamada, K.M., and Geiger, B. 2000. Physical state of the extracellular matrix regulates the structure and molecular composition of cell-matrix adhesions. Mol. Biol. Cell 11:1047-1060. Pankov, R., Cukierman, E., Katz, B.Z., Matsumoto, K., Lin, D.C., Lin, S., Hahn, C., and Yamada, K.M. 2000. Integrin dynamics and matrix assem-

bly: Tensin-dependent translocation of alpha(5)beta(1) integrins promotes early fibronectin fibrillogenesis. J. Cell Biol. 148:10751090.

Key References Cukierman et al., 2001. See above This is the paper upon which the procedures and materials in this unit are based. The Basic Protocol was modified from UNIT 10.4 in this manual, which was contributed by I. Vlodavsky in 1999.

Contributed by Edna Cukierman National Institutes of Health Bethesda, Maryland



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Purification and Analysis of Thrombospondin-1

UNIT 10.10

Thrombosponding-1 (TSP-1) is a trimeric matricellular protein that is expressed by many cells. It contains several different domains that allow it to participate in cell adhesion, cell migration, and cell signaling. Recently, TSP-1 has been shown to activate transforming growth factor-β (TGF-β) and to inhibit both angiogenesis and tumor growth. This unit describes two protocols: the purification of TSP-1 from platelet-rich plasma (see Basic Protocol 1) and the purification of TSP-1 proteolytic fragments (see Basic Protocol 2). ISOLATION OF THROMBOSPONDIN-1 FROM HUMAN PLATELETS TSP-1 is released from platelet α-granules in response to thrombin and can therefore be readily purified from the supernatant of thrombin-treated platelets. Human platelets can be obtained from the Red Cross or from hospital blood banks. Outdated pheresis units of platelet-rich plasma are a good source of TSP-1. Platelets are separated from plasma and other blood components by a series of centrifugation steps. The isolated platelets are washed repeatedly to remove plasma proteins and the washed platelets are then activated by exposure to thrombin. Next the TSP-1-containing supernatant is passed over a heparin-Sepharose column. Lower-affinity heparin-binding proteins are washed away and the TSP-1 is eluted under conditions of high salt. The TSP-1-containing fractions are pooled, precipitated, and loaded onto a 10% to 20% continuous sucrose gradient and subjected to ultracentrifugation. The gradient is divided into fractions and the protein concentrations are determined by measuring optical density. The level of purity is normally >95% as determined by SDS-PAGE (UNIT 6.1).

BASIC PROTOCOL 1

Materials Platelet-rich plasma Baenziger A buffer (see recipe) Baenziger B buffer (see recipe) 1 M CaCl2 (APPENDIX 2A) 1 N NaOH (optional) Thrombin Diisopropyl fluorophosphate (DFP) Heparin-Sepharose CL-6B (Amersham Pharmacia Biotech) 0.15, 0.25, 0.55, and 2.0 M heparin-Sepharose column buffers (see recipe) Anti-vitronectin immunoaffinity column: prepare in advance according to manufacturer’s instructions using an Affi-Gel Hz Immunoaffinity kit (Bio-Rad) and anti–human vitronectin antibody (e.g., GIBCO/BRL) Ammonium sulfate 10% and 20% (w/v) sucrose gradient solutions (see recipe) 15- and 50-ml centrifuge tubes (conical bottom preferred) Preparative centrifuge (Sorvall RC-B3 or equivalent) and rotor (H4000 or equivalent) 40-ml Oak Ridge centrifuge tubes High-speed centrifuge (Beckman J2-MC or equivalent) and rotor (JA-20 or equivalent) 1 × 12–cm chromatography column Fraction collector and appropriate tubes Spectrophotometer set at 280 nm Gradient maker 14-ml ultracentrifuge tubes Ultracentrifuge (Beckman LM-80 or equivalent) and rotor (SW 41Ti or equivalent) Contributed by Karen O Yee, Mark Duquette, Anna Ludlow, and Jack Lawler Current Protocols in Cell Biology (2003) 10.10.1-10.10.13 Copyright © 2003 by John Wiley & Sons, Inc.



NOTE: Platelets are temperature sensitive and activated by untreated glass surfaces; therefore, they should be handled at room temperature in plasticware, and centrifuges and buffers should be warmed to room temperature before use. Prepare platelets 1. Transfer platelet-rich plasma to 50-ml centrifuge tubes (conical bottom preferred) and centrifuge in a Sorvall RC-B3 preparative centrifuge 20 min at 1400 × g (2800 rpm in an H4000 rotor), 20°C. Pheresis units are preferable, but random donor units of platelet-rich plasma also work well.

2. Carefully pour off the supernatant. Gently resuspend the cell pellet in Baenziger A buffer at a ratio of 15 ml buffer per 2 ml packed cells. 3. Transfer the platelet suspension to 15-ml centrifuge tubes and centrifuge 8 min at 120 × g (800 rpm in an H4000 rotor), room temperature. Most of the platelets will remain in suspension following this centrifugation, while erythrocytes and leukocytes will pellet.

4. Leaving behind the red cell pellet, carefully transfer the platelet suspension to 50-ml centrifuge tubes (∼22 ml per tube). 5. Add Baenziger A buffer to a final volume of 50 ml. Mix by inverting the tube several times and centrifuge 20 min at 1400 × g (2800 rpm in an H4000 rotor), 20°C. Wash platelet pellet 6. Carefully pour off the supernatant. Resuspend each cell pellet in 15 ml Baenziger A buffer and then add buffer to a final volume of 50 ml. Invert the tube to mix and centrifuge 20 min at 1400 × g (2800 rpm in an H4000), room temperature. Repeat once. 7. Remove the supernatant and resuspend the pellet in 15 ml Baenziger B buffer. Add sufficient Baenziger B to achieve a ratio of 50 ml buffer per 2 to 3 ml packed cells. Mix the tube by inversion. 8. Add 100 µl of 1 M CaCl2 per 50 ml suspension. From this point on, 2 mM calcium must be present at all times to maintain the conformational integrity of the thrombospondin molecule.

9. Check the pH of the suspension using pH paper. Adjust to pH 7.6 by adding 1 N NaOH as necessary. Activate platelets 10. Optional: If the platelets are from outdated units, enhance their response to thrombin by incubating 5 min in a 37°C water bath. 11. Add 50 U thrombin per 50 ml platelet suspension and immediately mix by gentle inversion. Continue mixing 2 to 3 min at room temperature, then place on ice. Platelet aggregation should be evident upon examination of the suspension. The platelets will form large clumps and settle to the bottom of the tube, causing the supernatant to appear somewhat clear after 2 to 3 min. Outdated platelets respond more slowly than fresh ones. Outdated units should therefore be mixed for an additional 2 to 3 min.


12. Remove the cellular debris by centrifuging the tubes 5 min at 1400 × g (2800 rpm in an H4000 rotor), 4°C. Transfer supernatant to a 40-ml Oak Ridge centrifuge tube.



From this point on the TSP-1-containing supernatant must be kept on ice and all subsequent steps must be performed at 4°C.

13. Add sufficient DFP to achieve a final concentration of 1 mM (i.e., 0.181 µl/ml). CAUTION: DFP is a powerful serum protease inhibitor and is highly toxic. Great care should be taken in its use. DFP is volatile and should be used in a fume hood.

Isolate TSP-1 supernatant 14. Centrifuge 20 min in a Beckman J2-MC high-speed centrifuge at 34,957 × g (17,000 rpm in a JA-20 rotor), 4°C. 15. Transfer the supernatant to a clean 50-ml tube. Place the sample on ice and leave overnight at 4°C. This incubation step is necessary to allow formation of fibrin fibrils, which are then removed by centrifugation (step 17). If the supernatants are applied to the heparinSepharose column without performing this procedure, the fibrin fibrils will form on the top of the column and the flow rate will be decreased significantly.

Isolate TSP-1 16. Prepare and pour enough heparin-Sepharose CL-6B, according to the manufacturer’s instructions, to produce a 5-ml bed volume in a 1 × 12–cm chromatography column. Equilibrate the column with 50 ml of 0.15 M heparin-Sepharose column buffer. 17. Following the overnight incubation (step 15), centrifuge the supernatant 20 min at 1400 × g (2800 rpm in an H4000 rotor), 4°C. Transfer the supernatant to a new tube. 18. Load the supernatant onto the equilibrated heparin-Sepharose column at a flow rate of ∼3 ml/min. The TSP-1 will be immobilized on the column following this step. If necessary, the protocol may be paused at this point; however, the column should be washed extensively with 0.15 M heparin-Sepharose column buffer before pausing. TSP-1 is stable on the column for 3 to 4 days.

19. Connect the column to a fraction collector with appropriate tubes and elute the column with 40 ml of 0.15 M heparin-Sepharose column buffer at a flow rate of ∼3 ml/min, collecting twenty 2-ml fractions. Repeat with 0.25 M heparin-Sepharose column buffer. Little or no TSP-1 will be present in these first two elutions.

20. Elute TSP-1 by applying 40 ml of 0.55 M heparin-Sepharose column buffer and collect in 2-ml fractions. Determine which fractions contain protein by measuring their absorbance at 280 nm. Calculate the total amount of protein in milligrams using the following formula: total protein = OD280 × 1.08 × volume. After elution, >80% of total protein is TSP-1.

21. Strip the heparin-Sepharose column by applying 100 ml of 2.0 M heparin-Sepharose column buffer. Equilibrate and store the column in 0.15 M heparin-Sepharose column buffer at 4°C. The column can be used repeatedly if treated in this manner.

22. Pool the protein-containing fractions and apply to an anti-vitronectin immunoaffinity column. Although vitronectin is present in only trace amounts in the TSP-1-containing fraction (95% pure.


Baenziger A buffer 0.102 M NaCl 0.0039 M K2HPO4 0.0039 M Na2HPO4 0.022 M NaH2PO4 0.0055 M glucose Store up to 2 weeks at 4°C Baenziger B buffer 0.015 M Tris⋅Cl, pH 7.6 (APPENDIX 2A) 0.14 M NaCl 0.005 M glucose Store up to 2 weeks at 4°C Heparin-Sepharose column buffers, 0.15, 0.25, 0.55, and 2.0 M 0.015 M Tris⋅Cl, pH 7.6 (APPENDIX 2A) 0.002 M CaCl2 0.15, 0.25, 0.55, or 2.0 M NaCl Store up to 3 weeks at 4°C The molarity of the buffer refers to the concentration of the NaCl.

Sucrose gradient solutions, 10% and 20% (w/v) 0.015 M Tris⋅Cl, pH 7.6 (APPENDIX 2A) 0.14 M NaCl 0.002 M CaCl2 10% or 20% (w/v) sucrose Store up to 1 week at 4°C TBS (Tris-buffered saline) 0.015 M Tris⋅Cl, pH 7.6 (APPENDIX 2A) 0.015 M NaCl Store up to 2 weeks at 4°C COMMENTARY Background Information


The thrombospondins are a family of extracellular matrix proteins currently consisting of five members, thrombospondins 1 to 4 and cartilage oligomeric matrix protein (COMP). For comprehensive reviews, see Adams (2001) and Chen et al. (2000). These proteins are synthesized by many tissues with patterns of expression that are temporally and spatially

regulated. All thrombospondin family members are composed of a series of multidomain structures and have the ability to bind large numbers of calcium ions. Calcium binds to the thrombospondins through a cooperative mechanism that involves a significant conformational change in the protein. Through interactions with molecules on the cell surface and components of the extracellular matrix, the



thrombospondins play a role in cell adhesion, migration, differentiation, and proliferation. Thrombospondin-1 (TSP-1) was the first member of the gene family to be identified and has been the most extensively characterized. TSP-1 is a large multifunctional glycoprotein with a molecular weight of 420,000 Da, and is a trimer composed of identical subunits each with a molecular weight of 142,000 Da. TSP-1 is expressed by both normal and tumor cells and has a number of domains that allow it to interact with cells and other proteins. These include (1) a heparin-binding domain that interacts with proteoglycans, integrin α3β1, and cell-surface glycosaminoglycans (Clezardin et al., 1997; Merle et al., 1997); (2) three type 1 repeats that interact with CD36, matrix metalloproteinases, fibronectin, and heparan sulfate proteoglycans, and also activate latent TGF-β (Bornstein, 1995; Schultz-Cherry et al., 1995; Crawford et al., 1998); (3) an RGDA sequence within the last type 3 repeat, which interacts with integrin αvβ3; and (4) a C-terminal cellbinding domain that contains a recognition sequence for the integrin-associated protein CD47 (Gao et al., 1996). In this unit, the authors focus on the activities of TSP-1 that involve the type 1 repeats and the interaction of TSP-1 with integrins (Fig. 10.10.1). The interaction of TSP-1 with proteoglycans is discussed in detail in a recent review by Chen et al. (2000).

NH2

procollagen

FQGVLQNVRFVF

type 1

TSP-1 and transforming growth factor-â Recently, TSP-1 has been shown to activate transforming growth factor-β (TGF-β) by binding to the latency-associated protein and altering the conformation of TGF-β to make it accessible to its receptor (Schultz-Cherry et al., 1995; Crawford et al., 1998). The region of TSP-1 responsible for TGF-β activation is the amino acid sequence KRFK, which is found at the start of the second type I repeat (SchultzCherry et al., 1995; Crawford et al., 1998; Fig. 10.10.1). TGF-β is a 25-kDa homodimeric cytokine and a known tumor suppressor (Markowitz and Roberts, 1996). It is secreted in a latent complex consisting of mature TGF-β, the latency-associated protein, and sometimes an additional latent TGF-β-binding protein. The latent TGF-β-binding protein is thought to target latent TGF-β to sites in the extracellular matrix where it is sequestered until activated. Activation of TGF-β has been demonstrated in vitro by activators such as acids, plasmin, or cathepsin D (Munger et al., 1997). TSP-1 and the αvβ6 integrin have been shown to activate TGF-β in vivo (Crawford et al., 1998; Munger et al., 1999). Activation of TGF-β by TSP-1 was demonstrated in vivo when TSP-1-deficient mice were injected with a peptide containing the sequence KRFK. The lungs of the injected mice became morphologically more similar to wild-type mice and active TGF-β was detected in the bronchial epithelial cells (Crawford et al., 1998). In some contexts, however, TSP-1 does not appear to be a good activator of TGF-β

type 2

KRFK

CD36 CD36

α3β1

type 3

COOH

RFYVVMWK RGD

αvβ3 CD47

ss heparinbinding domain 25,000 Da

70,000 Da proteolytic fragment

Figure 10.10.1 Representative model of TSP-1 identifying the different structural and functional domains. The binding sites for the various integrins, CD36, and CD47 are indicated below the model. Amino acid sequences that mediate receptor binding and activation of TGF-β are indicated above the model. The proteolytic fragments isolated in the protocol are shown at the bottom. The FQGVLQNVRFVF sequence is a GAG-independent cell binding site and the RFYVMWK sequence is an integrin-associated protein (CD47) binding site.



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(Abdelouahed et al., 2000; Grainger and Frow, 2000). These data indicate that post-translational modification or other factors may regulate the ability of TSP-1 to activate TGF-β. Thus, co-expression of TSP-1 and TGF-β does not necessarily mean that TSP-1 will activate latent TGF-β in that tissue.


The role of TSP-1 in angiogenesis and cancer TSP-1 has been shown to be an effective inhibitor of angiogenesis, tumor progression, and metastasis (Chen et al., 2000; Lawler, 2002). While TSP-1 levels are very low in many tumor cells, expression of TSP-1 is high in the tumor stroma (Brown et al., 1999). Overexpression of TSP-1 in MDA-MB-435 human breast carcinoma cells decreased tumorigenesis and metastasis in vivo (Weinstat-Saslow et al., 1994). Furthermore, the tumors derived from cells formed by a fusion of low-TSP-1-expressing human breast cancer cells and high-TSP-1expressing normal breast epithelial cells were smaller in nude mice as compared to the tumors formed from the breast cancer cells alone (Zajchowski et al., 1990). Lastly, one group has shown that plasma TSP-1 secreted from primary HT1080 fibrosarcomas in nude mice inhibited growth of experimental metastases (Volpert et al., 1998). Moreover, if the implanted fibrosarcoma cells were transfected with an antisense TSP-1 construct prior to implantation, melanoma cell invasion of the lung was not inhibited. Recently, the authors have shown that recombinant proteins comprising the second type 1 repeat of TSP-1 and containing the TGF-β activating sequence KRFK inhibited B16F10 tumor growth in mice (Miao et al., 2001). Furthermore, it was observed that treatment with a TGF-β antibody or soluble TGF-β receptor reversed this inhibition, suggesting that TSP-1 activation of TGF-β is part of the inhibitory pathway. By contrast, an effect of TGF-β was not observed with Lewis lung carcinoma because these cells have acquired mutations that have rendered them unresponsive. Vascular density was decreased in both B16F10 and Lewis lung carcinoma tumors treated with the recombinant proteins through a TGF-β-independent mechanism. In another study, Streit et al. (1999) overexpressed full-length TSP-1 in A431 human carcinoma cells and implanted these cells in the flanks of nude mice. Decreased tumor growth and angiogenesis were observed in tumors expressing TSP-1. Recent work has demonstrated

that the KRFK sequence in the second type 1 repeat of TSP-1 is partly responsible for this growth inhibition and the decrease in tumor angiogenesis (K. Yee, unpub. observ.). In another recent study, TSP-1 null mice were crossed with c-neu transgenic mice to create a mouse that develops breast tumors and does not express TSP-1. These mice developed tumors that were larger and more vascular than the tumors of mice overexpressing TSP-1 (Rodrídguez-Manzaneque et al., 2001). The authors also determined that the absence of TSP-1 in these tumors resulted in an increase in the amount of active matrix metalloproteinase 9 (MMP-9). The effects of TSP-1 on endothelial cell migration and angiogenesis have been previously observed by several groups (Tolsma et al., 1993; Dawson et al., 1997; Qian et al., 1997; Iruela-Arispe et al., 1999; Jiménez et al., 2000; Nör et al., 2000). These studies demonstrate that TSP-1 is able to prevent tumor progression in several in vivo cancer models and that one of the ways TSP-1 inhibits tumor growth may be through decreasing tumor angiogenesis. In a different avenue of thinking, many groups have examined MMP-2 and MMP-9 with regards to breast cancer progression (Benaud et al., 1998; Martorana et al., 1998; Remacle et al., 1998; Rudolph-Owen et al., 1998; Lee et al., 2001). MMP-2 and -9 are gelatinases that degrade collagen types IV, V, VII, and X, as well as denatured collagen and gelatin (Dollery et al., 1995). Recently, TSP-1 has been shown to interact with MMP-2 and -9 and inhibit their activation (Bein and Simons, 2000; Rodrídguez-Manzaneque et al., 2001). This interaction is mediated by the type 1 repeats of TSP-1. Therefore, one of the mechanisms through which TSP-1 inhibits both tumor progression and tumor angiogenesis may be due to its ability to inhibit MMP activation and prevent growth factor and cell mobilization. Angiogenesis is a complex process that involves multiple cell types. TSP-1 does have possible effects on the recruitment of immune cells and on the proliferation and migration of vascular smooth muscle cells. In some assays, these effects can predominate, leading to the conclusion that TSP-1 supports angiogenesis. The preponderance of in vivo data indicates that the anti-angiogenic effects predominate in tumors. TSP-1 and CD36 CD36 is an integral membrane glycoprotein, a member of the class B scavenger receptor



family, and is located within the caveolae of the cell membrane. It is expressed in many cells including microvascular endothelium, adipocytes, skeletal muscle, dendritic cells, and hematopoietic cells including platelets and macrophages (Febbraio et al., 2001). CD36 is also a receptor for TSP-1 and binds to the specific sequence CSVTCG in the second and third type 1 repeats of TSP-1, while TSP-1 type 1 repeats bind the CD36 LIMP-II Emp sequence homology (CLESH) region of CD36 (Crombie and Silverstein, 1998). This binding initiates a signal that involves the nonreceptor tyrosine kinases fyn, lyn, and yes as well as p38MAPK (Huang et al., 1991). One of the endpoints of this cascade is activation of caspase 3 and endothelial cell apoptosis (Guo et al., 1997; Jiménez et al., 2000; Nör et al., 2000). CD36 signaling is one of the mechanisms by which TSP-1 inhibits angiogenesis and tumor progression (Dawson et al., 1997; Simantov et al., 2001). The initial work on exploring the anti-angiogenic effect of TSP-1 through CD36 utilized peptides containing the CSVTCG sequence. These peptides inhibited endothelial cell migration and angiogenesis (Iruela-Arispe et al., 1991; Tolsma et al., 1993; Dawson et al., 1999). Antibodies to CD36 also inhibited endothelial cell migration (Dawson et al., 1997) and, in CD36-null mice, TSP-1 did not inhibit angiogenesis in a cornea pocket assay (Jiménez et al., 2000). Therefore, binding of TSP-1 to CD36 on endothelial cells inhibits angiogenesis and tumor progression. TSP-1 and integrins Integrins are a family of cell surface receptors composed of both an α and a β subunit (Hynes, 1992). TSP-1, in both soluble and matrix-bound forms, can interact with β1 and β3 integrins; however, the physiological consequences of binding are dependent upon the integrin engaged, the cell type, and in some cases the involvement of accessory proteins. TSP-1 and â1 integrins In breast carcinoma cells, α3β1 is essential for chemotaxis towards TSP-1 and cell spreading on an immobilized TSP-1 matrix (Chandrasekaran et al., 1999). This interaction is mediated through binding of the integrin to residues 190 to 201 of the N-terminal region of TSP-1 (Krutzsch et al., 1999). In the presence of a β1-activating antibody, the adhesive properties of the carcinoma cells on TSP-1 are enhanced. This is characterized by rearrange-

ment of F actin filaments into filopodia that terminate at points that are rich in β1 and are in contact with TSP-1. Signaling through the insulin-like growth factor-I receptor (IGF-IR) can also potentiate this adhesion. Recent evidence suggests that IGF-IR signaling activates α3β1 by promoting association with the mitochondrial molecule heat shock protein 60 (Barazi et al., 2002). Small-cell lung carcinoma cells also bind residues 190 to 201 of TSP-1 through α3β1 (Guo et al., 2000). This interaction stimulates the cells to extend neurite-like processes and differentiate along a neuronal pathway. When epidermal growth factor is added to these cultures, binding to TSP-1 through this receptor also suppresses cell proliferation. This mechanism may be important for the antitumorigenic effects of TSP-1. In response to loss of cell-cell contact, endothelial cells engage immobilized TSP-1 through α3β1 and are stimulated to adhere to TSP-1 and proliferate (Chandrasekaran et al., 2000). This effect can be induced through disruption of cell contacts through wounding or by inhibiting vascular endothelial (VE) cadherin, indicating a role for TSP-1 in supporting repair of wounded endothelium. However, classically, TSP-1 is known for inhibiting endothelial cell proliferation and angiogenesis (Good et al., 1990). Indeed, endothelial cells exposed to a soluble TSP-1 peptide that recognizes α3β1 have decreased proliferation and motility (Chandrasekaran et al., 2000). These opposing effects on endothelial cells suggest that tight regulation of TSP-1/α3β1 interaction and signaling exists. Recent studies using melanoma cells demonstrated that the ability of TSP-1 to bind α3β1 is altered when TSP-1 is bound to fibronectin (Rodrigues et al., 2001). Conformational regulation of TSP-1 may represent one mechanism by which integrin-mediated cellular responses are controlled. Activated T-lymphocytes can adhere to intact TSP-1 through α4β1 and α5β1 integrins (Yabkowitz et al., 1993). This may have implications for mediating T cell activation, as stimulation of the ERK pathway by TSP-1 in these cells can be inhibited using anti-β1 function-blocking antibodies (Wilson et al., 1999). A role for TSP-1 in modulating the inflammatory response would not be surprising since TSP-1-deficient mice suffer from inflammatory disease (Lawler et al., 1998). Extracellular Matrix


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TSP-1 and â3 integrins In platelets, it was originally discovered that αvβ3 and, to a lesser extent, αIIbβ3 (GPIIbIIIa) function as adhesion receptors for TSP-1. The recognition site for these integrins is the RGD motif located in the type 3 repeats of TSP-1. TSP-1 can influence integrin function directly and indirectly through its interaction with nonintegrin receptors. In platelets, binding of the C terminus of TSP-1 to the transmembrane receptor integrin-associated protein (IAP or CD47) leads to assembly of a TSP-1/IAPαIIbβ3 complex on the platelet surface. This complex can further activate αIIbβ3 and cause phosphorylation of focal adhesion kinase, resulting in both augmentation of platelet aggregation and attachment to fibrinogen (Chung et al., 1997). A necessity for G-protein signaling has since been added to this cascade of events (Frazier et al., 1999). TSP-1/IAPαvβ3 complexes are also important in other cell types. On vitronectin substrates, C32 human melanoma cells are stimulated to spread in response to complex formation (Gao et al., 1996). More recently, an increase in latent TGF-β activation, induced by tamoxifen treatment of breast carcinoma cells, has been shown to be dependent on localization of TSP-1 to the cell surface by this mechanism (Harpel et al., 2001). Another example of TSP-1 affecting integrin function through cooperation with other receptors occurs in the clearance of apoptotic neutrophils. Here, TSP-1 associates with CD36 on the macrophage surface and αvβ3 associates on the neutrophils where it forms a bridge, allowing the recognition of neutrophils for ingestion (Savill et al., 1992). This process can be modulated on a second exposure of macrophages to neutrophils by ligation of αvβ3, α6β1, and α1β2 (Erwig et al., 1999). αvβ3 is also expressed on endothelial cells. In sickle cell anemia patients, both αvβ3 in the endothelium (Solovey et al., 1999) and TSP-1 plasma levels are elevated. These proteins have been implicated in recurring vaso-occlusion problems in sickle cell patients caused by exaggerated adhesion of the sickle cell red blood cells (SS-RBCs) to the endothelium. Indeed, it has been demonstrated that TSP-1 enhances adhesion of SS-RBCs to cultured endothelial cells and that antibodies to αvβ3 can block this event (Kaul et al., 2000). It is as yet unknown if this is a direct consequence of TSP-1/αvβ3 association.

Critical Parameters The response of the platelets to thrombin is a critical factor contributing to the success of the purification procedure. Since platelets become less responsive during storage, the platelet-rich plasma should be processed as soon as possible after collection. Since platelets are temperature sensitive, buffers and centrifuges used in the purification procedure should be warmed to room temperature before beginning the procedure. The platelets should also be handled gently during the resuspension steps to prevent mechanical activation. Moreover, since platelets are activated by untreated glass surfaces, all transfer pipets and tubes should be plastic. TSP-1 is susceptible to proteolysis following its secretion into the supernatant. It is important to work quickly following the activation step to minimize exposure to proteases secreted from the platelets and the thrombin used for the activation. The supernatant should be treated immediately with DFP following the debrisclearing centrifugation step in order to inactivate these proteases. The supernatant should be kept on ice at all times during the remaining purification steps. The association of TSP-1 with calcium maintains the confirmation of the molecule. It is therefore essential that calcium be present in all solutions during and subsequent to thrombin treatment. A concentration of 2 mM is recommended.

Troubleshooting The problem most likely to be encountered in the purification procedure is unresponsive platelets. To remedy this situation the procedure can be performed on a small scale using fresh platelets. This will provide a sense of how the aggregated platelets should appear following thrombin treatment. Another method for assaying platelet responsiveness is to perform electrophoresis on the supernatant from the thrombin-treated platelets. TSP-1 is a major component of the platelet α-granule and should appear as a prominent band running at an apparent molecular weight of 185,000 Da on discontinuous Laemmli SDS gels (UNIT 6.1). This anomalously high value for the molecular weight of the subunit is probably due to a decrease in the amount of SDS bound to the large number of negatively charged residues in the type 3 repeats.



Anticipated Results The purification procedure should result in producing ∼200 µg TSP-1 per 100 ml outdated platelet-rich plasma, which is most often >95% pure as determined by SDS-PAGE. There is evidence that some preparations of TSP-1 produced according to this method may contain trace amounts of active TGF-β bound to the TSP-1. It is possible to remove this contaminant by adjusting the pH of the sucrose gradient solutions to pH 11, as TGF-β will dissociate from TSP-1 under alkaline conditions (Murphy-Ullrich et al., 1992; Schultz-Cherry et al., 1994). The pH of the TSP-1-containing fractions should be returned to pH 7.6 immediately following centrifugation. Whereas the protocol for purifying TSP-1 proteolytic fragments does not require many steps and is reasonably efficient, it is important to bear in mind that the N-terminal domain only represents ∼18% of the total mass of the protein. Thus, if one starts with 5 mg total protein, a yield of 400 to 500 µg is appropriate. Since the 210,000-Da fragment represents about onehalf of the protein, yields of 1 to 1.5 mg can be expected.

Time Considerations The purification procedure is extended over a period of 3 days. The amount of time required to perform this procedure will depend in part on the amount of material to be processed. Approximately 3 to 4 hr should be allowed to isolate the TSP-1-containing supernatant (steps 1 to 15). Purification of TSP-1 (steps 17 to 28) will require another 3 to 4 hr. It is possible to leave the TSP-1 bound to the heparinSepharose column for a number of days prior to continuing the elution process. The purification of proteolytic fragments also takes ∼3 days. The limited tryptic digestion is done overnight. Elution of the heparinSepharose column can be done in ∼1 day and the elution of the G-200 column requires another day.

LITERATURE CITED Abdelouahed, M., Ludlow, A., Brunner, G. and Lawler, J. 2000. Activation of platelet-transforming growth factor β-1 in the absence of thrombospondin-1. J. Biol. Chem. 275:1793317936. Adams, J. 2001. Thrombospondins: Multifunctional regulators of cell interactions. Annu. Rev. Cell Dev. Biol. 17:25-51. Barazi, H.O., Zhou, L., Smyth Templeton, N., Krutzsch, H.C., and Roberts, D.D. 2002. Identification of heat shock protein 60 as a molecular

mediator of α3β1 integrin activation. Cancer Res. 62:1541-1548. Bein, K. and Simons, M. 2000. Thrombospondin-1 type 1 repeats interact with matrix metalloproteinase 2: Regulation of metalloproteinase activity. J. Biol. Chem. 275:32167-73. Benaud, C., Dickson, R.B. and Thompson, E.W. 1998. Roles of the matrix metalloproteinases in mammary gland development and cancer. Breast Cancer Res. Treatment 50:97-116. Bornstein, P. 1995. Diversity of function is inherent in matricellular proteins: An appraisal of thrombospondin 1. J. Cell Biol. 130:503-506. Brown, L.F., Guidi, A.J., Schnitt, S.J., Water, L.V.D., Iruela-Arispe, M.L., Yeo, T.-K., Tognazzi, K., and Dvorak, H.F. 1999. Vascular stroma formation in carcinoma in situ, invasive carcinoma and metastatic carcinoma of the breast. Clin. Cancer Res. 5:1041-1056. Chandrasekaran, S., Guo, N.-H., Rodrigues, R.G., Kaiser, J., and Roberts, D.D. 1999. Pro-adhesive and chemotactic activities of thrombospondin-1 for breast carcinoma cells are mediated by a3b1 integrin and regulated by insulin-like growth factor 1 and CD98. J. Biol. Chem. 274:11408-11416. Chandrasekaran, L., He, C.H., Al-Barazi, H., Krutzsch, H.C., Iruela-Arispe, M.L., and Roberts, D.D. 2000. Cell-contact-dependent activation of α3β1 integin modulates endothelial cell responses to thrombospondin-1. Mol. Biol. Cell 11:2885-2900. Chen, H., Herndon, M.E., and Lawler, J. 2000. The cell biology of thrombospondin-1. Matrix Biol. 19:597-614. Chung, J., Gao, A., and Frazier, W.A. 1997. Thrombospondin acts via integrin associated protein to activate the platelet integrin αIIbβ3. J. Biol. Chem. 272:14740-14746. Clezardin, P., Lawler, J., Amiral, J., Quentin, G., and Delmas, P. 1997. Identification of cell adhesive active sites in the N-terminal domain of thrombospondin-1. Biochem. J. 321:819-827. Crawford, S.E., Stellmach, V., Murphy-Ullrich, J.E., Ribeiro, S.M.F., Lawler, J., Hynes, R.O., Boivin, G.P. and Bouck, N. 1998. Thrombospondin-1 is a major activator of TGF-β1 in vivo. Cell 93:1159-1170. Crombie, R. and Silverstein, R. 1998. Lysomsomal integral membrane protein II binds thrombospondin-1. J. Biol. Chem. 273:4855-4863. Dawson, DW., Pearce, S.F.A., Zhong, R., Silverstein, R.L., Frazier, W.A., and Bouck, N.P. 1997. CD36 mediates the in vitro inhibitory effects of thrombospondin-1 on endothelial cells. J. Cell Biol. 138: 707-717. Dawson, D.W., Volpert, O.V., Pearce, S.F.A., Schneider, A.J., Silverstein, R.L., Henkin, J., and Bouck, N. 1999. Three distinct d-amino acid substitutions confer potent antiangiogenic activity on an inactive peptide derived from a thrombospondin-1 type 1 repeat. Molec. Pharmacol. 55:332-338.



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Markowitz, S.D. and Roberts, A.B. 1996. Tumor suppressor activity of the TGF-βpathway in human cancers. Cytokine Growth Factor Rev. 7:93102. Martorana, A.M., Zheng, G., Crowe, T.C., O’Grady, R.L., and Lyons, J.G. 1998. Epithelial cells upregulate matrix metalloproteinases in cells within the same mammary carcinoma that have undergone an epithelial-mesenchymal transition. Cancer Res. 58:4970-4979. Merle, B., Malaval, L., Lawler, J., Delmas, P., and Clezardin, P. 1997. Decorin inhibits cell attachment to thrombospondin-1 by binding to a KKTR-dependent cell adhesive site present within the N-terminal domain of thrombospondin-1. J. Cell. Biochem. 67:75-83. Miao, W.-M., Seng, W.L., Duquette, M., Lawler, P., Laus, C., and Lawler, J. 2001. Thrombospondin1 type 1 repeat recombinant proteins inhibit tumor growth through transforming growth factor β dependent and independent mechanisms. Cancer Res. 61:7830-7839.



Munger, J.S., Harpel, J.G., Gleizes, P.-E., Mazzieri, R., Nunes, I., and Rifkin, D.B. 1997. Latent transforming growth factor-β: Structural features and mechanisms of activation. Kidney Int. 51:1376-1382. Munger, J.S., Huang, X., Kawakatsu, H., Griffiths, M.J.D., Dalton, S.L., Wu, J., Pittet, J.-F., Kaminski, N., Garat, C., Matthay, M.A. et al. 1999. The integrin αvβ6 binds and activates latent TGFβ-1: A mechanism for regulating pulmonary inflammation and fibrosis. Cell 96:319-328. Murphy-Ullrich, J.E., Schultz-Cherry, S., and Höök, M. 1992. Transforming growth factor-βcomplexes with thrombospondin. Mol. Biol. Cell 3:181-188. Nielsen, B.S., Sehested, M., Kjeldsen, L., Borregaard, N., Rygaard, J., and Danø, K. 1997. Expression of matrix metalloprotease-9 in vascular pericytes in human breast cancer. Lab. Invest. 77:345-355. Nör, J.E., Mitra, R.S., Sutorik, M.M., Mooney, D.J., Castle, V.P., and Polverini, P.J. 2000. Thrombospondin-1 induces endothelial cell apoptosis and inhibits angiogenesis by activating the caspase death pathway. J. Vasc. Res. 37:209-218. Qian, X., Wang, T.N., Rothman, V. L., Nicosia, R.F., and Tuszynski, G.P. 1997. Thrombospondin-1 modulates angiogenesis in vitroby up-regulation of matrix metalloproteinase-9 in endothelial cells. Exper. Cell Res. 235:403-412. Remacle, A.G., Noël, A., Duggan, C., McDermott, E., O’Higgins, N., Foidart, J.M., and Duffy, M.J. 1998. Assay of matrix metalloproteinases types 1, 2, 3 and 9 in breast cancer. Br. J. Cancer 77:926-931. Rodrigues, R.G., Guo, N.-H., Zhou, L., Sipes, J.M., Williams, S.B., Smyth Templeton, N., Gralnick, H.R., and Roberts, D.D. 2001. Conformational regulation of the fibronectin binding and α3β1 integrin-mediated adhesive activities of thrombospondin-1. J. Biol. Chem. 276:27913-27922. Rodrídguez-Manzaneque, J.C., Lane, T.F., Ortega, M.A., Hynes, R.O., Lawler, J., and IruelaArispe, M.L. 2001. Thrombospondin-1 suppresses spontaneous tumor growth and inhibits activation of matrix metalloproteinase-9 and moblization of vascular endothelial growth factor. Proc. Natl. Acad. Sci. U.S.A. 98:1248512490. Rudolph-Owen, L.A., Chan, R., Muller, W.J., and Matrisian, L.M. 1998. The matrix metalloproteinase matrilysin influences early-stage mammary tumorigenesis. Cancer Res. 58:5500-5506. Savill, J., Hogg, N., Ren, Y., and Haslett, C. 1992. Thrombospondin cooperates with CD36 and the vitronectin receptor in macrophage recognition of neutrophils undergoing apoptosis. J. Clin. Invest. 90:1513-1522. Schultz-Cherry, S., Ribeiro, S., Gentry, L., and Murphy-Ullrich, J.E. 1994. Thrombospondin binds and activates the small and large forms of latent transforming growth factor-βin a chemically defined system. J. Biol. Chem. 269:26775-26782.

Schultz-Cherry, S., Chen, H., Mosher, D.F., Misenheimer, T.M., Krutzsch, H.C., Roberts, D.D., and Murphy-Ullrich, J.E. 1995. Regulation of transforming growth factor-βactivation by discrete sequences of thrombospondin-1. J. Biol. Chem. 270:7304-7310. Simantov, R., Febbraio, M., Crombie, R., Asch, A.S., Nachman, R.L., and Silverstein, R.L. 2001. Histidine-rich glycoprotein inhibits the antiangiogenic effect of thrombospondin-1. J. Clin. Invest. 107:45-52. Solovey, A., Gui, L., Ramakrishnan, S., and Hebbel, R.P. 1999. Sickle cell anemia as a possible state of enhanced anti-apoptotic tone: Survival effect of vascular endothelial growth factor on circulation and unanchored endothelial cells. Blood 93:3824-3830. Streit, M., Velasco, P., Brown, L.F., Skobe, M., Richard, L., Riccardi, L., Lawler, J., and Detmar, M. 1999. Overexpression of thrombospondin-1 decreases angiogenesis and inhibits the growth of human cutaneous squamous cell carcinomas. Am. J. Pathol. 155:441-452. Tolsma, S.S., Volpert, O.V., Good, D.J., Frazier, W.A., Polverini, P.J., and Bouck, N. 1993. Peptides derived from two separate domains of the matrix protein thrombospondin-1 have antiangiogenic activity. J. Cell Biol. 122:497-511. Volpert, O.V., Lawler, J., and Bouck, N.P. 1998. A human fibrosarcoma inhibits systemic angiogenesis and the growth of experimental metastases via thrombospondin-1. Proc. Natl. Acad. Sci. U.S.A. 95:6343-6348. Weinstat-Saslow, D.L., Zabrenetzky, V.S., VanHoutte, K., Frazier, W.A., Roberts, D.D., and Steeg, P.S. 1994. Transfection of thrombospondin 1 complementary DNA into a human breast carcinoma cell line reduces primary tumor growth, metastatic potential, and angiogenesis. Cancer Res. 54:6504-6511. Wilson, K.E., Li, Z., Kara, M., Gardner, K.L., and Roberts, D.D. 1999. β1 integrin- and proteoglycanmediated stimulation of T lymphoma cell adhesion and mitogen-activated protein kinase signaling by thrombospondin-1 and thrombospondin-1 peptides. J. Immunol. 163:3621-3628. Yabkowitz, R., Dixit, V.M., Guo, N., Roberts, D.D., and Shimizu, Y. 1993. Activated T-cell adhesion to thrombospondin is mediated by the α4β1 (VLA-4) and α5β1 (VLA-5) integrins. J. Immunol. 151:149-158. Zajchowski, D.A., Band, V., Trask, D.K., Kling, D., Connolly, J.L., and Sager, R. 1990. Suppression of tumor-forming ability and related traits in MCF-7 human breast cancer cells by fusion with immortal mammary epithelial cells. Proc. Natl. Acad. Sci. U.S.A. 87:2314-2318.

Contributed by Karen O Yee, Mark Duquette, Anna Ludlow, and Jack Lawler Beth Israel Deaconess Medical Center Boston, Massachusetts



Supplement 17

Purification of SPARC/Osteonectin

UNIT 10.11

SPARC (secreted protein acidic and rich in cysteine) is a founding member of the matricellular group of proteins that have been shown to mediate interactions between cells and the extracellular matrix (ECM; Bornstein and Sage, 2002). Other proteins within this family include thrombospondins 1 and 2, osteopontin, tenascins C and X, and Cyr61. Over the last several years, a wealth of data, largely from mice with targeted disruptions of the respective genes, has emerged identifying various targets of the matricellular proteins that influence cell behavior—e.g., growth factors, cell-cycle regulatory proteins, ECM components, adhesion proteins and/or their receptors, cell survival, collagen fibrillogenesis, and immune cell function. In vivo, these effects can be translated into abnormalities in blood vessel morphogenesis and connective tissues, wound healing, bone formation, and responses to various types of injury. Therefore, study of one or more of the matricellular proteins affords insight from a somewhat unusual and underexplored perspective: the interface between the cell surface and the extracellular milieu. SPARC belongs to a family of several genes, only one other of which, SC1/hevin, has been characterized beyond a limited degree (Brekken and Sage, 2000). SPARC-null mice exhibit many phenotypic abnormalities that follow logically from the effects of SPARC on cultured cells (i.e., de-adhesion, antiproliferation, interaction with growth factors and ECM, and regulation of collagen production). These characteristics include (1) accelerated dermal wound healing and fibrovascular invasion of sponge implants, (2) reduced foreign body response, (3) thin skin with decreased collagen, which is deposited as small-diameter fibrils, (4) excessive accumulation of adipose tissue, (5) osteopenia, and (6) cataract formation (Bornstein and Sage, 2002). Providing a mechanistic explanation for any one of these phenotypes requires experiments, largely in vitro, with active purified protein in clearly defined assays with quantitative endpoints. This unit presents several protocols for the purification of SPARC (see Basic Protocol and Alternate Protocols 1, 2, and 3), and for the measurement of its biological activity and conformation (see Support Protocols 1 and 2). Since the end product—i.e., natural SPARC or recombinant (rSPARC)—differs according to the source, guidelines for the choice of each protocol, and its advantages and limitations, have been included with the Basic Protocol (purification of SPARC from cultured cells), Alternate Protocol 1 (rSPARC from E. coli), Alternate Protocol 2 (rSPARC from insect cells), and Alternate Protocol 3 (SPARC from blood platelets). A method for determining endotoxin levels is presented in Support Protocol 3. NOTE: To prevent denaturation of SPARC due to adsorption to surfaces, only polypropylene or siliconized glass should be used. NOTE: All solutions and equiptment coming into contact with live cells should be sterile and a septic technique should be used accordingly PURIFICATION OF SPARC FROM PYS-2 CELLS This protocol describes the purification of SPARC from cultured PYS-2 cells. This cell line, originally derived from a mouse parietal yolk sac carcinoma, has been a consistent reproducible source of biologically active SPARC for nearly two decades (Sage and Bornstein, 1995). The following procedure can be applied to most cell culture supernatants and involves essentially three steps: (1) precipitation of culture medium, (2) ion-exchange chromatography, and (3) molecular-sieve chromatography. Advantages of the PYS-2 cell line are its immortality, its high rate of growth, its copious production (secretion) of SPARC, and the presence of few other secreted products in the culture medium. It is also possible to radiolabel SPARC metabolically if desired. A commercial Contributed by E. Helene Sage Current Protocols in Cell Biology (2003) 10.11.1-10.11.23 Copyright © 2003 by John Wiley & Sons, Inc.

BASIC PROTOCOL

Data Processing and Analysis


source of SPARC, isolated according to this protocol and of ∼80% purity, is available from Sigma-Aldrich. Materials 50% to 70% confluent PYS-2 cells (see recipe) DMEM (serum-free; APPENDIX 2A) 1100 Ci/mmol (12.5 Ci/ml) [trans-35S]methionine/cysteine (ICN; optional) DMEM minus methionine and cysteine (optional) 0.2 M PMSF stock solution (see recipe) N-Ethylmaleimide (NEM) Ammonium sulfate, ultrapure DEAE buffer, 4°C (see recipe) NaCl ∼2 × 20–cm DEAE column (see recipe) S-200 buffer (see recipe) Scintillation fluid (optional) Sephacryl molecular-sieve column (see recipe) 0.05 M acetic acid Plastic pipets 50-ml polycarbonate high-speed centrifuge tubes Low-speed GPKR (Beckman) centrifuge with swinging bucket rotor High-speed refrigerated centrifuge with GSA (Sorvall) or JA-17 rotors (Beckman) or equivalent 12,000- to 14,000-MWCO dialysis tubing (Spectrapor) or equivalent, prewashed with DEAE buffer Dialysis clips (optional) Standard gradient maker (e.g., Amersham Biosciences) Peristaltic pump Fraction collector Lyophilizer 50 or 250 ml centrifuge tubes Additional reagents and equipment for SDS-PAGE (UNIT 6.1) with autoradiography (UNIT 6.3), if appropriate, and determination of protein concentration by spectroscopy (APPENDIX 3B) CAUTION: When working with radioactivity, take appropriate precautions to avoid contamination of the experimenter and surroundings. Carry out the experiments and dispose of wastes in appropriately designated area, following guidelines provided by the local radiation safety officer (also see APPENDIX 1D). Collect and precipitate tissue culture medium containing secreted SPARC 1. Replace medium in 20 to 30 dishes or flasks of PYS-2 cells (grown to 50% to 70% confluency) with 12 to 13 ml serum-free DMEM and preincubate 15 min at 37°C. Replace with fresh medium and then incubate 18 to 24 hr. If desired, purification can be monitored by adding 500 ìCi of 1100 Ci/mmol [35S]methionine to one dish and processing the medium in parallel with nonlabeled medium from the other dishes. Alternatively, if radiolabeled SPARC of high specific activity is required for experimental purposes, [35S]methionine/cysteine can be added to all dishes. When using label, incubate cells in serum-free DMEM lacking methionine and cysteine. Purification of SPARC/Osteonectin

2. Collect the medium from the cell layer by gentle aspiration via plastic pipet and transfer to centrifuge tubes. Remove cellular debris by centrifuging in a clinical (i.e.,



tissue-culture) centrifuge 5 min at 1,000 × g, room temperature, or in GPKR centrifuge at 1000 × g, 4°C. 3. Pool all supernatants in a siliconized flask. Add 0.2 M PMSF drop-wise with stirring to a final concentration of 0.2 mM, and NEM to a final concentration of 10 mM. Stir on ice until medium reaches 4°C. For 100 ml medium, add 0.1 ml PMSF stock solution and 62.5 mg NEM. Take care not to lyse cells in any of these procedures.

4. Add solid ultrapure ammonium sulfate to the medium in an amount equivalent to 50% (w/v) of the starting volume over a period of several hours. Stir 12 to 24 hr at 4°C. For 100 ml medium, add 50 g ammonium sulfate, in very small increments (e.g., 1 to 2 g) over several hours (e.g., 3 to 5). This detail is important for maintenance of neutral pH and for efficient precipitation of protein, which consists mainly of laminin 1, type IV collagen, bovine serum albumin (BSA), and SPARC. Do not allow the solution to foam by stirring too rapidly, as this indicates the proteins are denaturing.

5. Transfer medium to 50-ml polycarbonate high-speed centrifuge tubes and centrifuge in a high-speed refrigerated centrifuge with JA-17 rotor 30 min at 40,000 × g, 4°C. Discard the supernatant. Keep tubes containing pellets on ice or store up to 1 to 2 months at –70°C. 6. Thaw, if necessary, and dissolve each pellet by gentle vortexing in 2 to 5 ml DEAE buffer, 4°C. Pool these solutions and transfer to 12,000- to 14,000-MWCO dialysis tubing, prewashed with DEAE buffer and closed on one end. Rinse each centrifuge tube with 1 ml buffer and add this solution to the bag. 7. Close the open end of the dialysis bag with double knots or dialysis clips, leaving 1 to 2 in. (2.5 to 5 cm) extra space to allow for change in volume. Immerse the bag (containing ∼40 ml) in a 500-ml graduated cylinder containing 500 ml DEAE buffer, 4°C. Dialyze with stirring overnight (or 4 to 6 hr), and change the dialysis buffer twice (2 to 3 hr each) for an additional 4 to 6 hr dialysis. Wear gloves when handling dialysis tubing to minimize exposure to radioactivity as well as to protect the sample from contamination. Mix bag contents several times by inversion.

8. Remove dialysis tubing, cut tip off carefully (if knotted) or remove clips, and empty contents into one or two 50-ml centrifuge tubes. Clarify the solution by centrifuging in a JA-17 rotor 20 min at 10,000 × g, 4°C. If appropriate, retain 10 to 25 µl for scintillation counting and for SDS-PAGE (UNIT 6.1) with autoradiography (UNIT 6.3), as assessment of starting material. The sample is now ready for ion-exchange chromatography.

Chromatograph on DEAE cellulose 9. Prepare gradient buffer B by adding 2.336 g NaCl to 200 ml DEAE buffer (200 mM NaCl final). Fill the front chamber of a standard gradient maker (containing a stir bar or paddle) with 200 ml DEAE buffer (gradient buffer A) and the second chamber with 200 ml gradient buffer B. Ensure that the narrow opening between the two chambers is filled with gradient buffer A before adding gradient buffer B. An air block will inhibit flow of B into A.

10. Use a peristaltic pump to add the entire sample onto an ∼2 × 20–cm DEAE column, and follow with one to two column volumes DEAE buffer. Discard this eluate, which contains unbound protein. 11. If phenol red (from DMEM) is seen to bind to the resin, wash the column until it is no longer visible, or until the A280 of the flowthrough is at baseline.



Supplement 17

Phenol red will interfere with the monitoring of the column effluent at 280 nm.

12. Connect the gradient maker to the peristaltic pump for delivery to the column bed. Connect a fraction collector to the column and set to collect 3-ml fractions of eluate in polypropylene or siliconized glass tubes. Elute bound proteins with a linear gradient of 0% to 100% buffer B over ~300 ml. All chromatographic procedures must be carried out at 4°C. A less complicated alternative to the continuous gradient is the use of two stepwise elutions, the first consisting of 100 ml of 75 mM NaCl in DEAE buffer, followed by 100 ml of 175 mM NaCl in DEAE buffer. SPARC will elute in the second buffer.

13. For radiolabeled SPARC (step 2), monitor the effluent by scintillation counting 20µl aliquots from alternate fractions suspended in 3 ml scintillation fluid. For nonradiolabeled SPARC, monitor alternate fractions by absorbance at 280 nm. SPARC is eluted at 150 to 175 mM NaCl. See Sage et al. (1989) for an example of the elution profile. If the location of the peak containing SPARC is in doubt, individual fractions can be analyzed by SDS-PAGE (UNIT 6.1).

14. Pool fractions containing SPARC, and dialyze the pooled sample (∼20 ml) against four changes of 4 liters (each) water over 24 to 48 hr, 4°C (see steps 6 to 8). After 24 to 48 hr, a precipitate containing SPARC, together with laminin and traces of BSA, should appear in the dialysis bag. Depending on the concentration of protein and/or the water used (pH 5.5 is optimal), precipitation may fail to occur. In this case, lyophilize the protein (step 16b), redissolve in DEAE buffer at 25% of the original volume, and repeat dialysis and precipitation (steps 14 and 15). If the column will be reused, it should be regenerated as described (see Reagents and Solutions).

15. Decant the entire contents of the bag into a centrifuge tube and centrifuge 30 min at 48,000 × g, 4°C. Discard the supernatant. 16a. For immediate use: Dissolve pellet in 2 ml S-200 buffer, clarify by microcentrifugation for 1 min at top speed or 10,000 × g, and proceed to molecular-sieve chromatography (step 18). 16b. For storage before chromatography: Resuspend pellet in 2 to 4 ml water, shell-freeze by twirling the tube in dry ice/ethanol to effect freezing of the solution on the sides of the vessel, and then lyophilize. Store up to 1 to 2 months at –70°C. Before use, resuspend in 1 to 2 ml S-200 buffer, stir 4 to 6 hr at 4°C, and clarify the solution by microcentrifugation at top speed for 1 min. Shell-freezing increases the efficiency of lyophilization and improves solubility of the protein after storage. Pellets from several preparations can be pooled prior to molecular-sieve chromatography.

Purify SPARC by molecular-sieve chromatography 17. Remove buffer from the top of a Sephacryl molecular-sieve column and apply the sample gently onto the resin. Allow the sample (optimally 1 to 2 ml) to flow into the bed. Add 2 to 4 ml S-200 buffer to the top of the column, reconnect the buffer reservoir, and allow effluent to flow by gravity at 8 to 10 ml/hr (0.17 ml/min) by adjustment of the pressure head (i.e., the reservoir containing S-200 buffer above the column). It is important not to disturb the column bed during sample loading, as the precision of elution can be affected. Purification of SPARC/Osteonectin



In some cases it may be necessary to use a peristaltic pump, pulling buffer from the bottom of the column, at ∼10 ml/hr. If the flow rate is too high, the column will pack too tightly and will cease to flow.

18. Collect 80 fractions of 1 to 1.5 ml each and monitor effluent by absorbance at 280 nm and/or by counting 10 to 25-µl aliquots in 3 ml scintillation fluid. The exact position of elution of SPARC will vary with chromatographic parameters (e.g., column size, sample size, flow rate). It is therefore advisable to monitor the column effluent and, if necessary, to check 10 to 25 ìl of each fraction by SDS-PAGE (see below). The initial peak (at Vo) contains laminin, whereas the leading shoulder of the peak corresponding to the elution position of SPARC contains most of the BSA.

19. Pool peak fractions corresponding to SPARC (approximately ten fractions, corresponding to 55 to 65 ml total column effluent). Dialyze this pool against four changes of 4 liters of 0.05 M acetic acid each, 4°C, and lyophilize. Alternatively, the sample can be stored at –70°C in S-200 buffer without dialysis or lyophilization, or it can be dialyzed directly into another buffer as desired.

20. Determine the concentration of SPARC by absorbance at 280 nm, using the extinction coefficient (ε) 0.838 mg ml−1 cm−1 (APPENDIX 3B). 21. Analyze the purified protein by SDS-PAGE (UNIT 6.1) with autoradiography (UNIT 6.3). When heating samples at 95°C, use reducing (i.e., 50 mM DTT) and nonreducing conditions. For detection using Coomassie blue, from 1 to 5 ìg SPARC is recommended; for detection by autoradiography, ∼104 cpm is recommended. A single broad band, or occasionally a doublet, should be obtained with an apparent Mr of 39,000 (with DDT) or 43,000 (without DDT), the latter co-migrating with an ovalbumin molecular weight standard. The yield of purified SPARC is ∼500 ìg per 30 maxiplates (150-mm diameter) of PYS-2 cells (2 to 3 × 108 cells).

PURIFICATION OF rSPARC FROM E. COLI The preceding procedure (see Basic Protocol) allows for the purification of murine SPARC from cultured (tumor) cells. Limitations of a mammalian cell culture system as a protein source are its cost, potential contamination of the product by serum and cellular proteins/proteinases, and the low yield of product. To circumvent these problems, Bassuk et al. (1996a) expressed human rSPARC with a C-terminal histidine tag in E. coli. A soluble (monomeric) form and an insoluble (aggregated) form of SPARC were recovered, the latter sequestered in inclusion bodies within the host. Soluble (monomeric) SPARC from E. coli is biologically active and can be purified in relatively large quantities with minimal contamination by endotoxin or bacterial proteins. Isolation of the soluble form is accomplished by anion-exchange, nickel-chelate affinity, and gel-filtration chromatographies. Anion-exchange chromatography on DEAE-Sepharose is used as an initial isolation step. Metal-chelate affinity chromatography provides an efficient purification of rSPARC that has been expressed with a (His)6 sequence. Gel-filtration chromatrography separates monomers of SPARC from dimers, trimers, and higher oligomers. This procedure is outlined below. It assumes that a competent strain of E. coli—e.g., BL21(DE3)—has been transformed with a SPARC expression plasmid—e.g., pSPARC wt (human)—with a hexahistidine (His)6 sequence at the 3′ end (Bassuk et al., 1996a) and has been propagated and frozen as a glycerol stock. Additionally, the aggregated form can be unfolded by urea treatment, purified by nickelchelate affinity chromatography, and renatured by gradual removal of the denaturant. After disulfide bond isomerization, the disaggregated monomers are further purified by




Supplement 17

high-resolution gel-filtration chromatography (Bassuk et al., 1996b). As the disaggregation/renaturation procedure is complicated and time consuming, the reader is referred to Bassuk et al. (1996b) for this additional protocol. Additional Materials (also see Basic Protocol) LB medium with appropriate selective reagents (APPENDIX 2A) E. coli strain transfected with SPARC expression vector (Bassuk et al., 1996a) Inducing agent (e.g., IPTG; APPENDIX 3A) 10 mM sodium phosphate, pH 7.0 (APPENDIX 2A)/10% (v/v) glycerol 90 mM sodium phosphate buffer (pH 7.8)/10% (v/v) glycerol/0.2 mM AEBSF, 4°C (see recipe), with and without 0.5 M NaCl DEAE-Sepharose Fast Flow anion-exchange resin (Amersham Biosciences): equilibrate in 90 mM sodium phosphate buffer (pH 7.8)/10% (v/v) glycerol/0.2 mM AEBSF and allow to settle 5 M NaCl (APPENDIX 2A) 0.2 M AEBSF stock solution (see recipe) Nickel/nitrilotriacetic acid (Ni-NTA) metal-chelate affinity resin (Qiagen) 50 mM sodium phosphate (pH 5.3, 6.0, and 7.8)/0.5 M NaCl/10% (v/v) glycerol (see recipe) 1.6 × 60–cm Superdex 70 column (see recipe) 50 mM Tris⋅Cl (pH 8.0)/0.15 M NaCl (see recipe) 1× PBS (APPENDIX 2A) containing 1 to 4 mM Ca2+ (optional) French press 2 × 20– and 1 × 10–cm chromatography columns Flow cell coupled to a UV monitor set at 280 nm Chart recorder Conductivity meter (optional) Disposable 10-ml gel-filtration column, sterile (optional) Additional reagents and equipment for transfecting SPARC expression vector (APPENDIX 3A) and for SDS-PAGE on minigels (UNIT 6.1) Extract E. coli 1. Inoculate 1.3 liters LB medium containing appropriate selective reagents with a suitable E. coli strain transfected with SPARC expression vector using standard techniques (APPENDIX 3A). Grow to midexponential phase (OD600 ∼0.5) and induce with the appropriate agent. Induction of rSPARC in midexponential phase cells is necessary for high levels of expression. The procedure and chemical(s) used depend on the E. coli strain and the vector into which SPARC cDNA is cloned. For example, IPTG was used at a final concentration of 1 mM for SPARC cloned into pET22b vector and transfected into strain BL21(DE3) (Bassuk et al., 1996a).

2. After the cells have been induced, grow an additional 1 to 4 hr. 3. Recover the cells by centrifuging 20 min at 7000 × g, room temperature. Discard the supernatant and resuspend the pellet in 20 ml of 10 mM sodium phosphate, pH 7.0, containing 10% (v/v) glycerol. Disrupt by performing two cycles in a French press at 20,000 psi. Cells can alternatively be broken open by sonication on ice. Purification of SPARC/Osteonectin

4. Separate soluble from insoluble material by centrifuging 30 min at 10,000 × g, 4°C. Decant soluble extract (supernatant) into a separate tube.



Soluble extracts and insoluble pellets at this stage can be stored up to 1 month at −80°C. Refer to Bassuk et al. (1996b) for details on processing pellets for aggregated SPARC.

Perform initial chromatography on DEAE-Sepharose 5. If necessary, thaw the soluble extracts on ice. Dilute to 100 ml with ice-cold 90 mM sodium phosphate buffer (pH 7.8)/10% (v/v) glycerol/0.2 mM AEBSF, 4°C. 6. Add 50 ml settled DEAE-Sepharose Fast Flow anion-exchange resin. Stir gently 12 to 18 hr at 4°C. 7. Pour slurry into a 2 × 20–cm chromatography column, allow to settle, and wash with ∼250 ml of 90 mM sodium phosphate buffer (pH 7.8)/10% (v/v) glycerol/0.2 mM AEBSF until the absorbance at 280 nm is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¤H[SUHVVHGSURWHLQFDQEHLPPX QRSUHFLSLWDWHG XVLQJ DQWL+$ DQWLERGLHV %HQHGLFWDQG&ODZVRQ $OWHUQDWLYHO\ LI DQ DQWLERG\ DJDLQVW RQH RI WKH SDUWQHUV LV DYDLODEOH GHWHFWLRQ FDQ EH DFFRPSOLVKHG E\

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Phosphoamino Acid Analysis

UNIT 14.5

It is often valuable to identify the phosphorylated residue in a protein. In the case of proteins phosphorylated at serine, threonine, or tyrosine, this is readily accomplished by partial acid hydrolysis in HCl followed by two-dimensional thin-layer electrophoresis of the labeled phosphoamino acid (see Basic Protocol). Phosphothreonine and phosphotyrosine are more stable to hydrolysis in alkali than are RNA and phosphoserine. Therefore, mild alkaline hydrolysis of protein samples can be used to enhance the detection of phosphothreonine and phosphotyrosine (see Alternate Protocol). Although this procedure can be carried out with a protein eluted from a preparative gel and concentrated by trichloroacetic acid (TCA) or acetone precipitation, it is most easily accomplished by transfer of the protein of interest to a PVDF membrane. This technique is obviously not ideal if the protein being studied does not transfer efficiently. NOTE: Wear gloves and use blunt-end forceps to handle membranes. CAUTION: When working with radioactivity, take appropriate precautions to avoid contamination of the experimenter and the surroundings. Carry out the experiment and dispose of wastes in an appropriately designated area, following the guidelines provided by the local radiation safety department (also see APPENDIX 1D). ACID HYDROLYSIS AND TWO-DIMENSIONAL ELECTROPHORETIC ANALYSIS OF PHOSPHOAMINO ACIDS

BASIC PROTOCOL

The protein to be acid hydrolyzed is transferred to a PVDF membrane using the same technique used for immunoblotting (UNIT 6.2) or for microsequencing. It is valuable, but not absolutely essential, to keep the filter wet following transfer. Following acid hydrolysis, phosphoamino acids are separated by two-dimensional thin-layer electrophoresis. Because electrophoresis equipment differs considerably in design, the details of the assembly and placement of the plate are not discussed here. It is assumed that a suitable apparatus is available for use by an experienced operator. Electrophoresis conditions are described for using the HTLE 7000 (CBS Scientific). They are almost certainly not correct for other equipment and will need to be altered according to the equipment manufacturer’s directions. Materials 32 P-labeled phosphoprotein (UNIT 14.4) India ink solution: 1 µl/ml India ink in TBS (UNIT 14.4)/0.02% (v/v) Tween 20, pH 6.5 (prepare fresh or store indefinitely at room temperature); or radioactive or phosphorescent alignment markers 6 M HCl Phosphoamino acid standards mixture (see recipe) pH 1.9 electrophoresis buffer (see recipe) pH 3.5 electrophoresis buffer (see recipe) 0.25% (w/v) ninhydrin in acetone in a freon (aerosol, gas-driven) atomizer/sprayer PVDF membrane (Immobilon-P, Millipore) 110° oven Screw-cap microcentrifuge tubes 20 cm × 20 cm × 100 µm glass-backed cellulose thin-layer chromatography plate (EM Sciences) Large blotter: two 25 × 25–cm layers of Whatman 3MM paper sewn together at the edges, with four 2-cm holes that align with the origins on the TLC plate Contributed by Bartholomew M. Sefton Current Protocols in Cell Biology (1999) 14.5.1-14.5.8 Copyright © 1999 by John Wiley & Sons, Inc.

Signal Transduction: Protein Phosphorylation

14.5.1 Supplement 3

Glass tray or plastic box Whatman 3MM paper Thin-layer electrophoresis apparatus (e.g., HTLE 7000, CBS Scientific) Fan Small blotters: 4 × 25–cm, 5 × 25–cm, and 10 × 25–cm pieces of Whatman 3MM paper 50° to 80°C drying oven Sheets of transparency film for overhead projector Additional reagents and equipment for SDS-PAGE (UNIT 6.1), immunoblotting (UNIT 6.2), and autoradiography (UNIT 6.3) Prepare sample 1. Run radiolabeled phosphoprotein on a preparative SDS-polyacrylamide gel (UNIT 6.1). It is difficult to obtain good results with 0.5 ml water. Place the piece of filter paper in a screw-cap microcentrifuge tube. Keep the excised piece as small as possible.

Hydrolyze sample 5. Add enough 6 M HCl to submerge the piece of filter. Screw the cap on the tube tightly and incubate 60 min in 110°C oven. 6. Let cool. Microcentrifuge 2 min at maximum speed, room temperature. Transfer the liquid hydrolysate to a fresh microcentrifuge tube and dry with a Speedvac evaporator. Drying takes ∼2 hr. Simultaneous drying of the hydrolysate and deblocked oligonucleotides in NH4OH must be avoided, as this will generate a cloud of ammonium chloride that will collect in the centrifuge tube and render the hydrolysate unsuitable for thin-layer electrophoresis.

7. Dissolve the sample in 6 to 10 µl water by vortexing vigorously. Microcentrifuge 5 min at maximum speed. Prepare plate for first-dimension electrophoresis 8. Spot 25% to 50% of the sample, in 0.25- to 0.50-µl aliquots, on one origin of a 20 cm × 20 cm × 100 µm glass-backed cellulose thin-layer chromatography plate (see Fig. 14.5.1 for arrangement of samples). Between each application, dry the sample spot with compressed air delivered through a Pasteur pipet plugged with cotton. Use long, thin plastic micropipet loading tips for loading, and do not let the tip touch the plate.


Four samples can be analyzed simultaneously. The complete hydrolysate can be spotted on a single origin, but some streaking in the first dimension may be observed due to overloading. This problem can be avoided by using a fraction of the sample.

14.5.2 Supplement 3


A

B

25 cm 8 cm

3 cm 25 cm

8 cm 3 cm plate with samples applied at 4 origins (+)

C

first-dimension blotter for electrophoresis at pH 1.9

D

phosphoamino acids blotter applied to plate in order to wet it

results of first-dimension electrophoresis at pH 1.9

Figure 14.5.1 First-dimension electrophoretic separation of phosphoamino acids at pH 1.9. (A) Positions of the four origins on a single 20 × 20–cm plate; (B) blotter used for wetting the plate with pH 1.9 electrophoresis buffer; (C) placement of the blotter on the plate (underneath; indicated by dashed outline); and (D) orientation of the plate between the + and − electrodes with the positions of the phosphoamino acids after electrophoresis.

9. Spot 1 µl nonradioactive phosphoamino acid standards mixture (containing phosphoserine, phosphothreonine, and phosphotyrosine) on top of each sample in 0.25to 0.50-µl aliquots as above. 10. Wet the large blotter (with four holes) by submerging it in pH 1.9 electrophoresis buffer in a large glass tray or plastic box. Briefly allow the excess buffer to drain off. Lower the wet blotter onto the prespotted plate with the origins on the plate in the centers of the four holes in the blotter (Fig. 14.5.1). Press on the blotter gently to achieve even wetting of the cellulose and concentration of the samples. When the plate is uniformly wet, remove the blotter. The blotter should be quite damp but not sopping wet. Excess buffer can be wicked off onto filter paper. Areas of the plate that are too dry can be seen through the blotter and will appear to be whiter than the rest of the plate. If this happens, dab the blotter with a Kimwipe wetted with pH 1.9 electrophoresis buffer. If there are puddles of buffer on the plate, let them dry before carrying out electrophoresis.



Supplement 3

A

B 10 cm

5 cm

4 cm 25 cm second-dimension blotters for electrophoresis at pH 3.5

C

blotters applied to plate in order to wet it

D

phosphamino acids

90o counterclockwise rotation of plate

results of second-dimension electrophoresis at pH 3.5 showing ninhydrin-stained standards

Figure 14.5.2 Second-dimension electrophoretic separation of phosphoamino acids at pH 3.5. (A) The three pieces of Whatman 3MM paper used for wetting the plate with pH 3.5 electrophoresis buffer; (B) proper placement of the blotters on the plate (underneath; indicated by dashed outline); (C) reorientation of the plate for electrophoresis in the second dimension; and (D) orientation of the plate between the + and - electrodes with the position of the phosphoamino acids after electrophoresis.

11. Place the thin-layer plate in the electrophoresis apparatus and overlap 0.5 cm of the right and left sides of the plate with wicks made of Whatman 3MM paper. If the apparatus has an air bag, be sure to inflate it. Close the cover and start electrophoresis. With an HTLE 7000, double-thickness Whatman 3MM wicks, and a plate with four samples, electrophorese 20 min at 1.5 kV. For the HTLE 7000 apparatus, use folded-over Whatman 3MM wicks that are 20 cm wide (the same as the plate) and not overly wet. Overly wet wicks will flood the plate and cause sample diffusion.


For other electrophoresis apparatuses the appropriate duration of electrophoresis can be determined empirically by examining the rate of migration of the phosphoamino acid standards.

14.5.4 Supplement 3


Pi

phosphoserine phosphothreonine phosphotyrosine phosphopeptides generated by partial hydrolysis origin

pH 3.5

pH 1.9

Figure 14.5.3 Hypothetical autoradiogram of a two-dimensional separation. Four samples of acid-hydrolyzed, 32P-labeled proteins are applied at the origins, one in each of the four quandrants. This diagram shows the origins, the directions of electrophoresis, the positions of phosphoserine, phosphothreonine, and phosphotyrosine, the position of Pi, and the position of partially hydrolyzed fragments of the proteins for the upper right-hand sample. Every protein generates different partial hydrolysis peptide fragments.

12. Following electrophoresis, remove the plate and quickly air dry with a fan without heating. It takes ∼20 min to dry the plate.

Perform second-dimension electrophoresis 13. Wet the small blotters in pH 3.5 electrophoresis buffer and use them to wet the plate using the method described in step 10 to achieve even wetting without puddling (Fig. 14.5.2). After electrophoresis at pH 1.9, phosphoamino acids are present as a streak extending from the origin towards the + electrode. Blotters are not applied directly over the phosphoamino acids to prevent sample blurring or smearing.

14. Remove the blotters, rotate the plate 90° counterclockwise, and electrophorese 16 min at 1.3 kV in pH 3.5 electrophoresis buffer if using the HTLE 7000 apparatus. 15. At the end of the electrophoresis run, remove the plate and dry 20 to 30 min in an oven at 50° to 80°C. When dry, spray with 0.25% ninhydrin in acetone, then reheat in the oven 5 to 10 min to visualize the phosphoamino acid standards. 16. Place radioactive or phosphorescent alignment marks on the plate and autoradiograph with an intensifying screen overnight to 10 days at −70°C. 17. Following autoradiography, trace the alignment markers and the stained phosphoamino acid markers onto a transparent sheet used for overhead projectors. Save this template. Align the film with the plate and identify radioactive phosphoamino acids (Fig. 14.5.3). Use of fluorography or autoradiography to detect the labeled phosphoamino acids is preferable to use of a phosphorimager. The image on film is precisely the same size as the thin-layer plate, which allows the transparent film to be overlaid on the plate for an unambiguous spot identification. A phosphorimager can subsequently be used for quantification.



Supplement 3

ALTERNATE PROTOCOL

ALKALI TREATMENT TO ENHANCE DETECTION OF TYR- AND THR-PHOSPHORYLATED PROTEINS BLOTTED ONTO FILTERS Phosphothreonine and phosphotyrosine are much more stable to hydrolysis in alkali than RNA or phosphoserine. Detection of proteins containing phosphothreonine and phosphotyrosine in impure samples containing 32P-labeled RNA and serine-phosphorylated proteins can often be enhanced by mild alkaline hydrolysis of gel-fractionated samples. Although this technique was first developed for the treatment of fixed polyacrylamide gels, it is much more easily performed with proteins that have been first transferred to a PVDF membrane. Alkaline hydrolysis does not preclude subsequent phosphoamino acid analysis. A band from a blot that has been treated with alkali can be excised and subjected to acid hydrolysis as described in the Basic Protocol. Additional Materials (also see Basic Protocol) 1 M KOH TN buffer: 10 mM Tris⋅Cl (pH 7.4 at room temperature)/0.15 M NaCl 1 M Tris⋅Cl, pH 7.0 at room temperature Covered plastic container (e.g., Tupperware box) 55°C oven or water bath 1. Run radiolabeled sample on a preparative SDS–polyacrylamide gel and transfer proteins electrophoretically to a PVDF membrane (see Basic Protocol steps 1 and 2). A band containing as few as 10 cpm is detectable under optimal conditions with this technique. A nylon membrane may be used in place of a PVDF membrane, but in that case, the bands cannot subsequently be analyzed by acid hydrolysis, as nylon membrane will dissolve in 6 M HCl.

2. Wash membrane thoroughly with water: three 2-min incubations in 1 liter water are sufficient. These washes remove buffer and detergent.

3. Incubate membrane 120 min at 55°C in an oven or water bath in sufficient 1 M KOH to cover the filter in a covered Tupperware container. 4. Discard KOH. Wash membrane and neutralize remaining KOH by rinsing once for 5 min in 500 ml TN buffer, once for 5 min in 500 ml of 1 M Tris⋅Cl (pH 7.0), and twice for 5 min in 500 ml water. Wrap the membrane in plastic wrap and autoradiograph (UNIT 6.3) overnight with flashed film and an intensifying screen at −70°C. Identification of the band of interest is most easily accomplished by coelectrophoresis of a radioactive marker protein of known identity.



pH 1.9 electrophoresis buffer 50 ml 88% formic acid (0.58 M final) 156 ml glacial acetic acid (1.36 M final) 1794 ml H2O Store indefinitely in a sealed bottle at room temperature

14.5.6 Supplement 3


pH 3.5 electrophoresis buffer 100 ml glacial acetic acid (0.87 M final) 10 ml pyridine [0.5% (v/v) final] 10 ml 100 mM EDTA (0.5 mM final) 1880 ml H2O Store indefinitely in a sealed bottle at room temperature Phosphoamino acid standards mixture Prepare a solution of phosphoserine, phosphothreonine, and phosphotyrosine (Sigma) in water at a final concentration of 0.3 mg/ml each. Store in 1-ml aliquots indefinitely at −20°C. COMMENTARY Background Information Phosphoamino acid analysis by the two-dimensional electrophoretic technique described in the basic protocol was first carried out with proteins isolated by elution from unfixed SDSpolyacrylamide gels (Hunter and Sefton, 1980). However, this technique is laborious, especially if it involves grinding up pieces of high-percentage acrylamide gels, and the yields can be disappointing. Additionally, because the eluted protein must be precipitated in the presence of a carrier protein, spotting the whole sample on a single origin usually yields a badly smeared pattern. The grind-and-elute technique is, however, advantageous with proteins that are very refractory to electrophoretic transfer to PVDF membranes. The alkaline treatment of protein described in the alternate protocol was first developed by Jon Cooper and Tony Hunter, who treated fixed gels with KOH (Cooper and Hunter, 1981). The original technique is tricky because the gel becomes extremely sticky during incubation with KOH and swells. Additionally, the manipulations needed to recover proteins from the gel following treatment are very involved because the proteins are contaminated with products of the hydrolysis of polyacrylamide.

Critical Parameters and Troubleshooting It is essential to use PVDF membranes to immobilize proteins for acid hydrolysis rather than nylon or nitrocellulose membranes, both of which dissolve in 6 M HCl. Proteins immobilized on either PVDF or nylon membranes may be subjected to alkaline hydrolysis with KOH (Contor et al., 1987), but nitrocellulose membranes are not suitable. Proteins immobilized on nylon cannot subsequently be analyzed by acid hydrolysis because nylon is dissolved by 6 M HCl.

Two-dimensional thin-layer electrophoresis is required for unambiguous identification of phosphorylated residues, as some spots after one-dimensional electrophoresis do not represent pure species. For example, uridine monophosphate, which is generated during acid hydrolysis of RNA (a frequent contaminant of phosphoproteins), comigrates with phosphotyrosine during one-dimensional electrophoresis at pH 3.5. Streaking of the sample in the first dimension is a symptom of overloading, either with the phosphoprotein itself or with contaminants in the sample. This problem can be corrected by loading less sample. Streaking in the second dimension is usually the result of problems with wetting or running the plate and cannot be corrected by loading less sample. Some batches of blotting paper contain calcium, which interferes with electrophoresis of phosphoamino acids (probably by precipitating them). In the author’s experience Whatman 3MM paper is quite reliable; other blotting papers are probably suitable as well. Inclusion of EDTA in the pH 3.5 buffer alleviates this problem. This unit calls for glass-backed cellulose thin-layer plates rather than the plastic-backed variety, which are lighter and less expensive. This is because plastic-backed plates can under some circumstances cause sample streaking. They are, however, probably satisfactory for most experiments. If use of plastic-backed plates results in streaking, try glass-backed plates to see if that corrects the problem.

Anticipated Results To detect a phosphoamino acid by autoradiography, a minimum of 10 cpm must be spotted and the plate exposed for a week with flashed film and an intensifying screen. Only 15% to 20% of the radioactivity in a phospho-



Supplement 3

protein is recovered as phosphoamino acids. The majority is present as 32Pi, which is released by dephosphorylation of phosphoamino acids, with the remainder being peptide products resulting from partial acid hydrolysis. As a result, the thin-layer plates will contain a number of radioactive spots that are not phosphoamino acids (see Fig. 14.5.3). Partial hydrolysis products remain near the origin during electrophoresis at pH 1.9, but exhibit some mobility at pH 3.5 (see Fig. 14.5.3). After two-dimensional electrophoresis, they are found above the origin and below the phosphoamino acids. 32Pi has a high mobility at both pH 1.9 and pH 3.5 and is found in the upper left-hand corner of each quadrant of the plate (see Fig. 14.5.3). Because of these additional radioactive spots, it is essential to localize internal phosphoamino acid standards by staining with ninhydrin.

Time Considerations After the preparative gel has been run and the protein transferred to the membrane, isolation of the membrane fragment containing the protein, followed by acid hydrolysis, takes 5 × 105 cells/minigel lane); (4) following the proper techniques for SDS-PAGE and immunoblotting (see UNITS 6.1 & 6.2). Accurate comparison and quantification by densitometry of the amounts of phosphorylated Akt in different samples can be achieved if the detection system is kept in a linear range. The enhanced chemiluminescence (ECL) system should be optimized to obtain linearity by adjustments of the amount of the protein loaded on the gel, concentrations of primary and secondary antibody, and the X-ray film exposure time. If the weakest signal is detectable and the strongest signal is still within the linear range of the film (e.g., not saturated), then the rest of the samples are also in the linear range of the system, which can be used for quantification.

Anticipated Results Typical results expected after probing for Akt activity (see Basic Protocol) or Akt membrane translocation (see Support Protocol) are presented in Figure 14.6.2A and B, respectively. Comparison between the starved cells (contr) and cells after PDGF stimulation (PDGF) should demonstrate a several-fold increase in the amount of phosphorylated Akt. Attenuation of this response after experimental manipulations of the cell cultures may indicate effects on the upstream pathways leading to Akt activation. Evaluation of the amount of phosphorylated Akt in the samples taken after PDGF

stimulation at different time points provides information about the rate of Akt dephosphorylation (inhibition). While in the samples from β1 cells, this decrease is modest, the Akt in the samples from mutant cells is rapidly dephosphorylated. Such a result indicates activation of some of the systems for Akt inhibition (see Background Information). Performing the same experiment in the presence of a PP2A inhibitor, okadaic acid completely reverses this effect, indicating that activation of this phosphatase is involved in the observed increased dephosphorylation rate in W/A mutant cells. If OA is ineffective, additional experiments should be designed to test the role of other Akt inhibitors (see Background Information). Activation of Akt is dependent on its membrane translocation. A typical increase in membranebound Akt after growth factor stimulation is presented in Figure 14.6.2B. A modest but detectable increase in total Akt in the membrane fraction is observed in both cell lines after PDGF stimulation. Failure to detect such translocation may indicate defects in the function of the PH domain of Akt or insufficient phosphatidylinositol 3,4,5-trisphosphate and phosphatidylinositol 3,4-bisphosphate (see Background Information).

Time Considerations The entire procedure described in the Basic Protocol can be completed in 3 days. This period includes the time for cell attachment after plating (4 hr); starvation (12 hr); PDGF stimulation and the necessary incubations up to preparation of the SDS-PAGE samples (3 hr); SDS-PAGE and electrotransfer (6.5 hr for mini gels or 9.5 hr for normal size gels); overnight incubation with the primary antibody; and completion of the immunoreactions with ECL processing (4 hr). Since this procedure takes >1 day, it is helpful to use one night for starvation of the cells and the next night for incubation with the primary phosphospecific antibody. There are a number of points where the procedure can be interrupted: (1) after the preparation of the SDS-PAGE samples; (2) after the electrotransfer (membranes can be stored wet or dry in resealable plastic bags at 4°C); and (3) after the completion of the first immunoreaction (membranes can be stored wet in resealable plastic bags at 4°C). A similar timeframe applies for the procedure described in the Support Protocol. Signal Transduction: Protein Phosphorylation


Supplement 22

Literature Cited Andjelkovic, M., Alessi, D.R., Meier, R., Fernandez, A., Lamb, N.J.C., Frech, M., Cron, P., Cohen, P., Lucoc, J.M., and Hemmings, B.A. 1997. Role of translocation in the activation and function of protein kinase B. J. Biol. Chem. 272:31515-31524. Brazil, D.P. and Hemmings, B.A. 2001. Ten years of protein kinase B signalling: A hard Akt to follow. Trends Biochem. Sci. 26:657-664. Chan, T.O., Rittenhouse, S.E., and Tsichlis, P.N. 1999. AKT/PKB and other D3 phosphoinositide-regulated kinases: Kinase activation by phosphoinositide-dependent phosphorylation. Annu. Rev. Biochem. 68:965-1014. Datta, S.R., Brunet, A., and Greenberg, M.E. 1999. Cellular survival: A play in three Akts. Genes Dev. 13:2905-2927. Fassler, R., Pfaff, M., Murphy, J., Noegel, A., Johansson, S., Timpl, R., and Albrecht, R. 1995. Lack of beta 1 integrin gene in embryonic stem cells affects morphology, adhesion, and migration but not integration into the inner cell mass of blastocysts. J. Cell Biol. 128:979-988.

Marte, B.M. and Downward, J. 1997. PKB/Akt: Connecting phosphoinositide 3-kinase to cell survival and beyond. Trends Biochem. Sci. 22:355-358 Meier, R. and Hemmings, B.A. 1999. Regulation of protein kinase B. J. Recept. Signal Transduct. Res. 19:121-128. Millward, T.A., Zolnierowicz, S., and Hemmings, B.A. 1999. Regulation of protein kinase cascades by protein phosphatase 2A. Trends Biochem. Sci. 24:186-191. Pankov, R., Cukierman, E., Clark, K., Matsumoto, K., Hahn, C., Poulin, B., and Yamada, K.M. 2003. Specific beta 1 integrin site selectively regulates Akt/protein kinase B signaling via local activation of protein phosphatase 2A. J. Biol. Chem. 278:18671-18681. Persad, S. and Dedhar, S. 2003. The role of integrinlinked kinase (ILK) in cancer progression. Cancer Metastasis Rev. 22:375-384. Scheid, M.P. and Woodgett, J.R. 2003. Unravelling the activation mechanisms of protein kinase B/Akt. FEBS Lett. 546:108-112.

Franke, T.F., Kaplan, D.R., and Cantley, L.C. 1997. PI3K: Downstream AKT ion blocks apoptosis. Cell 88:435-437.

Testa, J.R. and Bellacosa, A. 1997. Membrane translocation and activation of the Akt kinase in growth factor-stimulated hematopoietic cells. Leuk. Res. 21:1027-1031.

Hemmings, B.A. 1997. Akt signaling: Linking membrane events to life and death decisions. Science 275:628-603.

Vanhaesebroeck, B. and Alessi, D.R. 2000. The PI3K-PDK1 connection: More than just a road to PKB. Biochem. J. 346:561-576.

Hill, M.M. and Hemmings, B.A. 2002. Inhibition of protein kinase B/Akt. Implications for cancer therapy. Pharmacol. Ther. 93:243-251.

Yamada, K.M. and Araki, M. 2001. Tumor suppressor PTEN: Modulator of cell signaling, growth, migration and apoptosis. J. Cell Sci. 114:23752382.

Kobayashi, S., Shirai, T., Kiyokawa, E., Mochizuki, N., Matsuda, M., and Fukui, Y. 2001. Membrane recruitment of DOCK180 by binding to PtdIns(3,4,5)P3. Biochem. J. 354:73-78. Maira, S.M., Galetic, I., Brazil, D.P., Kaech, S., Ingley, E., Thelen, M., and Hemmings, B.A. 2001. Carboxyl-terminal modulator protein (CTMP), a negative regulator of PKB/Akt and v-Akt at the plasma membrane. Science 294:374-380.

Contributed by Roumen Pankov National Institute of Dental and Craniofacial Research National Institutes of Health Bethesda, Maryland

Determination of Akt/PKB Signaling



CHAPTER 15 Protein Trafficking INTRODUCTION

A

s discussed in the introduction to Chapter 3, eukaryotic cells are not simply sacs of amorphous protoplasm but are instead organized into an array of membrane-bound structures known as organelles. Subcellular fractionation (Chapter 3) and microscopic analyses (Chapter 4) have revealed a wondrous diversity of subcellular organelles, including the nucleus, endoplasmic reticulum (ER), Golgi complex, lysosomes, endosomes, secretory granules, plasma membrane, mitochondria, peroxisomes, chloroplasts, and a variety of other structures found in specialized cells. Each of these organelles fulfills a distinct function, which is carried out by a specific set of organellar proteins. Genetic information directing the synthesis of most organellar proteins, as well as cytosolic and extracellular proteins, is encoded by DNA contained within the nucleus. Genomic DNA is transcribed into messenger RNA (see UNIT 11.6) and this in turn is translated into protein (see UNIT 11.1). It is at the time of their translation that proteins begin a journey that will take them to different locations within the cell, or out to the extracellular space. Some proteins are synthesized on ribosomes that exist free in the cytosol. These proteins either remain soluble in the cytosol, become incorporated into supramolecular structures such as the cytoskeleton, or are imported into the nucleus, mitochondria, peroxisomes, chloroplasts, and in some cases the ER. Other proteins are synthesized on ribosomes that are associated with the cytosolic face of the ER membrane. These proteins are translocated across that membrane and subsequently distributed to different compartments of the secretory and endocytic pathways including the ER proper, the Golgi complex, lysosomes, endosomes, secretory granules, and the plasma membrane; in other cases they are secreted into the extracellular medium. Thus, the interior of the cell is a highly dynamic environment, in which heterogeneous yet precisely controlled protein distributions are achieved via various protein trafficking pathways. In vitro assays to examine specific protein transport steps and their molecular mechanisms have been described in Chapter 11. However, in vitro assays are valid only if they reflect the transport processes that occur in intact cells. In addition, the itineraries followed by proteins as they traffic through different cellular compartments are often too complex to be reconstituted in vitro and are best studied in intact cells. Chapter 15 is devoted to biochemical and morphologic assays used to study protein trafficking in vivo. The chapter begins with an overview of protein trafficking in the secretory and endocytic pathways (UNIT 15.1), which provides useful background information for some of the succeeding protocol units. Most proteins destined for organelles of the secretory or endocytic pathways, or for secretion into the extracellular milieu, are co-translationally modified by addition of carbohydrate moities. These moieties are progressively remodeled as the proteins traverse the different compartments of the secretory pathway. The type of carbohydrate present on a protein thus serves as a record of its itinerary through the cell. Gross analyses of the carbohydrate moities of glycoproteins can be conveniently carried out by treatment of metabolically labeled, immunoprecipitated proteins (see UNITS 7.1 & 7.2) with an assortment of specific glycosidases. UNIT 15.2 presents a comprehensive description of carbohydrate modifications along the secretory pathway, and provides a series of protocols for analysis of the general structure of carbohydrate moieties of glycoproteins labeled in vivo. Contributed by Juan S. Bonifacino Current Protocols in Cell Biology (2004) 15.0.1-15.0.3 Copyright © 2004 by John Wiley & Sons, Inc.

Protein Trafficking


Even after proteins reach their allotted destinations within the cells, they remain in constant flux. Indeed, most proteins are not fixed at a particular location but attain a particular steady-state distribution as a result of opposing exit and retrieval pathways. This is very neatly exemplified by some plasma membrane proteins, such as endocytic receptors, that are rapidly internalized but return to the cell surface after a sojourn in the endosomal system. UNIT 15.3 describes a set of biochemical techniques for measuring the steady-state distribution, internalization, and recycling of the transferrin receptor, a typical endocytic receptor, using radioiodinated transferrin as a probe. Protocols are also provided to measure the steady-state distribution, internalization, and recycling of any plasma membrane protein, even those that do not have a physiologic ligand, by using radioiodinated antibodies or Fab fragments. Internalized ligands and antibodies used to tag plasma membrane proteins are often degraded in lysosomes, a process that can be measured using another protocol included in this unit. Finally, the unit also covers methods for measuring fluid-phase uptake from the medium and for inhibiting endocytosis. Integral membrane proteins expressed at the cell surface are synthesized in the ER and transported to the plasma membrane following the secretory pathway. By the time that these proteins reach the cell surface, almost all post-translational modifications have taken place. For this reason, the carbohydrate analyses described in UNIT 15.2 cannot be used to determine the kinetics of protein arrival at the cell surface. However, the accessibility of cell surface proteins to externally added reagents has been exploited to develop assays for biosynthetic protein transport to the plasma membrane. UNIT 15.4 contains several protocols that combine pulse-chase metabolic labeling (see UNIT 7.1) with treatments that modify surface-exposed proteins on intact cells: hydrolysis of sialic acid residues with sialidase, biotinylation, protease digestion, and antibody binding. Modified surface-exposed proteins are detected by electrophoretic or immunoprecipitation techniques sensitive to the modifications introduced. Although one often refers to the plasma membrane as if it were a single compartment, most eukaryotic cells exhibit more than one plasma membrane domain. Polarized epithelial cells are perhaps the best characterized example of cells that have specialized plasma membrane domains. In epithelial cells, these domains are known as apical and basolateral. Protein trafficking to and from these two domains follows distinct pathways, the analysis of which is the focus of UNIT 15.5. For analysis of polarized sorting to be possible, these cells need to be grown in special chambers fitted with porous filters. The cells form a tight monolayer in which the apical and basolateral domains are segregated. In the chambers the apical and basolateral media are separately available for addition of reagents or collection of secreted proteins. The unit contains protocols for metabolic labeling of polarized epithelial cells and collection of proteins secreted from the apical and basolateral surfaces, stable transfection and selection of transfected clones, culture of epithelial cells on filters, determination of leakiness of epithelial monolayers, monitoring the arrival of newly synthesized proteins at both plasma membrane domains, and indirect immunofluorescence microscopy of epithelial cells grown as polarized monolayers.

Introduction

UNIT 15.6 takes us back to early events in the maturation of newly synthesized proteins in the ER. Upon emergence into the lumen of the ER, nascent polypeptide chains undergo a series of post-translational modifications that lead to the development of a mature protein (see UNIT 15.1). Among these modifications are glycosylation (UNIT 15.2), folding, and disulfide bond formation. UNIT 15.6 describes a series of protocols for the analysis of protein folding and disulfide bond formation in the ER. In these protocols, newly synthesized proteins are first labeled with radiolabeled amino acids either in intact cells (also see UNIT 7.1) or in cell-free (also see UNIT 11.4) or semi-permeabilized cell systems.



Protein folding can then be followed by limited proteolytic digestion or immunoprecipitation with conformation-specific antibodies. Disulfide bond formation can be analyzed by monitoring the disappearance of sulfhydryl groups or the appearance of faster-moving species on nonreducing SDS-PAGE (also see UNIT 6.5). The subject of UNIT 15.7 is phagocytosis—the process by which specialized cells such as macrophages and monocytes ingest particles such as bacteria, yeast, and apoptotic bodies. Because of the size and complexity of the internalized particles, phagocytosis is distinct from the receptor-mediated or fluid-phase endocytosis described in UNIT 15.3. The first step in phagocytosis is the recognition of specific molecules on the surface of the particle by the phagocytic cell. These molecules can be intrinsic to the particle or deposited onto the particle by the host (i.e., “opsonization”). The unit starts with microscope-based protocols to measure phagocytosis of red blood cells or latex beads opsonized with immunoglobulin G (IgG), which are internalized upon binding to Fcγ receptors on the phagocytic cells. These protocols are followed by others designed to measure complement-mediated phagocytosis of red blood cells or latex beads opsonized with complement factor C3bi, a process that is mediated by binding to complement receptors. Phagocytosis can be measured more quantitatively by flow cytometry, for which a protocol is also included in this unit. The next protocols deal with events that follow the uptake of particles, such as the acidification and maturation of the phagosome. The final protocol describes a procedure to inhibit phagocytosis with wortmannin or cytochalasin D. Juan S. Bonifacino

Protein Trafficking


Supplement 22

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Figure 17.1.1 (A) Cy3-PY72 photobleaching releases FRET-quenched EGFR-GFP emission. The histogram shows the distribution of calculated FRET efficiencies in the cell. (B) FRET measured by fluorescence lifetime imaging microscopy. The histogram shows the distribution of measured lifetime (nsec) and corresponding calculated FRET efficiencies prior and after photobleaching of the acceptor. See color figure.

the photobleached area. The heterogeneity in this distribution contains information about variability in the formation of complexes of the proteins carrying the donor and acceptor fluorophores. Figure 17.1.1 shows an MCF-7 mammary carcinoma cell expressing EGFR-GFP that was stimulated with 100 ng/ml EGF for 5 min. This cell was incubated with Cy3-labeled anti-phos-

photyrosine antibodies (e.g., 10 µg/ml PY72 monoclonal) and EGFR-GFP tyrosine phosphorylation was measured by FRET using the unquenching of GFP fluorescence by acceptor photobleaching (Fig. 17.1.1A) and FLIM (Fig. 17.1.1B). The EGFR-GFP distribution is shown in the D1 image. Most EGFR-GFP is localized at the plasma membrane in addition to internalized (punctate) EGFR-GFP. The cor-

Molecular Interactions in Cells


Supplement 18

responding antibody-staining A1 shows phosphorylation at the plasma membrane. A rectangular region was subjected to acceptor photobleaching (white box). The GFP fluorescence in this region increased after photobleaching as shown in the difference image (D2 − D1), demonstrating positive FRET efficiencies [(D2 − D1)/D2]. Highest FRET efficiencies can be observed in the plasma membrane corresponding to fully phosphorylated receptor at this location. The large peak in the energy-transfer efficiency histogram that is distributed around zero originates from the area outside the photobleached region and indicates proper image registration. The additional population, ranging from 15% to 35% efficiency, corresponds to the photobleached region. Figure 17.1.1B shows the analogous result of a fluorescence lifetime measurement. The steady-state fluorescence distribution of EGFR-GFP is shown in (D) and the corresponding lifetime map in (τ1). The anti-phosphotyrosine Cy3 immunofluorescence (A) is photobleached to obtain an intracellular reference lifetime in absence of acceptor (τ2). As can be seen in the fluorescencelif etim e h istog ram, these values are homogeneously distributed around an average of ∼2.0 nsec. The decrease in lifetime due to FRET is shown in the difference image (τ2 − τ1). The energy-transfer efficiency is given by the normalization of this difference to the reference EGFR-GFP lifetime [E = (τ2 − τ1)/τ2] and again shows highly phosphorylated receptor in the plasma membrane.

Time Considerations

Imaging Protein-Protein Interactions by FRET Microscopy

From cell seeding, the entire procedure can be performed in 3 to 4 days. This period includes 1 to 2 days for expression of the GFP construct after transfection or microinjection; 1/2 day for treatment, fixation, and antibody incubation of cells; 1/2 day of data acquisition; and 1/2 day for data analysis. Removal of gelatin/BSA from commercial antibody takes ∼3 hr. The most time-consuming step in the procedure for labeling antibodies is the buffer exchange using a Microcon concentration device. Depending on the amount of contaminating free amino groups in the original buffer, this can take between 2 and 5 hr. The labeling reaction and subsequent gel filtration chromatography takes ∼1 hr. There are a number of points where the procedure can be interrupted: (1) optimal expression after overnight incubation rather than a 3- to 4-hr incubation post microinjection can

be achieved by lowering the concentration of DNA; (2) after fixation and permeabilization, the cells can be stored overnight in PBS at 4°C before antibody incubation; (3) after mounting, the cells can be viewed immediately rather than the following day, by application of rubber cement instead of Mowiol; and (4) the samples can be stored at −20°C for weeks without appreciable loss of antibody staining or FRET.

Literature Cited Adams, S.R., Harootunian, A.T., Buechler, J., Taylor S.S., and Tsien, R.Y. 1991. Fluorescence ratio imaging of cyclic AMP in single cells. Nature 349:694-697. Bastiaens, P.I.H. and Jovin, T.M. 1996. Microspectroscopic imaging tracks the intracellular processing of a signal-transduction protein: Fluorescent labeled protein kinase C beta I. Proc. Natl. Acad. Sci. U.S.A. 93:8407-8412. Bastiaens, P.I.H. and Jovin, T.M. 1998. Fluorescence resonance energy transfer microscopy. In Cell Biology a Laboratory Handbook, Vol. 3 (J.E. Celis, ed.), pp. 136-146. Academic Press, New York. Bastiaens, P.I.H. and Squire, A. 1999. Fluorescence lifetime imaging microscopy: Spatial resolution of biochemical processes in the cell. Trends Cell Biol. 9:48-52. Bastiaens, P.I.H., Majoul, I.V., Verveer, P.J., Soling, H.D., and Jovin, T.M. 1996. Imaging the intracellular trafficking and state of the AB(5) quaternary structure of cholera-toxin. EMBO J. 15:4246-4253. Clegg, R.M. 1996. Fluorescence resonance energy transfer spectroscopy and microscopy. In Fluorescence Imaging Spectroscopy and Microscopy (X.F. Wang and B. Herman eds.) pp. 179-251. John Wiley & Sons, New York. Day, R.N. 1998. Visualization of Pit-1 transcription factor interactions in the living cell nucleus by fluorescence resonance energy transfer microscopy. Mol. Endocrinol. 12:1410-1419. Förster, T. 1948. Zwischenmolekulare Energiewanderung und Fluoreszenz. Ann. Phys. 2:55-75. Gadella, T.W.J. and Jovin, T.M. 1995. Oligomerization of epidermal growth-factor receptors on A431 cells studied by time-resolved fluorescence imaging microscopy—a stereochemical model for tyrosine kinase receptor activation. J. Cell Biol. 129:1543-1558. Gadella, T.W.J., Jovin, T.M., and Clegg, R.M. 1993. Fluorescence lifetime imaging microscopy (FLIM)—spatial resolution of microstructures on the nanosecond time-scale. Biophys. Chem. 48:221-239. Gordon, G.W., Berry, G., Huan Liang, X., Levine, B., and Herman, B. 1998. Quantitative fluorescence resonance energy transfer measurements using fluorescence microscopy. Biophys. J. 74:2702-2713.



/DNRZLF]-5DQG%HUQGW./LIHWLPHVHOHF WLYHIOXRUHVFHQFHLPDJLQJXVLQJDQUISKDVHVHQ VLWLYHFDPHUD5HY6FL,QVWUXP 0DKDMDQ13/LQGHU.%HUU\**RUGRQ*: +HLP5DQG+HUPDQ%%FODQGED[ LQWHUDFWLRQVLQPLWRFKRQGULDSUREHGZLWKJUHHQ IOXRUHVFHQW SURWHLQ DQG IOXRUHVFHQFH HQHUJ\ WUDQVIHU1DWXUH%LRWHFKQRO 0DW] 09 )UDGNRY $) /DEDV − H[S NRQ F $ + NRII W @ NRII + NRQ F $ Equation 17.6.3

5GLVVRF W = 5DVVRF W G H[S> −N RII W − W G @ Equation 17.6.4

directly to the data (with td denoting the end of the association and the beginning of the dissociation phase, where cA = 0; O’Shannessy et al., 1993). This should be done without truncation of the data beyond possible regions of artifacts from buffer changes or injections. Global analysis should be employed, which allows one to analyze all experimentally observed curves at all analyte concentrations simultaneously. This results in unambiguous best-parameter estimates for the binding constants, and provides residuals that should be inspected for comparison of the model and the data. Software needed for global analysis can be obtained from the manufacturer or as shareware. It should be noted that exclusion of specific data regions that may be poorly described by the model will introduce a bias into the analysis. If the global analysis cannot be performed, and separate analyses of the individual binding curves of each experiment according to Equations 17.6.3 and 17.6.4 are applied, then tests for consistency of the results are crucial (Schuck and Minton, 1996a). Important tests are: (1) the estimated extrapolated values of Rassoc (t → ∞) at different concentrations should be consistent with a Langmuir isotherm for the estimated KD (e.g., approaching half-saturation at cA = KD; see Figure 17.6.2); and (2) the extrapolated values of (kassoccA + koff) from the association phases should be consistent with the value of koff as estimated in the dissociation phase. An analysis of the accuracy of the kinetic rate constants can be found in Ober and Ward (1999a).

Measuring Protein Interactions by Optical Biosensors

Possible problems The data may be only poorly described by Equations 17.6.3 and 17.6.4 (leading to a poor fit and systematically distributed residuals) and instead show multiphasic behavior. Although it is in principle possible to apply global modeling with multiple binding-sites models, this is not recommended, because it is very difficult and can easily lead to very large errors (up to several orders of magnitude). Such modeling is usually not very robust, and many different models may fit the same data equally well (Glaser and Hausdorf, 1996). The reader is also advised to be extremely cautious with some of the implemented models in commercial or shared software, because they may not be valid descriptions of real surface binding processes (for example, mass transport models or models for bivalent analytes). Frequently, the deviations of the data from the single exponential binding



progress predicted in Equations 17.6.3 and 17.6.4 are signatures of experimental artifacts, such as heterogeneity of the ligand (subpopulations of ligand with different binding properties through nonspecific immobilization), steric hindrance of binding to neighboring sites, or the presence of mass transport limitations or multivalent aggregates of the analyte (see discussion of Common Experimental Obstacles). Steady-State Experiments and Binding Isotherm Analysis This approach can yield both equilibrium constants and, if appended by a dissociation analysis, kinetic constants. Its main advantage with respect to the previously described technique (see discussion of Kinetic Analysis) is that it is much more robust and reliable because it is less affected by problems of potential multiphasic binding. Experimental If in the course of the association experiments a steady-state signal is observed, which is generally possible for interactions with relatively high koff, then a binding isotherm can be constructed (Figs. 17.6.3 and 17.6.4). In an experimental procedure similar to the kinetic analysis, a concentration range as wide as possible should be covered, with two-fold concentration steps starting from both far below (≤1/10) KD up to almost complete saturation of the surface sites, far above the KD (≥ 10 times the KD). The injection times required to attain steady state at the lower concentrations are longer (see Equations 17.6.3 and 17.6.4), and exponential extrapolation may be used, in particular in the Biacore system, since the association times may be limited in the by the limited volume in the injection loop of the microfluidics. Exponential extrapolation is not advised, however, for concentrations above KD. For interactions that do not yield steady-state binding within a tolerable time interval for the association phase, or where the ligand does not withstand a regeneration procedure, equilibrium titration represents an experimental variant (Fig. 17.6.4). In this approach, the analyte concentration is increased in two-fold steps, allowing for attainment of steady state in each step, avoiding regeneration between the steps. This can be achieved by addition of a small aliquot of the analyte into a fixed volume of solution above the sensor surface (in cuvette-based sensors; Hall and Winzor, 1997), into a loop of recycling sample (slightly modified Biacore X; Schuck et al., 1998), or into the running buffer reservoir of a standard Biacore (Myszka et al., 1998). The fixed-volume techniques require far less sample volume. After the highest analyte concentration has been applied, the dissociation phase can be observed and analyzed in terms of Equation 17.6.4. This can give an estimate of koff, which can be used in conjunction with Equation 17.6.2 to calculate kon as koff/KD. Data analysis Although the data analysis could in principle be performed by Scatchard analysis, this is not recommended for statistical reasons. A better method is a plot of Rsteady-state versus log(cA), in which the KD can be easily estimated as the inflection point at half saturation of the typical sigmoid isotherm (Fig. 17.6.4). The functional form for modeling is:

5VWHDG\ −VWDWH F$ =

+

5PD[ . ' F$

Equation 17.6.5

Even if the data cannot be fitted well with this expression, the 50% saturation will still represent a good estimate of the order of magnitude of an average KD. It is important, however, in such an approach, to have enough data points at high analyte concentrations

Macromolecular Interactions in Cells


Supplement 22

for a reliable estimate of Rmax and the 50% saturation level, respectively. Then the robustness will be greater than that of the kinetic analysis of Equations 17.6.3 and 17.6.4. Competition Analysis Competition experiments can be performed in different variations, but conceptually all use a calibration of the sensor signal as a function of free analyte concentration, followed by experiments with mixtures of analyte and soluble ligand in different molar ratios. The basic presumption is that the soluble ligand competes with the immobilized ligand for the

A

100 4KD

80

Signal (%)

2KD 10KD

60 KD 0.5KD

40

20

0.25KD cA = 0.1KD

0 0

Steady-state signal (%)

B

200

400

600

800

1000

1200

100

80

60

40

20

0 0.01KD

0.1KD

KD

10KD

100KD

Analyte concentration cA


Figure 17.6.4 (A) Time course of an equilibrium titration experiment. Arrows indicate the timepoints of the step-wise increase in the analyte concentration. (B) Steady-state binding analysis according to Equation 17.6.5. In a plot of the steady-state signal versus the base-10 logarithm of the analyte concentration, the binding isotherm exhibits a typical sigmoid shape. The inflection point at 50% saturation determines KD, and the width of the curve is characterized by ∼10% saturation at 1/10 KD and ∼90% saturation at 10 KD (dotted lines). Visible inspection of the data in this representation already allows a robust parameter estimation.



free analyte, which leads to a reduction of the surface binding. This gives information on the solution affinity of ligand and analyte, while the freedom from the need to explicitly model the surface binding drastically diminishes the influence of potential surface artifacts on the results (Nieba et al., 1996; Schuck et al., 1998). This analysis can be extended to any other competitor that completely inhibits ligand binding of the analyte when bound to the analyte. Experimental For low-affinity systems with high chemical off-rate constants, in general, steady-state conditions can be achieved. Accordingly, the binding isotherm can be taken as a “calibration” of the sensor signal as a function of the free analyte concentration. Experiments to derive the surface binding constant can be conducted as described above (see Steady-State Experiments and Binding Isotherm Analysis). In a second variation, for high-affinity systems, mass-transport limited conditions are established by using a large immobilization density. Under these conditions, the initial binding rate constant will be directly proportional to the free analyte concentration (dR/dt = ktcA). The initial binding rate constant should be measured in equidistant concentration intervals for analyte concentrations up to a few times (two-fold) the KD. After either one of these sets of calibration experiments, a set of experiments is performed using a constant concentration of analyte, cA,tot (which should be taken in the range of KD), premixed and equilibrated with different concentrations of the soluble form of the ligand. The ligand concentrations, cL, should be taken in two-fold steps, spanning the range from far below to far above the KD. Choice of the fixed analyte concentration is important, since too high a value for cA,tot would show competition only in the range of stoichiometric binding. For systems that do not withstand regeneration, a recycling competitive titration procedure has been developed (Schuck et al., 1998). Data analysis For the high-affinity variant of the experiment with a linear initial rate of binding, dR/dt = ktcA, a plot of dR/dt versus cA can be used to graphically calculate the amount of free analyte, cA,free, during the competition experiment, or cA,free is calculated according to the analytical inversion cA,free(cL) = dR/dt(cL)/kt. Similarly, in place of the linear function, an empirical calibration function for dR/dt(cA) could be used. The obtained concentration of free analyte cA,free in experiments at different soluble ligand concentrations cL can be plotted as cA,free versus log(cL), which gives a competition isotherm with the characteristic sigmoid shape. It can be fitted with Equation 17.6.6, the solution of the quadratic equation of the mass action law combined with mass balance:

F

$IUHH

=F

$WRW

−

(F

$WRW

+F +. /

VRO '

−

F

$WRW

+F +. /

VRO '

− F

$WRW

F

/

)

Equation 17.6.6

which gives the solution binding constant of analyte and ligand, KDsol. If the low-affinity variant of the experiment was used, the steady-state analysis of the sensor response in the absence of soluble ligand can be performed according to Equation 17.6.5, giving Rmax and KDsurface. This can be used in a direct analysis of the competition steady-state response using:

5 F

/

= +

.

VXUI '

 F 

$WRW

−

(F

5

PD[

$WRW

+F +. /

VRO '

−

Equation 17.6.7

F

$WRW

+F +. /

VRO

'

− F

$WRW

)

F  /



Supplement 22

Alternatively, both titrations can be analyzed simultaneously in a global fit to both isotherms, assuming the binding constants to the surface sites and in solution to be identical. COMMON EXPERIMENTAL OBSTACLES Two of the most important experimental problems are mass transport limitations and the effect of aggregates on the binding kinetics. The first difficulty is found most frequently (but not exclusively) in systems of medium to high affinity with high kon, whereas the second is observed predominantly in systems with lower affinity, where higher analyte concentrations are required. Other obstacles can be steric hindrance effects of binding to adjacent surface sites (O’Shannessy and Winzor, 1996), or the contribution of second-order kinetics to the binding process in cuvette systems (Edwards et al., 1998). Most of these processes scale with the immobilization level of the ligand. For this reason, a low ligand density is generally advantageous in minimizing potential problems. Special considerations for working at very low ligand densities are described in Ober and Ward (1999a,b). The use of control experiments (e.g., comparison of the KD from solution competition experiments with the KD obtained from surface binding kinetic measurements) is very useful in verifying the absence of artifacts. The role of mass transport and analyte aggregates will be explained in detail below. Mass Transport Limitation If the supply of analyte to the sensor surface is the limiting factor in the surface binding rate, the observed binding kinetics are mass transport limited. This will depend directly on both the density of the immobilized sites, and on the chemical on-rate constant of the interaction. In the association phase, this will produce a zone of depleted analyte concentration in the vicinity of the surface, while in the dissociation phase, this will lead to a zone of nonvanishing free analyte concentration, which is subject to rebinding-retention effect (Silhavy et al., 1975; Fig. 17.6.5). The quantitative influence of this process on the observed surface binding kinetics is governed by a highly complex reaction/diffusion/convection process (Yarmush et al., 1996; Schuck, 1996). Although the processes in the association and dissociation phase are conceptually closely related, it cannot be assumed that the ratio of the “apparent” rate constants still reflects a good estimate of the equilibrium constant. Also, it is strongly recommended that the nonspecialist reader refrain from modeling using mass-transport-corrected kinetic models. Instead, the primary goal is to identify mass transport limitations, and to establish experimental conditions avoiding transport effects. The following is a list of diagnostic indicators, in the order of reliability, for mass transport limited binding. 1. Dependence on surface capacity. This is a very good indicator of mass transport limitation, since the transport limit always directly scales with the density of active surface sites. For this reason, variations of the immobilization density of the ligand are recommended as routine tests demonstrating the absence of transport limitation and other surface-related artifacts. This test can fail in the presence of significant non-specific analyte binding to the surface.


2. Increased dissociation rate when a soluble form of the ligand is injected (Fig. 17.6.5D). This always strongly indicates rebinding (mass transport limitation) if detected. Since the ligand by itself should not interact with the surface, its effect is the binding to analyte near the sensor surface, preventing it from rebinding and allowing diffusion and washout from the surface (Fig. 17.6.5C). Unfortunately, this



A

analyte flow cA

depletion zone

c 0

C

competitor in flow

cA=0

Signal (%)

D

100 competitor c L

80 60

rebinding

40 20

no rebinding

0 0

100

200

300

400

500

Time (sec)

Figure 17.6.5 Mass transport effects on the surface binding process. (A) In the association phase, insufficient transport does not fully replenish the free analyte in a zone near the sensor surface (depletion zone). The inability to maintain the concentration cA in the depletion zone leads to a limited surface binding rate. (B) The corresponding effect of mass transport limitation in the dissociation phase is that the surface is insufficiently washed and dissociated analyte is insufficiently removed. This generates a zone near the surface in which free analyte (after dissociation from the ligand) can rebind to empty ligand sites before diffusing into the bulk flow. This zone is indicated as retention zone. (C) Introduction of an excess of the soluble form of the ligand as a competitor into the buffer of the dissociation phase helps prevent rebinding to the surface. Soluble ligand can bind to dissociated analyte before rebinding takes place, and the soluble complex can diffuse into the buffer flow. This can allow the measurement of koff free of mass transport effects. (D) Time course of dissociation during a mass transport induced rebinding situation (as depicted in panel B), and after introduction of a soluble competitor into the washing buffer (arrow). In the presence of the competitor, the signal reflects the chemical off-rate constant koff, while in the rebinding situation, the apparent rate governing the signal is reduced.



Supplement 22

effect may not be present for large ligands or ligands/analytes with high nonspecific binding if they exhibit limited diffusivity. 3. Double exponential dissociation phase. This will be seen only after dissociation from more than 50% occupation of all available surface sites. 4. Weak dependence on the flow rate (stirring speed in cuvette-based systems). Since the transport parameters only change with the cube root of the flow rate, i.e., generating only a factor of 2 when changing the flow rate by a factor of 10, this flow rate dependency can be difficult to detect. This is particularly true if the flow rate is only varied by a factor of 2, and the reaction is only partially transport influenced. Then, only a ∼10% change in the apparent koff would be expected in cases where the true koff is 100% larger than the apparent koff. 5. Linear association phase. This effect may not always be present, in particular at substantial transport limitation (Schuck, 1996; Schuck, 1997b; Schuck and Minton, 1996b). Generally, tests 1 and 2 are the most reliable. They also lead to experimental techniques that can be utilized to reduce or eliminate mass transport artifacts. The most effective way for reducing mass transport influence is lowering the surface density of the immobilized ligand. Higher flow rates give only comparatively very small improvements, but are connected with strongly increased sample volume requirements. If the surface density of the ligand cannot be reduced further without leading to an insufficient signal-to-noise ratio, then switching from kinetic experiments to steady-state or competition steady-state experiments is the best solution. This will give information on the equilibrium constant. The kinetic rate constants can then be estimated best from a saturation experiment (approaching complete saturation of all surface sites), followed by a dissociation phase during which soluble ligand is coinjected. The soluble ligand will minimize rebinding and allow the estimation of the chemical off-rate constant, from which the chemical on-rate constant can be determined via Equation 17.6.3. Analyte Aggregates Oligomeric aggregates of analyte can be troublesome in biosensor experiments in two different ways. First, if trace amounts of higher oligomers are present in the analyte sample, this will lead, in the association phase, to a slow accumulation at the sensor surface, which can be visible as a slower second phase of binding. As depicted in Figure 17.6.2, these multimeric analyte aggregates can have multiple interactions with immobilized ligand molecules, and therefore they will dissociate much more slowly than the monomeric analyte. Consequently, they will appear in the dissociation phase as a submoiety with a very low off-rate constant (Davis et al., 1998). The troublesome trace amounts of oligomers can be eliminated by careful chromatographic purification, or their influence can be minimized by exchanging the role of analyte and ligand (Davis et al., 1998; Andersen et al., 1999). This is illustrated in the example of Figure 17.6.6, which demonstrates the importance of both the sample preparation and the choice of ligand and analyte.


The second potentially problematic form of aggregation is a surface-induced multimerization of the analyte. Because the local macromolecular concentrations at the sensor surface are very high (e.g., in the order of 10 mg/ml at a signal of 1000 RU in Biacore instruments), local crowding effects combined with non-specific interactions of the analyte can promote oligomer formation at the sensor surface (Fig. 17.6.7A; Minton, 1995, 1998). As with the influence of preformed aggregates, this process will lead to biphasic association and dissociation profiles, with the slower phase resulting from oligomer accumulation and dissociation, respectively. This process will be favored by



B F2

F3

Signal (RU)

300

A

F3

F2

200

F1 100

0 20

0

40

60

800

F1

700 Signal (RU)

C

600 500 400 300 200 100

0 0

Steady-state signal (RU)

D

100

200

300 400 Time (sec)

500

350 300 250 200 150 100

50 0 1

10 Concentration (µM)

100

Figure 17.6.6 Exemplary data of the interaction of TCR with superantigen (for details of this interaction see Andersen et al., 1999). (A) Elution profile of the TCR from size-exclusion chromatography, indicating the fractions used for kinetic analysis of the interaction with immobilized superantigen, as shown in (B). It should be noted that fraction 1 (bold solid line), fraction 2 (dashed line), and fraction 3 (solid line) all exhibit multiphasic binding, with different relative magnitudes of the slower component. It should also be noted that despite the lower concentration of fraction 1, which results in the lowest response in the association phase, the signal in the dissociation phase is highest. This slower kinetic component in the binding progress curve reflects different amounts of aggregates bound to the sensor surface. The aggregates have a slower dissociation because of their multivalent attachment. For comparison, the same interaction is shown in the reverse orientation, with immobilized TCR and soluble superantigen (dotted line). (C) Sequence of association-dissociation curves at different superantigen concentrations, binding to immobilized TCR. In this orientation, the association and the dissociation is much faster, virtually monophasic, and the binding is completely reversible, which provides further evidence that the slow kinetic components introduced in the different fractions of the soluble TCR sample in Panel B are artifacts. For quantitative analysis, the signal measured at a nonfunctionalized surface (dotted line) is subtracted in order to remove the bulk refractive index contribution of the analyte. (D) A plot of the net steady-state binding signal versus superantigen concentration allows the measurement of the affinity of the interaction free from artifacts due to TCR aggregation that would be introduced in the configuration of Panel B. Macromolecular Interactions in Cells


Supplement 22

A

500 KD >> 1 µM

Signal (RU)

400

300

200

100

0 0

B

20

40

60

1400

80

100

KD = 0.3 µM

Signal (RU)

1200 1000 800 600 400 200 0 0

100

200

300

400

Time (sec)

Figure 17.6.7 Effects of the ligand density on the binding kinetics for interactions with relative low affinity (A) and medium affinity (B). Each panel shows the interaction of immobilized superantigen with single-chain TCR, observed at high ligand surface density (1700 RU, solid lines) and low ligand surface density (700 RU, dashed lines). For easier comparison, the signal obtained at the lower density surface was scaled proportionally. In panel A, a slow phase of binding in the association phase and a residual binding (possible slow dissociation of multivalently bound aggregates) is introduced at high ligand density, under otherwise identical conditions. For the interaction in panel B, the chemical off-rate constant is smaller than for the low-affinity interaction shown above. Nevertheless, from comparison of the binding curves at different ligand surface densities (under otherwise identical conditions) it is obvious that an increased ligand density has significant effects on the surface binding kinetics. This observation could be due to aggregation or to mass transport limitations, both of which are more likely at higher ligand densities.




Table 17.6.1

Troubleshooting Guide for Measuring Protein Interactions by Optical Biosensors

Problem

Solution

No electrostatic preconcentration achieved; poor immobilization

Desalt protein (e.g., using spin column or microdialysis); decrease pH of buffer used for immobilization (should be below pI of protein). Analyte may be too hydrophobic, or there may be electrostatic interaction with surface. Increase salt or detergent concentration in running buffer.a If this does not result from high analyte concentrations, dialyze the analyte against running buffer, or use a spin column for buffer exchange. Use increasingly harsher conditions for regeneration; test procedures used in affinity chromatography; check for strong nonspecific binding of the analyte; check for possible incomplete blocking of activated surface sites after immobilization. Check for presence of free biotin in sample. Biotin on the analyte may not be accessible by surface-immobilized streptavidin; in this case try biotinylated linker. Check for mass-transport artifacts; check for possible traces of aggregates and for formation of aggregates at the surface; change immobilization method (avoid random coupling, avoid large surface densities). If this is not successful, go to steady-state analysis methods. Use longer injections by increasing injection volume and/or decreasing flow rate. If sample volume is limiting, try an equilibrium titration. Decrease immobilization density.b If immobilization density cannot be lowered further, go to steady-state analysis, combined with establishment of lower limit of kdis from dissociation after saturation. Potential signature of mass transport; lower the immobilization density. Signature of mass transport limitation; lower the immobilization density.

Nonspecific binding is high

High signal from buffer; refractive index changes

Analyte does not come off after regeneration

No binding of biotinylated sample to streptavidin surface

Kinetics does not follow 1:1 binding

No steady-state binding is reached in a flow system Mass transport limitation

Increasing slope in the association phase Increasing signal in the dissociation phase

aBe aware of effects of nonspecific binding on the binding kinetics, which substantially decreases the diffusivity of the

analyte across the sensor surface, potentially leading to mass transport artifacts that cannot be detected through change of the ligand density. bIncreasing the flow rate affects transport much less, but consumes much more sample.

higher surface concentrations—i.e., higher density of immobilized ligand, and by higher ligand affinity (Fig. 17.6.7). As with mass transport limitations, they can be detected by variation of the surface density of the ligand. They also can be reduced by lowering the surface density of the immobilized ligand, by size-exclusion chromatography immediately prior to the experiment, or by exchanging the role of ligand and analyte. Macromolecular Interactions in Cells


Supplement 22

Troubleshooting As with other complex biophysical techniques and all investigations involving biological samples, it is impossible to give guidelines which are even nearly complete or generally helpful for troubleshooting. Nevertheless, Table 17.6.1 presents a small list of possible solutions to potential problems, given in the hope that some readers may find them helpful. SUMMARY Binding studies with optical biosensors can be very powerful and versatile. Among the most important virtues are their high sensitivity and utility for a broad range of affinities, real-time detection allowing studies of binding kinetics, and relatively low requirements of sample volume. We have outlined some general strategies and described some of the most commonly used techniques. The work with protein interactions at a surface can introduce additional experimental difficulties as compared to solution methods. As a general rule in experiments with optical biosensors, it is highly recommended that analyses be performed in different ways so that the consistency of the results can be tested. Biosensors can be an excellent tool in the study of protein interactions, and become particularly powerful if combined with other methods. ACKNOWLEDGMENT The authors are grateful for helpful comments by D. Margulies in the preparation of the manuscript. They acknowledge D. Kranz and K. Karjalainen for providing the samples used for some of the experiments shown. Peter Andersen acknowledges grant support by the Danish Natural Sciences Research Council. LITERATURE CITED Andersen, P.S., Lavoie, P.M., Sekaly, R.P., Churchill, H., Kranz, D.M., Schlievert, P.M., Karjalainen, K., and Mariuzza, R.A. 1999. Role of the T cell receptor alpha chain in stabilizing TCR-superantigen-MHC class II complexes. Immunity 10:473-483. Buckle, P.E., Davies, R.J., Kinning, T., Yeung, D., Edwards, P.R., Pollard-Knight, D., and Lowe, C.R. 1993. The resonant mirror: A novel optical sensor for direct sensing of biomolecular interactions. Part II: Applications. Biosens. Bioelectron. 8:355-363. Davis, S.J., Ikemizu, S., Wild, M.K., and van der Merwe, P.A. 1998. CD2 and the nature of protein interactions mediating cell-cell recognition. Immunol. Rev. 163:217-236. Edwards, P.R., Gill, A., Pollard-Knight, D.V., Hoare, M., Buckle, P.E., Lowe, P.A., and Leatherbarrow, R.J. 1995. Kinetics of protein-protein interactions at the surface of an optical biosensor. Anal. Biochem. 231:210-217. Edwards, P.R., Maule, C.H., Leatherbarrow, R.J., and Winzor, D.J. 1998. Second-order kinetic analysis of IAsys biosensor data: Its use and applicability. Anal. Biochem. 263:1-12. Garland, P.B. 1996. Optical evanescent wave methods for the study of biomolecular interactions. Q. Rev. Biophys. 29:91-117. Gershon, P.D. and Khilko, S. 1995. Stable chelating linkage for reversible immobilization of oligohistidine tagged proteins in the Biacore surface plasmon resonance detector. J. Immunol. Methods. 183:65-76. Glaser, R.W. and Hausdorf, G. 1996. Binding kinetics of an antibody against HIV p24 core protein measured with real-time biomolecular interaction analysis suggest a slow conformational change in antigen p24. J. Immunol. Methods. 189:1-14. Hall, D.R. and Winzor, D.J. 1997. Use of a resonant mirror biosensor to characterize the interaction of carboxypeptidase A with an elicited monoclonal antibody. Anal. Biochem. 244:152-160. Hermanson, G.T. 1996. Bioconjugate Techniques. Academic Press, San Diego. Measuring Protein Interactions by Optical Biosensors

Khilko, S.N., Jelonek, M.T., Corr, M., Boyd, L.F., Bothwell, A.L.M., and Margulies, D.H. 1995. Measuring interactions of MHC class I molecules using surface plasmon resonance. J. Immunol. Methods. 183:77-94. Knoll, W. 1998. Interfaces and thin films as seen by bound electromagnetic waves. Annu. Rev. Phys. Chem. 49:569-638.



Leckband, D.E. 1997. The influence of protein and interfacial structure on the self-assembly of oriented protein arrays. Adv. Biophys. 34:173-190. Leckband, D.E., Kuhl, T., Wang, H.K., Herron, J., Muller, W., and Ringsdorf, H. 1995. 4-4-20 anti-fluorescyl IgG Fab′ recognition of membrane bound hapten: Direct evidence for the role of protein and interfacial structure. Biochemistry 36:11467-11478. Lukosz, W. 1991. Principles and sensitivities of integrated optical and surface plasmon sensors for direct affinity sensing and immunosensing. Biosens. Bioelectronics. 6:215-225. Malmborg, A.C. and Borrebaeck, C.A. 1995. Biacore as a tool in antibody engineering. J. Immunol. Methods. 183:7-13. Margulies, D.H., Corr, M., Boyd, L.F., and Khilko, S.N. 1993. MHC Class I/peptide interactions: Binding specificity and kinetics. J. Mol. Recognit. 6:59-69. Margulies, D.H., Plaksin, D., Khilko, S.N., and Jelonek, M.T. 1996. Studying interactions involving the T-cell antigen receptor by surface plasmon resonance. Curr. Opin. Immunol. 8:262-270. Minton, A.P. 1995. Confinement as a determinant of macromolecular structure and reactivity. 2. Effects of weakly attractive interactions between confined macrosolutes and confining structures. Biophys. J. 68:1311-1322. Minton, A.P. 1998. Molecular crowding: Analysis of effects of high concentrations of inert cosolutes on biochemical equilibria and rates in terms of volume exclusion. Methods Enzymol. 295:127-149. Muller, K.M., Arndt, K.M., and Plückthun, A. 1998. Model and simulation of multivalent binding to fixed ligands. Anal. Biochem. 261:149-158. Myszka, D.G., Jonsen, M.D., and Graves, B.J. 1998. Equilibrium analysis of high affinity interactions using Biacore. Anal. Biochem. 265:326-30. Nieba, L., Krebber, A., and Plückthun, A. 1996. Competition Biacore for measuring true affinities: Large differences from values determined from binding kinetics. Anal. Biochem. 234:155-165. Ober, R.J. and Ward, E.S. 1999a. The influence of signal noise on the accuracy of kinetic constants measured by surface plasmon resonance experiments. Anal. Biochem. 273:49-59. Ober, R.J. and Ward, E.S. 1999b. The choice of reference cell in the analysis of kinetic data using Biacore. Anal. Biochem. 271:70-80. O’Shannessy, D.J. and Winzor, D.J. 1996. Interpretation of deviations from pseudo-first-order kinetic behavior in the characterization of ligand binding by biosensor technology. Anal. Biochem. 236:275-283. O’Shannessy, D.J., Brigham-Burke, M., and Peck, K. 1992. Immobilization chemistries suitable for use in the Biacore surface plasmon resonance detector. Anal. Biochem. 205:132-136. O’Shannessy, D.J., Brigham-Burke, M., Soneson, K.K., Hensley, P., and Brooks, I. 1993. Determination of rate and equilibrium binding constants for macromolecular interactions using surface plasmon resonance: Use of nonlinear least squares analysis methods. Anal. Biochem. 212:457-468. O’Shannessy, D.J., O’Donnell, K.C., Martin, J., and Brigham-Burke, M. 1995. Detection and quantitation of hexa-histidine-tagged recombinant proteins on western blots and by a surface plasmon resonance biosensor technique. Anal Biochem. 229:119-124. Plant, A.L., Brigham-Burke, M., Petrella, E.C., and O’Shannessy, D.J. 1995. Phospholipid/alkanethiol bilayers for cell-surface receptor studies by surface plasmon resonance. Anal. Biochem. 226:342-348. Ramsden, J.J., Bachmanova, G.I., and Archakov, A.I. 1996. Immobilization of proteins to lipid bilayers. Biosens. Bioelectronics 11:523-528. Schuck, P. 1996. Kinetics of ligand binding to receptor immobilized in a polymer matrix, as detected with an evanescent wave biosensor. I. A computer simulation of the influence of mass transport. Biophys. J. 70:1230-1249. Schuck, P. 1997a. Use of surface plasmon resonance to probe the equilibrium and dynamic aspects of interactions between biological macromolecules. Annu. Rev. Biophys. Biomol. Struct. 26:541-566. Schuck, P. 1997b. Reliable determination of binding affinity and kinetics using surface plasmon resonance biosensors. Curr. Opin. Biotechnol. 8:498-502. Schuck, P. and Minton, A.P. 1996a. Minimal requirements for internal consistency of the analysis of surface plasmon resonance biosensor data. Trends Biochem. Sci. 252:458-460. Schuck, P. and Minton, A.P. 1996b. Analysis of mass transport limited binding kinetics in evanescent wave biosensors. Anal. Biochem. 240:262-272. Schuck, P., Millar, D.B., and Kortt, A.A. 1998. Determination of binding constants by equilibrium titration with circulating sample in a surface plasmon resonance biosensor. Anal. Biochem. 265:79-91. Schuster, S.C., Swanson, R.V., Alex, L.A., Bourret, R.B., and Simon, M.I. 1993. Assembly and function of a quaternary signal transduction complex monitored by surface plasmon resonance. Nature 365:343-347.



Supplement 22

Sigal, G.B., Bamdad, C., Barberis, A., Strominger, J., and Whitesides, G.M. 1996. A self-assembled monolayer for the binding and study of histidine-tagged proteins by surface plasmon resonance. Anal. Chem. 68:490-497. Silhavy, T.J., Szmelcman, S., Boos, W., and Schwartz, M. 1975. On the significance of the retention of ligand by protein. Proc. Natl. Acad. Sci. USA. 72:2120-2124. Stein, T. and Gerisch, G. 1996. Oriented binding of a lipid-anchored cell adhesion protein onto a biosensor surface using hydrophobic immobilization and photoactive crosslinking. Anal. Biochem. 237:252-259. van der Merwe, P.A. and Barclay, A.N. 1996. Analysis of cell-adhesion molecule interactions using surface plasmon resonance. Curr. Opin. Immunol. 8:257-261. Yarmush, M.L., Patankar, D.B., and Yarmush, D.M. 1996. An analysis of transport resistances in the operation of Biacore; implications for kinetic studies of biospecific interactions. Mol. Immunol. 33:1203-1214.

KEY REFERENCES Davis et al, 1998. See above. Contains a detailed description of analyte aggregation effects on the measured surface binding. Nieba et al., 1996. See above. Demonstration how competition approaches can be used to circumvent kinetic artifacts. O’Shannessy et al., 1992. See above. Collection of immobilization techniques. Schuck, 1997b. See above. General review of the method and its application.

INTERNET RESOURCES http://www.biacore.com Web site for Biacore; extensive list of published biosensor applications http://www.affinity-sensors.com Web site for Affinity Sensors; extensive list of published biosensor applications.

Contributed by Peter Schuck and Lisa F. Boyd National Institutes of Health Bethesda, Maryland Peter S. Andersen University of Maryland Rockville, Maryland




Chromatin Immunoprecipitation for Determining the Association of Proteins with Specific Genomic Sequences In Vivo

UNIT 17.7

Chromatin immunoprecipitation (ChIP) is a powerful and widely applied technique for detecting the association of individual proteins with specific genomic regions in vivo. In this technique, live cells are treated with formaldehyde to generate protein-protein and protein-DNA cross-links between molecules in close proximity on the chromatin template in vivo. A whole-cell extract is prepared, and the cross-linked chromatin is sheared by sonication to reduce average DNA fragment size to ∼500 bp. The resulting material is immunoprecipitated with an antibody against a desired protein, modified (e.g., acetylated, phosphorylated, methylated) peptide, or epitope (in situations where the protein of interest is epitope-tagged). DNA sequences that directly or indirectly cross-link with a given protein (or modified variant) are selectively enriched in the immunoprecipitated sample. Thus, the method is not restricted to sequence-specific DNA-binding proteins. Reversal of the formaldehyde cross-linking by heating permits the recovery and quantitative analysis of the immunoprecipitated DNA. The amounts of specific genomic regions in control and immunoprecipitated samples are determined individually by quantitative PCR. The fold enrichment of certain chromosomal sequences (e.g., presumed binding sites) relative to other chromosomal sequences (e.g., presumed nonbinding sites) provides quantitative information about the relative level of association of a given protein with different genomic regions. Protein association with specific genomic regions can be performed under a variety of conditions (e.g., environmental change, cell-cycle status) and/or in wild-type versus mutant strains. Furthermore, as formaldehyde inactivates cellular enzymes essentially immediately upon addition to cells, ChIP provides snapshots of protein-protein and protein-DNA interactions at a particular time point, and hence is useful for kinetic analysis of events occurring on chromosomal sequences in vivo. In addition, ChIP can be combined with microarray technology to identify the location of specific proteins on a genome-wide basis (see Commentary). This unit describes the ChIP protocol for cells of the baker’s yeast Saccharomyces cerevisiae (see Basic Protocol); however, it is also applicable to other organisms, although some organism-specific modifications related to cell lysis and sonication are necessary. A protocol for eluting immunoprecipitated protein-DNA complexes is also provided (see Alternate Protocol 1). As an alternative to gel electrophoretic analysis of the PCR products, a quantitative PCR analysis in real time with SYBR Green is also provided (see Alternate Protocol 2).

CHROMATIN IMMUNOPRECIPITATION Materials Saccharomyces cerevisae cells to be studied 37% formaldehyde: store up to 1 year at room temperature 2.5 M glycine, heat sterilized TBS (APPENDIX 2A), ice cold FA lysis buffer with and without 2 mM PMSF (see recipe), ice cold ChIP elution buffer (see recipe) 20 mg/ml Pronase (Roche) in TBS; store up to 1 year at −20◦ C TE buffer, pH 7.5 (APPENDIX 2A) 20 mg/ml DNase-free RNase A (see recipe) 10× loading buffer (see recipe)

Contributed by Oscar Aparicio, Joseph V. Geisberg, and Kevin Struhl Current Protocols in Cell Biology (2004) 17.7.1-17.7.23 C 2004 by John Wiley & Sons, Inc. Copyright

BASIC PROTOCOL



Primary antibody against protein or epitope of interest 50% (v/v) protein A-Sepharose beads (Amersham Pharmacia Biotech) or equivalent in TBS FA lysis buffer (see recipe), room temperature FA lysis buffer (see recipe)/0.5 M NaCl ChIP wash buffer (see recipe) Primers (see Critical Parameters and Troubleshooting) 3000 Ci/mmol [32 P]dATP (optional; see annotation to step 30) 2-ml screw-cap microcentrifuge tubes with (relatively) flat bottoms ∼0.5-mm-diameter silica-zirconia (BioSpec; preferred) or glass beads Mini bead beater (BioSpec; preferred) or individual or multivortexer 5-ml syringe 15-ml conical tubes, disposable 25-G needles Sonicator with microtip probe (e.g., Branson Sonifier 250) End-over-end rotator 0.5-ml PCR tube Spin-X centrifuge-tube filter (e.g., Corning) 65◦ C water bath PCR-purification spin column (Qiagen) Software for analyzing PCR primers and products Additional reagents and equipment for growth of Saccharomyces cerevisiae cultures (APPENDIX 3A), phenol/chloroform extraction and ethanol precipitation (APPENDIX 3A), PCR (APPENDIX 3F), agarose gel electrophoresis (APPENDIX 3A), and nondenaturing acrylamide gel electrophoresis (UNIT 6.5) CAUTION: When working with radioactive materials, take appropriate precautions to avoid contamination of the experimenter and the surroundings. Carry out the experiment and dispose of wastes in an appropriately designated area, following guidelines provided by the local radiation safety officer (also see APPENDIX 1D).

Cross-link protein-DNA complexes in vivo 1. For each sample, grow 200 ml Saccharomyces cerevisiae to OD600 = 0.6 to 0.8. CAUTION: Keep cultures covered or work in a fume hood to avoid noxious formaldehyde fumes. The volumes of culture can be reduced (20 ml is a reasonable minimum) or increased depending on need. Typically, 20 to 40 ml yeast is used for an individual immunoprecipitation, so the 200-ml volume permits multiple immunoprecipitations from the same cells. This is particularly useful for experiments involving the analysis of multiple factors or for carrying out independent immunoprecipitations involving the same factor for data reproducibility.

2. Add 5.5 ml of 37% formaldehyde (1% final). Cross-link 15 to 20 min at room temperature by occasionally swirling flask or shaking slowly on a platform. 3. Add 30 ml heat-sterilized 2.5 M glycine and incubate an additional 5 min at room temperature. Glycine stops the cross-linking by reacting with formaldehyde. Determining the Association of Proteins with Specific Genomic Sequences

Harvest cells 4. Centrifuge cells 5 min at 2500 × g, 4◦ C. Discard supernatant into a chemical waste container and resuspend pellet in 50 to 200 ml ice-cold TBS. Repeat once.



5. Centrifuge cells for a third time 5 min at 2500 × g, 4◦ C. Discard supernatant and resuspend cells in 10 ml ice-cold FA lysis buffer. 6. Pellet cells by centrifuging in a benchtop centrifuge 5 min at 3000 rpm, 4◦ C. Discard supernatant. The cells can remain on ice for a few hours while other samples are being collected so that all samples may be processed as a group from this point onward. Alternatively, the cells may be frozen in liquid nitrogen or a dry ice/ethanol bath and stored up to several months at −80◦ C. This is particularly helpful if multiple samples are being generated during a time-course experiment. If cells are frozen, they must be thawed on ice before continuing with the procedure.

Lyse cells and isolate chromatin For lysis using a mini bead beater (preferred) 7a. Resuspend the cell pellet in 1 ml ice-cold FA lysis buffer/2 mM PMSF. Fill threequarters of a 2-ml flat-bottomed screw-cap microcentrifuge tube with ∼0.5-mmdiameter silica-zirconia or glass beads. Add cells, taking care to avoid introduction of bubbles, and screw the cap on tightly. Make sure there are no leaks. The mini bead beater is recommended because it is more efficient at breaking cells (multiple samples can be broken simultaneously). Silica-zirconia beads are more efficient at breaking cells than glass beads and are also recommended. To facilitate cell breakage with the mini bead beater, it is important that the final suspension nearly fill the tube. Do not break >160 OD600 units of cells (i.e., 160 OD600 units of cells (i.e., 1 min. Repeat two more times. Take great care that the sample does not get too hot. If a different sonication device is used, empirically determine the conditions necessary to achieve the desired level of DNA shearing. The shear size is determined as described below (see Critical Parameters and Troubleshooting).

14. Microcentrifuge 30 min at maximum speed, 4◦ C. Transfer the supernatant to a fresh 15-ml disposable conical tube, add 4 ml ice-cold FA lysis buffer, and gently mix by inversion. Remove 250 µl for checking DNA fragment size and freeze the remaining chromatin solution in 800-µl aliquots in liquid nitrogen. Upon sonication, the cross-linked chromatin is solubilized and purified away from the pelleted material which contains cell debris and unbroken cells. The resulting chromatin solution constitutes the input sample for the subsequent immunoprecipitation. The frozen aliquots are stable for many months when stored at −70◦ C and are suitable for immunoprecipitations.

Check chromatin-fragment size 15. Add 250 µl ChIP elution buffer and 20 µl of 20 mg/ml Pronase in TBS to the 250-µl chromatin aliquot. Incubate 2 hr at 42◦ C, followed by 6 hr at 65◦ C. Phenol extract and ethanol precipitate sample (APPENDIX 3A). While it is convenient to perform the reaction in a PCR machine overnight, it could just as easily be done in heat blocks or water baths. The same is true of the incubation described in step 26.

16. Resuspend in 30 µl TE buffer, pH 7.5, add 1 µl of 20 mg/ml DNase-free RNase A, and incubate 15 min at 37◦ C. Add 3 µl of 10× loading buffer and electrophoretically separate the material on a 1.5% agarose gel (APPENDIX 3A). Fragments should be between 100 to 1000 bp, with an average length of 400 to 500 bp. It is important to shear DNA fragments down to an average length of 400 to 500 bp. Longer fragments will increase the background and will decrease the resolution of the region to which the protein associates (see Commentary). Determining the Association of Proteins with Specific Genomic Sequences

Immunoprecipitate 17. Incubate 800 µl chromatin solution with 10 µl primary antibody against the protein or epitope of interest and 20 µl of 50% (v/v) protein A–Sepharose beads in TBS on an end-over-end rotator 90 min at room temperature.



The actual amount of antibody needed has to be empirically determined and can vary considerably. The idea is to have an excess of antibody to efficiently precipitate at least 50% of the antigen in question. One way to assess the efficiency of antigen immunoprecipitation is to determine the amount of antigen present in the sample before and after the immunoprecipitation. An aliquot of 30 µl chromatin solution, taken before and after immunoprecipitation, is usually sufficient to visualize the protein of interest via immunoblotting (UNIT 6.2) and standard chemiluminescent detection; however, the samples have to be boiled in SDS/PAGE sample buffer for 30 min prior to loading in order to reverse the formaldehyde cross-links. The immunoprecipitation conditions can be varied (e.g., time, temperature, salt concentration, presence of detergents) if necessary. Protein A–Sepharose beads are used here because they work well with most monoclonal and polyclonal sera derived from mouse and rabbit, respectively. In some cases, the use of other beads (e.g., protein G–Sepharose) may improve binding of some antibodies, including rat IgG (see Table 7.2.1).

18. Microcentrifuge beads 1 min at 3000 rpm, room temperature. Transfer 300 µl supernatant into a 0.5-ml PCR tube labeled “INPUT.” Discard the rest of the liquid.

Wash beads 19. Resuspend beads in 700 µl FA lysis buffer, room temperature, and transfer mixture into a Spin-X centrifuge-tube filter. The use of Spin-X filters aids in the recovery of the beads after washes and results in better uniformity between different samples. The procedure is also substantially faster with the filters, particularly when multiple samples are processed simultaneously. Alternatively, one could use conventional microcentrifuge tubes for the washes and aspirate the supernatant with a narrow-bore pipet tip after each spin.

20. Place the filter into a 1.5-ml microcentrifuge tube and mix sample 3 min on an endover end rotator. Microcentrifuge 2 min at 3000 rpm, room temperature. Discard the flow-through liquid at the bottom of the tube. 21. Add 700 µl FA lysis buffer, room temperature, to the beads and repeat step 20.

Elute protein from beads 22. Wash beads for 3 min each with 700 µl FA lysis buffer/0.5 M NaCl, 700 µl ChIP wash buffer, and finally 700 µl TE. For many polyclonal antibodies, the more stringent washes in this step result in a cleaner signal, while gentle washes frequently lead to an unacceptably high background. For some antibodies (e.g., monoclonal against peptide epitopes; see Alternate Protocol 1), repeated washes with FA lysis buffer, which are gentler, might be more appropriate.

23. Place filter unit containing the beads into a new 1.5-ml microcentrifuge tube and add 100 µl of ChIP elution buffer. Gently pipet up and down two or three times in order to dislodge beads from the filter. Incubate 10 min in a 65◦ C water bath. A water bath is used instead of other heating apparatuses in order to improve heat transfer.

24. Microcentrifuge beads 2 min at 3000 rpm, room temperature. Discard filter with beads. Transfer the eluate into a 0.5-ml PCR tube labeled “IP.”

Reverse cross-links and purify DNA 25. Add 80 µl TE and 20 µl Pronase in TBS to the IP tube. Combine 20 µl INPUT material (step 18), 100 µl ChIP elution buffer, 60 µl TE, and 20 µl TBS into a new 0.5-ml PCR tube. 26. To reverse cross-links, place tubes into a PCR machine. Incubate 2 hr at 42◦ C, followed by 6 hr at 65◦ C. Store samples at 4◦ C until use.



Supplement 23

The incubation at 42◦ C allows for Pronase digestion of cross-linked polypeptides, while the 65◦ C incubation results in a reversal of the formaldehyde cross-links.

27. Purify DNA using a Qiagen PCR-purification spin column as per manufacturer’s instructions. This will require double loading of the spin column (i.e., 600 µl spin through and then repeat). Alternatively, add 20 µl of 4 M LiCl and purify by extracting with 25:24:1 phenol/ chloroform/isoamyl alcohol, followed by extraction with chloroform and ethanol precipitation (APPENDIX 3A). It is useful to add 2 µl of Pellet Paint (Novagen) prior to the addition of ethanol, as this aids both the ethanol precipitation and visualization of the very small pellet.

28. Resuspend in 300 µl TE and store at −20◦ C. DNA pellets stored in this fashion should be stable for years.

Perform quantitative PCR 29. Design primer pairs for the desired genomic regions to be examined. Success in obtaining high-quality data is critically dependent on good primer design (see Critical Parameters and Troubleshooting). In general, primers should be 20 to 30 bases long with a Tm of 55◦ to 60◦ C. The design of good primers is greatly facilitated by commercially available software packages such as Oligo 6.6 (see http://www.oligo.net) or Primer Express 1.5 (see http://www.appliedbiosystems.com). Most primers require no purification or special treatment prior to PCR. Amplification products should be 75 to 350 bp; longer products should be avoided, as the amplification efficiency is substantially lower. A final primer concentration of 1 µM works well for most primers, but in some instances, improved product specificity may be obtained by lowering the final primer concentration 5 to 10 fold. Refer to APPENDIX 3F for more information.

30. Dilute INPUT DNA (obtained from step 18) in three separate tubes by a factor of 5, 10, and 20. Set up standard PCR reactions (APPENDIX 3F) with 2 µl DNA sample, primers at 1 pmol/µl, and total reaction volumes of 10 to 50 µl. If PCR products will be detected by radioactivity, add 1 µCi of 3000 Ci/mmol [32 P]dATP. For a typical measurement, the three dilutions of input DNA are tested along with duplicate immunoprecipitated samples (or undiluted and 5-fold diluted immunoprecipitated samples). This permits an assessment of whether the assay is being performed in the linear range as well as of the reproducibility of the PCR reaction. The immunoprecipitated DNA is typically used without dilution, although it is useful to analyze different amounts to ensure that it is also in the linear range. There are several key parameters for achieving an optimum PCR reaction. For example, it is very important to have a quality repeat pipettor that can reproducibly dispense 2 µl DNA. Pipetting inaccuracies at this stage will lead to greater well-to-well variability and poorer reproducibility among identical samples. Additionally, multiple primer pairs (up to 4 to 5) can be included in the same reaction, provided that the PCR products can be unambiguously resolved from each other by gel electrophoresis. This permits simultaneous and internally controlled analysis of multiple genomic regions in a single reaction. However, it is critical to ensure that there is no competition between the different primer pairs and PCR products. Also, comparable results are obtained when PCR reactions are performed in volumes between 10 to 50 µl; using smaller volumes reduces the cost and facilitates loading of the reaction products on gels. See Critical Parameters and Troubleshooting for a discussion of primer choice. Determining the Association of Proteins with Specific Genomic Sequences

Detection of PCR products by [32 P]label is recommended over detection by ethidium bromide or SYBR Green (see Alternate Protocol 2) staining as it improves the sensitivity and extends the linear range of detection; however, it necessitates using the usual precautions in working with radioactivity.



31. Carry out hot-start PCR using the following thermal cycling parameters.

Initial step: 26 cycles:

Final step:

10 min 30 sec 30 sec 1 min 4 min

95◦ C 95◦ C 55◦ C 72◦ C 72◦ C

(denaturation) (denaturation) (annealing) (extension) (final extension).

These conditions are generally appropriate for most situations. The annealing temperature may have to be adjusted if the melting temperatures of the primers is substantially above or below 55◦ C. The number of cycles might also have to be adjusted in some cases if reactions are not in the linear range. See Critical Parameters and Troubleshooting for more details.

Analyze PCR products 32. Add the appropriate loading buffer to the PCR products, and analyze by electrophoresis on nondenaturing polyacrylamide (UNIT 6.5) or agarose gels (APPENDIX 3A). The gels should be stained either with ethidium bromide or SYBR Green dyes, or analyzed by autoradiography or PhosphorImager.

33. Quantitate the relative amount of PCR products using appropriate software for the accompanying instrument. 34. Calculate the apparent immunoprecipitation efficiency for a specific fragment by dividing the amount of PCR product obtained with the immunoprecipitated DNA by the amount obtained with the input DNA. A volume of 2 µl immunoprecipitated DNA sample (1/150 total immunoprecipitated material) contains ∼200 times the number of cell equivalents as 2 µl INPUT sample that has been diluted 5-fold (1/30,000 of the original aliquot that was immunoprecipitated). Thus, if the amount of PCR product in the immunoprecipitated sample is equal to the amount of PCR product in the 5-fold diluted INPUT sample, the apparent immunoprecipitation efficiency is 0.5%. The apparent immunoprecipitation efficiency for the background signal is typically ∼0.025% to 0.05%, and it should not be higher than 0.1%.

SPECIFIC PEPTIDE ELUTION OF PROTEIN-DNA COMPLEXES IMMUNOPRECIPITATED FROM CROSS-LINKED CHROMATIN Peptide elution represents an alternative method for removal of immunoprecipitated protein-DNA complexes from beads. In this procedure, beads containing the immunoprecipitated complexes are incubated with high concentrations of a peptide recognized by the antibody used in the immunoprecipitation. The added peptide competes with the protein antigen of interest for binding to the antibody, and specifically liberates the protein-DNA complexes from the beads. The high specificity of peptide elution reduces the nonspecific background (typically by a factor of 2 to 4), which makes it the method of choice, particularly for applications where the expected immunoprecipitation signal is low. Peptide elution is especially useful for chromatin immunoprecipitation experiments involving proteins that are tagged with the HA or myc epitopes (in single or multiple copies); however, it would also be appropriate in cases where the antibody used for the immunoprecipitation was generated against a defined peptide sequence. Peptide elution is slightly more expensive than conventional elution, due to the cost of the peptide. In general, peptide elution should be used in conjunction with gentle washes during the immunoprecipitation procedure described below, which minimizes antigen leaching. Stringent washes, such as those employed in the main method (see Basic Protocol), will often result in signals that are several-fold lower, with little or no improvement in background. Finally, peptide elution may vary in quality depending on factors such as the number of epitopes in the antigen and the relative stability of the antibody-antigen interaction.




Supplement 23

Additional Materials (also see Basic Protocol) 1 mg/ml peptide (e.g., myc, HA) in TBS (see APPENDIX 2A for buffer) For this protocol, follow steps 1 to 21 of the main method (see Basic Protocol), replace steps 22 to 25 with the following, and continue with step 26 onwards. 22. Repeat FA lysis buffer wash (see Basic Protocol, steps 20 and 21) three additional times for a total of five washes. Repeated washes with FA lysis buffer are much more gentle than the single washes with FA lysis buffer/0.5 M NaCl, ChIP wash buffer, and TE used in the Basic Protocol and result in higher signal-to-background ratios.

23. Place the Spin-X centrifuge-tube filter unit containing the beads into a new 1.5-ml microcentrifuge tube and add 100 µl of 1 mg/ml peptide (typically myc or HA) in TBS. Gently pipet up and down two or three times in order to dislodge beads from the filter. Incubate 30 min at 30◦ C. 24. Microcentrifuge beads 2 min at 3000, room temperature. Discard filter with beads. Transfer the eluate into a 0.5-ml PCR tube suitable for PCR labeled “IP.” 25. Add 150 µl TE, pH 7.5, 250 µl of ChIP elution buffer, and 20 µl of 20 mg/ml Pronase in TBS. ALTERNATE PROTOCOL 2

ANALYSIS OF CHROMATIN IMMUNOPRECIPITATION EXPERIMENTS BY REAL-TIME QUANTITATIVE PCR WITH SYBR GREEN Quantitative PCR (QPCR) analysis in real time with SYBR Green has several advantages over the analysis of PCR reactions by gel electrophoresis (see Basic Protocol, step 32). First, the method saves considerable time because no gels are involved and because quantitative values are obtained directly from the data curves and do not require densitometry or phosphor imager analysis. As a consequence, this approach permits very rapid analysis of much larger numbers of chromatin immunoprecipitation samples than can be performed with the Basic Protocol. Using standard 96-well instruments, it is a straightforward procedure to analyze 100 to 200 samples/day (in replicates of three) with only 1 to 2 hr of total setup time. With newer 384-well instruments and automated robotics equipment, sample throughput can be further increased to thousands per week. Second, the data generated by this procedure are more accurate and reproducible, because quantitative values are determined from continuous sampling throughout the PCR reaction rather than a single end-point determination. Furthermore, the quality and “linear range” of every PCR reaction are directly visualized. Third, the procedure is significantly safer for the researcher, as no radioactive materials or toxic acrylamide are used. The major disadvantage of this procedure is that the measurements are performed individually and hence are not internally controlled, whereas the Basic Protocol permits the simultaneous analysis of multiple genomic regions in a single PCR reaction (provided the individual primer pairs function independently). As such, the Basic Protocol is more useful for analyzing the same small set of genomic regions under multiple experimental conditions and for simultaneous analysis of electrophoretically distinguishable alleles of a given genomic region.

Determining the Association of Proteins with Specific Genomic Sequences

SYBR Green is a sensitive and highly selective double-stranded DNA (dsDNA)–binding dye that remains associated even at the high temperatures normally used for PCR template extension. Real-time PCR reactions involving SYBR Green are performed with standard oligonucleotide primers, and hence are much less expensive than real-time PCR reactions using fluorophore-conjugated oligonucleotides (e.g., TaqMan or Lux probes). Measurements of SYBR Green fluorescence at the polymerase extension step of PCR, when



plotted against PCR cycle number, provide both a qualitative assessment of the progress of the PCR and a way to quantitate the relative amount of DNA template initially present in the reaction. Typical real-time QPCR graphs feature the plot of the log10 (Net fluorescence) on the y axis versus the PCR cycle number on the x axis, and usually contain three well-defined stages: (1) baseline, (2) linear, and (3) plateau. In the baseline stage, the amount of DNA product formed is still below the sensitivity threshold of SYBR Green, so product formation is undetectable. This part of the curve is typically used as a baseline for SYBR Green signal drift. The linear part of the curve is the most important from the analytical standpoint, because it is at this stage that the rate of PCR product accumulation is both constant on a per-cycle basis and readily detectable by increased SYBR Green fluorescence. Finally, as all of the SYBR Green in the reaction becomes bound to the recently synthesized PCR products, the amount of fluorescence stays constant from cycle to cycle and the reaction reaches a plateau. In the protocol described below, PCR is performed under special conditions that minimize the inhibitory effects of SYBR Green on Taq activity and maximize the linear range of product detection. After amplification is complete, raw data are stripped of outliers and exported in a format readable by a spreadsheet program such as Microsoft Excel. Finally, data points from replicate samples are averaged, and mean values are further manipulated and ultimately compared to some internal reference or control.

Additional Materials (also see Basic Protocol) Input DNA (see Basic Protocol, step 28) Immunoprecipitated fragments (“IP” sample; see Basic Protocol, step 23) 2× SYBR Green Taq mix (see recipe) Real-time PCR machine and corresponding software (e.g., ABI) 96-well PCR plates (ABI, cat. no. 4306737) and optical adhesive covers Centrifuge with swinging-bucket rotor and microtiter plate adapter Spreadsheet program (e.g., Microsoft Excel) Set up PCR reactions 1. Dilute input DNA to an approximate equivalent of 1 × 106 cells/ml in TE buffer, pH 7.5. If immunoprecipitations were performed as described in the Basic Protocol, then a 1:25 dilution of the input sample from step 28 will result in 1:1000 overall dilution and will correspond to ∼5 × 108 to 1 × 109 cell equivalents.

2. If necessary, resuspend immunoprecipitated fragments in TE buffer, pH 7.5, so that the approximate cell equivalent is 1 × 109 cells/ml. Immunoprecipitated DNA derived from the IP sample obtained by the Basic Protocol (step 23) is appropriately diluted and needs no further treatment.

3. Prepare PCR primer stocks by mixing each primer pair at a final concentration of 3.3 µM in TE buffer, pH 7.5. It is critical to test newly obtained primer pairs for amplification specificity and performance under conditions that will be used for real-time PCR with SYBR Green (see Critical Parameters and Troubleshooting). SYBR Green can inhibit PCR reactions, and primer pairs that are appropriate for quantitative PCR analysis in the absence of SYBR Green may not work well in the presence of SYBR Green. High-quality primer pairs should result in ∼1.9-fold amplification/cycle (this can be determined from quantitative analysis of raw fluorescence data for each cycle, which is generally available on commercial instruments). Amplified material at the completion of the PCR should contain only one band



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(as assayed on high-percentage agarose or polyacrylamide gels). Specificity information may also be obtained by running dissociation curves on reactions following the conclusion of the PCR run. Typically, samples are melted for 15 min at 95◦ C, cooled to 60◦ C, and then slowly heated back up to 95◦ C over a period of 20 min. Plotting the first derivative of the fluorescence against the temperature allows for simple visual identification of sample heterogeneity. Some instrument-specific software packages have built-in modules for dissociation curve analysis.

4. Select and label the wells to be used in the run. In general, individual samples should be run in triplicate. Obvious outliers occur with some frequency, generally at 500 bp (this is not recommended), the extension time at 72◦ C should be increased to 1 min. See Critical Parameters and Troubleshooting for more details.

6. Using a small-volume automatic pipettor (20-µl capacity), place a 2-µl aliquot of each DNA template into the appropriate wells of a 96-well PCR plate. Gently tap the plate to allow the sample droplets to fall to the bottoms of the wells. It is very important to have a quality repeating pipettor that can reproducibly dispense small volumes of sample into the wells. Pipetting inaccuracies at this stage will lead to greater well-to-well variability and poorer reproducibility among identical samples.

7. Using a small-volume automatic pipettor (20-µl capacity), place a 3-µl aliquot of primer mix (see step 3) into the relevant wells and tap the plate a few times to settle the contents. On many real-time PCR machines, results from 10-µl reactions are virtually indistinguishable from those of 25- and 50-µl reactions in their accuracy and reproducibility. The use of 10-µl reactions provides substantial savings in reagent costs. On some machines, the minimal reaction volume needed for accurate and reproducible results may be greater. Determining the Association of Proteins with Specific Genomic Sequences

8. With a larger automatic pipettor (100-µl capacity), add 5 µl of 2× SYBR Green Taq Mix to every assayed well. Place microtiter plate into appropriate microtiter plate adapter and centrifuge 1 min at 200 × g, room temperature, in a swinging-bucket rotor.



The 2× SYBR Green Taq Mix contains a variety of components that considerably reduce the inhibitory effect of SYBR Green, thereby resulting in more reproducible signals that require fewer amplification cycles. Comparable mixes containing proprietary buffers can be obtained commercially. It is critical that quantitative PCR reactions containing SYBR Green be performed under conditions of efficient amplification (e.g., 1.9-fold amplification/ cycle).

9. Seal plate with clear optical adhesive covers, overlay foam compression pad with gold side facing up, and place into the real-time PCR machine. Secure lid. The details of this step may differ, depending on the machine.

10. Start the PCR protocol (see step 5). After completion, save the run for future analysis.

DATA analysis 11. Open the file containing the real-time data according to the manufacturer’s instructions for the instrument. Although the specific protocol will depend on the software and instrument, the overall logic and approach to the analysis of real-time data is generally applicable.

12. Look at the different curves and set the baseline as needed. Generally, the baseline should be set from cycle 3 to the cycle just prior to where the curves start increasing in a linear fashion. It is desirable to have at least 10 cycles for the calculation of the baseline, as this results in increased accuracy in the subsequent calculations of the threshold cycle.

13. Change the value in the Threshold box to be about halfway up in the linear range, and apply changes to the data set. The threshold cycle is defined to be the PCR cycle at which the fluorescence is 10 times (10 is the default multiplier) the standard deviation obtained in the baseline calculation. When the multiplier is set to 10, the fluorescence at the threshold cycle is considered the lowest fluorescence value that is significantly above the background. In practice, this number frequently lies in the nonlinear range of many of the curves. For later calculations, it is easier to manually set the fluorescence value used to calculate the threshold cycles to 0.04. At this value, all the curves should be in the linear range and well above the baseline, allowing for far more accurate comparisons of the threshold cycles. On occasion, however, it will be necessary to adjust this value either up or down to better reflect the linear range of net fluorescence for most of the curves.

14. Manually select one group of triplicates and visually inspect their amplification plots. If curves are essentially superimposable and the threshold cycle (CT ) values are close to each other (maximal and minimal replicates within 1 cycle, preferably within 0.5 cycles), proceed to the next triplicate sample. Otherwise, remove the outlier and continue to the next triplicate. Decisions regarding the removal of some outliers could either be straightforward or judgment calls, depending on the circumstances. In cases where two out of three curves are superimposable while the third is clearly off by more than a cycle, it is a fairly easy decision to consider the third replicate an outlier. If the curves are closer, the decision on which one to eliminate, if any, becomes much more difficult. As a general guide only, if the spread between the lowest and highest CT values is less than 0.5, it is probably safe to average all the CT values (see step 12). If the CT range is 0.5, the data are less reliable and the decision to remove any data points should probably be made on a case-by-case basis. It is highly recommended that the PCR be repeated for samples where the CT ranges are >1 with no two curves superimposable.

15. Proceed to analyze the data for all triplicates in the manner described above. Save the results in a different file.



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16. Export the data to a spreadsheet program such as Microsoft Excel by using builtin filters. The file should not contain omitted wells (see step 14) and should be in a column format containing well positions, descriptors, and CT values for each selected well. Final calculations are most easily handled in a spreadsheet but could also be performed with a scientific calculator.

17. Open the exported file. Proceed to average triplicate measurements for each sample in a new column (AVERAGE CT ). For some samples, there may be only two measurements left as a result of the removal of outliers in step 14.

18. For each primer pair, subtract the AVERAGE CT (INPUT) from new column. This number is the NET CT .

AVERAGE C T

(IP) in a

This value represents the difference in cycles between the immunoprecipitated sample and the input DNA.

19. Subtract NET CT for one primer pair (experimental; EXPT) from the NET CT of another primer pair that serves as a reference or a control (CTRL) in a new column. The resulting value is NET CT EXPT − CTRL. Repeat NET CT subtraction of control primer for all other experimental primers. It is very desirable to have a control primer pair that can be used to assess the relative cross-linking efficiency at promoters of interest. Frequently, the control primer pair could be specific for a DNA region that does not bind to the immunoprecipitated protein of interest. The signal from the control primer pair could then be considered the background, and the binding efficiency of the protein to different promoter regions could be expressed as fold over background binding.

20. Evaluate the expression POWER ([mean primer slope], [− NET CT EXPT − CTRL]), where the [mean primer slope] is the base and [− NET CT EXPT − CTRL] is the exponent. Repeat the process with other primers by using the different NET CT EXPT − CTRL values calculated in step 19. The actual value calculated in the POWER expression above is the degree of occupancy of the immunoprecipitated protein at the sequence of interest relative to that of a control (or background) DNA region. Perfectly efficient PCR, in which the number of amplified molecules doubles every cycle, has a primer slope of 2. As defined, this value is independent of primer pair sequence, target sequence length, and other variables that under normal circumstances may adversely affect the efficiency of amplification. In practice, however, the mean primer slope is almost always 100-fold, but even a factor of two can be meaningful if the experiment is repeated enough times and the experimentally determined error is sufficiently low. The advantage of this approach is that the identical samples are used to directly determine relative protein association to different genomic regions. Furthermore, differences in fold enrichments for different genomic regions represent relative quantitative measurements of protein association in vivo. Additional controls may be used depending on the specific application. For example,

where binding to a putative binding site is being tested, a mutation in the binding site is a critical control (especially if such a mutation previously was shown to eliminate binding in vitro). In a related manner, it might be useful to determine the protein association in mutant strains or under particular environmental conditions that are suspected to be important for the protein of interest. Cross-linking The extent of formaldehyde cross-linking is an important variable that in principle may be modified by changing the duration of crosslinking, the concentration of formaldehyde, or the temperature at which the cross-linking is performed. The use of 1% (final concentration) formaldehyde for 15 min at temperatures ranging from 12◦ C to 37◦ C usually works well; however, at temperatures above 30◦ C, background sometimes increases. Therefore, when fixation at a higher temperature is required, reducing the duration of cross-linking or the formaldehyde concentration may be helpful. Excessive cross-linking can interfere with cell breakage by bead beating and effective fragmentation and solubilization of the DNA by sonication (see below). For some applications where protein cross-linking is particularly efficient (e.g., histones), it might be useful to decrease the cross-linking time or formaldehyde concentration. In particular, histone tails have a number of lysine residues that are likely to be modified by formaldehyde, and such modified lysines may interfere with the binding of antibodies against specific peptides corresponding to modified histones (e.g., by acetylation, phosphorylation, methylation). Cell lysis Although complete lysis of all cells is not absolutely necessary (and may be difficult to achieve), it is very important that lysis be as efficient as possible. Efficient lysis is important to obtain a reproducible degree of cell breakage among a group of samples to reliably compare results. Significant differences in cell lysis efficiency will result in immunoprecipitations with different ratios of antibody to chromatin, which will possibly alter immunoprecipitation efficiency. Cell breakage by a mini bead beater is generally more efficient than breakage by a multi-vortexing apparatus, although both methods work. In both cases, it is important to use flat-bottomed 2-ml microcentrifuge tubes. When using the mini bead beater, the sample and beads should nearly fill the tube, whereas for vortexing it is important



to maintain a small volume. The extent of cell breakage may be monitored microscopically by comparing the number of intact cells (determined by counting on a hemacytometer) in small, diluted aliquots of the sample taken before and after vortexing. In addition, the size of the remaining pellet (unbroken cells and debris) obtained in the first centrifugation following sonication (see Basic Protocol, step 14) is a good general indicator of the extent of lysis. The size of this pellet should be routinely checked (by rapid visual inspection) to compare the extent of lysis among samples. The final yield of genomic DNA in the extract is also an important indicator of the extent of cell breakage, although the DNA yield is also dependent on the solubilization of chromatin by sonication (see below). Poor or variable cell breakage may result from excessive cross-linking that toughens the yeast cell wall and other structures. The procedure for lysis of Saccharomyces cerevisiae is appropriate for other yeast species. However, modified procedures are necessary for breaking mammalian cells. Sonication Shearing DNA to a small size (∼500 bp average) by sonication is the critical factor in achieving resolution between a DNA sequence where a particular protein is bound and a nearby (cis-)DNA sequence that does not bind that protein. In addition, fragmentation of the chromatin is essential for its solubilization from the ruptured cells. As indicated above, the ability to fragment and solubilize the chromatin depends on the extent of chromatin cross-linking. In general, more cross-linking results in larger fragment size and lower solubility, resulting in lower yield (Orlando et al., 1997). Because of the importance of this variable, the shear size of the DNA should be assessed to confirm that the desired degree of fragmentation has been achieved, and it should be reassessed if fixation conditions are altered. The shear size is determined by electrophoresing DNA from step 16 of the Basic Protocol on a 1.5% to 2.0% agarose gel and visualizing with ethidium bromide. A smear of DNA should be apparent with an average size of 500 bp and most of the DNA (>90%) should be in the size range of 100 to 1000 bp. As an alternative to sonication, DNA fragment size can be reduced by treatment of the cross-linked chromatin with micrococcal nuclease. Micrococcal nuclease preferentially cleaves DNA located in the linker re-

gions between nucleosomes. By varying the concentration of micrococcal nuclease, it is possible to generate samples in which average DNA size varies. The minimal useful size is about 150 bp, which corresponds to a mononucleosome. However, cleavage to mononucleosome-sized fragments may also result in a preferential loss of certain genomic regions due to the sequence-specificity of micrococcal nuclease. Immunoprecipitation The success of this procedure relies on the use of an antibody that will specifically and tightly bind its target protein in the buffer and wash conditions used. In addition, antibody should be present in excess with respect to its target protein so that differences in the amounts of the protein-DNA complexes of interest will be accurately measured. Perform preliminary experiments to confirm avid immunoprecipitation and determine an approximate amount of antibody to use. Chromatin extracts should be prepared without prior crosslinking of the cells and subjected to immunoprecipitation with varying concentrations of antibody (20 µg/ml antibody may be a good starting point). The amount of the protein of interest in the extracts before and after immunoprecipitation should be analyzed by immunoblotting (UNIT 6.2) to determine the lowest antibody concentration that depletes >90% of the protein of interest from the extract. This antibody concentration is a good starting point for the full protocol and may later be modified to maximize the signal-to-noise ratio (see Anticipated Results). With cross-linked chromatin, immunodepletion of the target protein is less efficient (∼50%), presumably due to masking or modification of the epitopes, and a significant amount of the protein remains refractory to immunoprecipitation even with higher antibody concentrations. Thus, the ideal antibody concentration is ultimately determined empirically to maximize the yield of specific coprecipitated DNA while minimizing precipitation of nonspecific DNA. Both monoclonal and polyclonal antibodies have been used in this procedure. The monoclonal antibodies 12CA5 (anti-HA), 17D09 (anti-HA), and 9E10 (anti-myc) have been used successfully in different laboratories. In general, triple-HA epitope tags work well (Hecht et al., 1996; Aparicio et al., 1997; Tanaka et al., 1997), and larger multi-myc epitope tags have also been successful (e.g., myc-9, myc-18; Tanaka et al., 1997). Protein G–Sepharose, Protein A–Sepharose, and



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anti-mouse immunoglobulin-coupled magnetic beads have all been used to precipitate the immune complexes, although it should be noted that certain classes of mouse and rat immunoglobulins are not strongly bound by protein A (Harlow and Lane, 1988; see Table 7.2.1). For optimal results, it is critical to minimize the background level of material that inevitably comes down during the immunoprecipitation. The procedures described here work well with a diverse set of antibodies, but it might be necessary to modify the binding and elution conditions in specific cases. Peptide elution is clearly preferred over heat elution, as it is more specific and results in lower experimental backgrounds and hence higher-fold inductions. However, peptide elution is only possible for experiments using antibodies against peptides (typically for analyzing epitope-tagged proteins, but analysis of native proteins should also be possible). In performing peptide elution, it is important to add enough peptide such that the protein-DNA complexes are efficiently eluted from the beads. Another consideration is that the epitope of interest in the chromatin-bound protein might be inaccessible to the antibody due to associated proteins or DNA structures. In such a case, one might obtain a false-negative result. Whereas the majority of a given protein may be efficiently immunoprecipitated from the cross-linked cells, the fraction that is actually cross-linked might be undetectable. The

Determining the Association of Proteins with Specific Genomic Sequences

use of polyclonal antibodies (which often recognize multiple determinants within a protein) or epitope-tagged proteins (the epitope is unlikely to have a specific interaction with other proteins or DNA sequences, particularly if the epitope does not affect the biological function as determined by genetic complementation) minimizes, but does not eliminate this concern. Because of this caveat, negative results should be interpreted cautiously and alternative methods (e.g., in vitro DNA binding or association of the protein with bulk chromatin) should be tried. This concern is particularly relevant when a protein of interest does not appear to interact with any genomic sequence. However, if a protein selectively associates with some genomic sequences, this concern is significantly reduced—i.e., it is unlikely that epitope masking will occur at some loci, but not others. PCR strategy The choice of primers depends on the experimental goals. If binding to a specific site is being tested, a primer pair that flanks the site and at least one control primer pair recognizing a DNA sequence not expected to bind the protein of interest are the minimal requirements (see Fig. 17.7.2). When choosing primers, it is important to remember that resolution between adjacent sequences is limited by the shear size of the DNA. For an average DNA size of 500 bp, a significant fraction of the DNA molecules will be in the 500 to 1000 bp

Figure 17.7.2 Anticipated results from chromatin immunoprecipitation analysis of origin recognition complex (ORC) with replication origin and nonorigin DNA sequences.



range, and hence DNA sequences 1000 bp distal from the actual protein binding site may be coprecipitated. Therefore, primer pairs used as controls should amplify a region of DNA that is far enough away from the expected binding site (e.g., >1 kbp) that coprecipitation of adjacent DNA is not detected. A good strategy is to design multiple sets of primers at increasing distances from a suspected binding site. Such a strategy has also been used to probe the “spreading” and “movement” of proteins on chromatin (Hecht et al., 1996; Aparicio et al., 1997; Strahl-Bolsinger et al., 1997). Success in obtaining high-quality quantitative data is critically dependent on good primer design! In general, primers should be 20 to 30 bases long with a Tm of 55◦ to 60◦ C. Most primers require no purification or special treatment prior to PCR. Amplification products should be 75 to 300 bp. Longer PCR products should be avoided, because the amplification efficiency is substantially lower, and DNA fragments that do not bind to both primers will not be amplified (this can be a significant problem since the size of DNA fragments in the samples averages ∼500 bp and ranges between 100 to 1000 bp). A final primer concentration of 1 µM works well for most primers, but in some instances, improved product specificity may be obtained by lowering the final primer concentration 5 to 10 fold. The design of good primers is greatly facilitated by commercially available software packages such as Oligo 6.6 or Primer Express 1.5. These packages allow for extensive customization of many different parameters, including Tm , oligonucleotide length, GC content, and more. While the success of each individual primer pair in the specific amplification of its target sequence is dependent on many variables, special care must be taken to minimize primer-dimers and hairpins. Finally, it is a good idea to check primers for hybridization to other genomic sequences through the use of a web-based program such as BLAST. Newly obtained primer pairs must be tested for amplification specificity and performance under the conditions that will be used in quantitative PCR. Primer pairs that are suitable for reactions performed by the Basic Protocol might not be suitable for real-time PCR reactions using SYBR Green, because SYBR Green can inhibit Taq polymerase. It is particularly informative to analyze input DNA amplification by the primers in question on highpercentage agarose or polyacrylamide gels after completion of the PCR. The presence of multiple product bands indicates poor speci-

ficity and will invariably lead to unreliable results. For the Basic Protocol, the best test for quality of a given primer pair is to carry out a standard curve using different dilutions of DNA. For a high-quality primer pair, the amount of PCR product should be directly proportional to the amount of DNA, with an error of less than ±20%. The number of PCR cycles is determined empirically. Usually, 25 to 28 cycles is appropriate. More than 28 cycles can result in detection of nonspecifically precipitated sequences and/or lead to variable results due to inactivation of Taq polymerase. Multiple primer pairs can be used in combination if the PCR products are separable by gel electrophoresis (as many as five have been used), but some combinations interfere with efficient amplification of one or more products. It is essential to test primer pairs singly and in combination, with titrations of template DNA, to determine if this is a problem. The advantage of using multiple primer pairs is that individual reactions can generate data for multiple genomic regions in an internally controlled manner. In addition, the Basic Protocol can be used to simultaneously analyze two alleles of a given locus in an internally controlled manner, provided the individual alleles result in different-sized PCR products. When quantitative PCR will be performed in real time using SYBR Green (see Alternate Protocol 2), high-quality primer pairs should result in ∼1.9-fold amplification/cycle. Such amplification efficiency can be determined from quantitative analysis of raw fluorescence data for each cycle. Amplification efficiencies