Q 2011 by The International Union of Biochemistry and Molecular Biology
BIOCHEMISTRY AND MOLECULAR BIOLOGY EDUCATION Vol. 39, No. 4, pp. 309–315, 2011
Laboratory Exercise From Green to Blue: Site-Directed Mutagenesis of the Green Fluorescent Protein to Teach Protein Structure–Function Relationships Received for publication, July 19, 2010, and in revised form, September 10, 2010 Marı´a D. Giro´n and Rafael Salto‡ From the Department of Biochemistry and Molecular Biology II, School of Pharmacy, University of Granada, Campus de Cartuja sn, E-18071 Granada, Spain
Structure–function relationship studies in proteins are essential in modern Cell Biology. Laboratory exercises that allow students to familiarize themselves with basic mutagenesis techniques are essential in all Genetic Engineering courses to teach the relevance of protein structure. We have implemented a laboratory course based on the site-directed mutagenesis of the green fluorescent protein (GFP) from the jellyfish Aequorea victoria. The GFP is ideal because the students are able to correlate the changes introduced into the structure of the protein with the observable modification of its fluorescence properties. By using noncommercial kits, we set up a non PCR-thermocycling reaction using mutagenic primers, followed by removal of the original plasmid template by DpnI digestion. By introducing only one (Y66H) or two mutations (Y66H/Y145F) in the ‘‘cycle 3’’ variant of GFP (F99S, M153T, and V163A) or GFPuv, students are able to analyze the changes from green to blue in the fluorescence emission of the mutated proteins and to correlate these differences in fluorescence with the structural changes using three-dimensional structure visualization software. This inexpensive laboratory course familiarizes the students with the design of mutagenic oligonucleotides, site-directed mutagenesis, bacterial transformation, restriction analysis of the mutated plasmids, and protein characterization by SDS-PAGE and fluorescence spectroscopy. Keywords: Green fluorescent protein, site-directed mutagenesis, structure–function relationship, laboratory course.
Knowledge of the three-dimensional structure of proteins is essential for a detailed understanding of protein functions, because structure–function relationship studies are the basis of Cell Biology. In the School of Pharmacy at the Granada University, Spain, we offer an undergraduate course entitled ‘‘genetic engineering applied to drug design.’’ This includes laboratory exercises to familiarize the student with the concept of structure–function relationship in proteins. The aim of these exercises is to teach the students how to manipulate three-dimensional protein structural models and to analyze, using site-directed mutagenesis, key residues in the protein structure. The green fluorescent protein (GFP)1 from the jellyfish Aequorea victoria is ideal for this purpose, for several reasons. First, GFP possesses the property of emitting light in the green wavelength upon excitation with an appropriate UV
This work was supported by a grant from Fundacio´n Marcelino Botı´n and by University of Granade ‘‘Proyecto de Innovacion Docente 2009-146’’. ‡ To whom correspondence should be addressed. R. Salto. Tel.: (34)-958-246363; Fax: (34)-958-248960. E-mail:
[email protected]. 1 The abbreviations used are: GFP, green fluorescent protein; LB, Luria-Bertani; TAE, tris-acetate-EDTA buffer. This paper is available on line at http://www.bambed.org
light source without the need for any additional substrate or cofactor [1], facilitating the detection and assay of the protein fluorescent activity. Second, numerous mutant forms of GFP with different fluorescence excitation and emission spectra have been produced through mutagenesis, and the threedimensional structures of several mutants have been elucidated, enabling structure–function relationship studies to be carried out. Third, GFP can be easily expressed in bacteria and is structurally very stable, thereby enabling the student to perform phenotype analysis of the mutants produced. Although the changes in spectral properties of GFP usually involve the mutation of several residues in the protein [2], making it nonpractical in a laboratory course for undergraduate students, we propose the implementation of only two mutations, Y66H and Y145F in the ‘‘cycle 3’’ variant of GFP (F99S, M153T, and V163A) or GFPuv with an excitation maximum at 385 nm [3]. The single Y66H mutation in GFPuv is enough to produce a change in fluorescence emission from green (510 nm) to blue (450 nm) and therefore allows students to correlate the phenotype of the mutated protein with the induced structural changes. The additional mutation Y145F further enhances the emission of blue fluorescence, probably either due to the stabilization of the protein folding or the interaction of the mutated amino acid with the chromophore, changing
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DOI 10.1002/bmb.20467
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its protonation state. Thus, this Y145F mutation is suitable for introducing students to handle the models of the wildtype and mutated proteins and correlating the mutations introduced with the new functions. Additionally, students are provided with a mutated plasmid in which Thr 64 has been deleted. As the GFP DT64 lacks any fluorescence, students can confirm that restoration of Thr 64 or the simultaneous introduction of the mutations described above restores green or blue fluorescence, respectively.
TABLE I Timetable for practical course on site-directed mutagenesis Days
Methods
Day 1
Introduction Design of mutagenic oligonucleotides Mutagenic PCR/DpnI digestion DNA electrophoresis of mutagenic reactions Competent cells transformation Bacterial plate visualization Individual colonies; selection and growth in liquid media Purification of plasmid DNA (minipreps) Plasmid DNA digestion with restriction enzymes/DNA electrophoresis of restricted DNA Bacterial crude extract preparation SDS-PAGE Fluorescence spectrum analysis of bacterial crude extracts
Day 2 Day 3 Day 4
COURSE DESIGN
The laboratory exercises presented last 5 days (3 hours daily) and are aimed at students taking the genetic engineering applied to drug design course and who have previously taken a biochemistry and molecular biology course as well as a term of microbiology. During the genetic engineering course, the students are informed about the techniques used in gene engineering and introduced to site-directed mutagenesis methods. Before starting the laboratory exercises, a timetable (Table I) is distributed and safety concerns about the specific laboratory techniques used are explained to the students. Next, the mutations to be assayed by the group are distributed. Students work in pairs. Each student is required to keep a laboratory notebook. This notebook, containing a brief report of the principles, materials and methods, together with the results obtained, and their discussion constitutes 60% of the evaluation. The remaining 40% of the final mark is based on the student’s aptitude, relevance of the student’s questions and answers, as well as an evaluation of their performance and skills during the practical work. MATERIAL AND METHODS
Day 5
Each day comprises 3 hours practical work in the laboratory. Computer-assisted modeling of the mutated proteins; discussion of the results obtained and theoretical background in site-directed mutagenesis and structure-function relationships are taught during the term in addition to the 5 days of laboratory exercises.
Design of Mutagenic Oligonucleotides Mutagenic oligonucleotides, 39–45 bases long, were designed to contain the codon coding for the desired mutation in the central position. To facilitate annealing, primers contained at least 40% GC and also exhibited one or more Gs and Cs at the 50 and 30 ends. Finally, oligonucleotides were designed to include silent mutations that allowed restriction screening of the mutated plasmids. This was accomplished using online software programs such as Sitefind V4.2, available online or as a Java independent application (http://www.utmb.edu/scccb/Software/sitefind.html), or Silent Site Selector for DNA Mutagenesis online software (http://rana.lbl.gov/SSS/). Desalted oligonucleotides were used for mutagenesis without any further purification.
Bacterial Strain and Plasmids The bacteria used in the course are E. coli XL1-blue [endA1 gyrA96(nalR) thi-1 recA1 relA1 lac glnV44 F’[ ::Tn10 proAB1 lacIq D(lacZ)M15] hsdR17(rK2 mKþ)] (Stratagene, Agilent Technologies, Santa Clara, CA). Cells were grown in Luria-Bertani (LB) broth or LB agar at 37 8C. Ampicillin (60 lg/mL) and tetracycline (30 lL/mL) were used as antibiotics. Plasmid pGFPCR [4] carries the cycle 3 variant of GFP (F99S, M153T, and V163A) described by Crameri et al. [3] and is derived from pGFPuv plasmid (Clontech, Mountain View, CA). Either plasmid can be used in this course. The production of the mutated plasmid pGFPCR DT64 is described below.
PCR Mutagenic Reaction Site-directed mutagenesis of the pGFPCR plasmid used is based on a non-PCR-thermocycling reaction using mutagenic primers, followed by removal of the original plasmid template by DpnI digestion. The mutagenic primers are shown in Table II. For the mutagenesis reactions, commercial kits are not used. The PCR reactions are prepared according to Laible and Boonrod [5]: plasmid template 60 ng, the selected pair of mutagenic oligonucleotides (0.4 lM final concentration each), 0.25 mM nucleotides triphosphates, 20 mM Tris-HCl (pH 8.8 at 25 8C), 10 mM (NH4)2SO4, 10 mM KCl, 0.1 mg/mL bovine serum albumin,
TABLE II Oligonucleotides used in the course Name DT64 Green Y66H Y145F
Secuence 0
0
5 -CCA ACA CTT GTC ACT --- TTC TCT TAT GGT GTT CAA TGC-3 (Forward) 50 -GCA TTG AAC ACC ATA AGA GAA --- AGT GAC AAG TGT TGG-30 (Reverse) 50 -CCA ACA CTT GTC ACT ACT TTC TCT TAT GGT GTT CAA TGC-30 (Forward) 50 -GCA TTG AAC ACC ATA AGA GAA AGT AGT GAC AAG TGT TGG-30 (Reverse) 50 -CCA ACA CTT GTC ACT ACT TTC AGC CAT GGT GTT CAA TGC-30 (Forward) 50 -GCA TTG AAC ACC ATG GCT GAA AGT AGT GAC AAG TGT TGG-30 (Reverse) 50 -GGA CAC AAG CTT GAG TAC AAC TTT AAC TCA CAC AAT GTA TAC ATC-30 (Forward) 50 -GAT GTA TAC ATT GTG TGA GTT AAA GTT GTA CTC AAG CTT GTG TCC-30 (Reverse)
Mutated nucleotides are underlined.
Mutation
Restriction
DT64
none
none
none
Y66H
NcoI
Y145F
HindIII
311 0.1% (v/v) Triton X-100, 2 mM MgSO4, and 2.5 U of recombinant Pfu DNA Polymerase (Fermentas GmbH, St. Leon-Rot, Germany) in a final volume of 50 lL. PCR conditions were as follows: an initial denaturation phase (30 sec, 95 8C), followed by 18 cycles of denaturation (30 sec, 95 8C), hybridization (30 sec, 55 8C), and extension (4 min, 72 8C). After cycling, PCR was programmed for an additional extension of 10 min at 72 8C followed by cooling to 4 8C. Once the thermocycling reaction was completed, an aliquot of 10 lL was reserved for agarose electrophoresis analysis and 1 lL (10 U) of DpnI enzyme was added to the rest of the reaction and incubated at 37 8C for at least 2 hours to remove the plasmid template. To confirm that a successful mutagenic reaction has been produced, students check the amplification efficiency by running 10 lL of the amplification reaction before and after DpnI digestion, as well as the equivalent amount of template used for the mutagenic reaction, in a tris-acetate-EDTA buffer (TAE) 0.8% agarose gel (Fig. 2a).
Transformation of Competent Cells and Visualization of Colonies Containing Mutated Plasmids Students are provided with XL1-blue competent cells obtained in our laboratory by the method of Nishimura et al. [6]. For transformation, 100 lL competent cells aliquots were thawed and incubated on ice with 5 lL of the mutagenic reaction for 1 hour. Cells were heat shocked for 2 min at 42 8C and then kept on ice for an additional period of 10 min. Next, 900 lL of liquid LB media without antibiotics were added to the transformed cells. These were incubated at 37 8C for 1 hour to induce antibiotic resistance. Finally, 250 lL of the transformed bacteria were plated on selective (ampicillin plus tetracycline) agar media and allowed to grow at 37 8C overnight. Next morning, plates were cooled to 4 8C to facilitate GFPuv folding and therefore to enhance its fluorescence and were examined under an UV transilluminator (365 nm) for the presence of colonies expressing wild-type or mutated GFPuv.
Plasmid DNA Purification and Restriction Analysis of Mutated Plasmids Selected colonies were picked and grown in 3 mL liquid LB media plus antibiotic overnight for plasmid preparation the next day. This was carried out using the alkaline lysis method (without phenol:chloroform extraction) using standard protocols [7]. Purified plasmid was digested with the corresponding restriction enzyme (Fast Digest NcoI for the mutants at position 66 and Fast Digest HindIII for the mutants at position 145) according to the manufacturer (Fermentas) for 15 min and analyzed by TAE agarose electrophoresis (Figs. 2b and 2c).
Expression of GFPuv Mutants and SDS-PAGE and Fluorescence Characterization Colonies of the desired fluorescence were picked from agar plates containing bacteria transformed with wildtype or mutated pGFPCR plasmid and allowed to grow overnight at 37 8C to saturation. Although the GFP gene
is driven by the Lac promoter, its basal level of expression is great enough to allow detection of the protein in the absence of isopropyl-D-thiogalactopyranoside, which was therefore not added. Cell density was quantified by turbidity at 600 nm. Cell aliquots were pelleted by centrifugation and resuspended in lysis buffer containing SDS and boiled or not for 4 min. Samples were electrophoresed on 12% SDS-PAGE and photographed under 365 nm UV light using a transilluminator, as described above, and later Coomassie-blue stained. For the determination of the fluorescence spectra of GFPuv mutants, cell cultures from bacteria transformed with the wild-type or mutated pGFPCR plasmids were allowed to grow overnight, cell density was normalized and bacteria pelleted by centrifugation. Cells were resuspended in PBS and disrupted by sonication. Cell debris was removed by centrifugation and crude extract spectra were recorded in a Shimadzu RF-5301PC spectrofluorophotometer. Emission spectra were recorded using an excitation wavelength of 380 nm. Excitation spectra were measured using the maximum emission wavelength for each mutant.
Analysis of the Structure of the Mutants Analysis and handling of the crystallographic models of GFP variants was carried out by students using the Swiss-PdbViewer/DeepView software [8]. The pdb files used were 1B9C (GFPuv) [9], 2EMD (GFPuv Y66H) [10], and 1BFP (GFPuv Y66H/Y145F) [11]. RESULTS AND DISCUSSION
The implementation of laboratory exercises that familiarize students with the structure–function relationships in proteins by means of site-directed mutagenesis is mandatory in courses involving genetic engineering or advanced drug design. However, these practical exercises are sometimes time-consuming, and the results obtained by the students are often unsatisfactory. There are two main reasons for this, technical and economical. First, there are the difficulties involved in the different site-directed mutagenesis techniques. Uracyl-laden single-stranded DNA or the Kunkel mutagenesis method involve a cumbersome step to obtain single-stranded DNA and are not well suited for practical teaching. Mutagenesis techniques based on the joining of PCR fragments obtained from two PCR amplifications using two central mutated complementary oligonucleotides and two flanking oligonucleotides are time-consuming and require the posterior ligation of the fragment in the appropriate vector (which is usually complicated for the student and has a very low ligation efficiency). Finally, the methods based on a non-PCR thermocycling amplification using mutated oligonucleotides followed by the DpnI digestion of the template are commercially available (e.g. Stratagene’s QuikChangeTM Site-Directed Mutagenesis Kit), but expensive. The second problem in teaching site-directed mutagenesis is that the students are unable to fully comprehend that the changes introduced in the genotype have a clear correspondence with the phenotype or properties of the mutated protein. Subtle changes in the kinetic
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FIG. 1. Primary and secondary structure of GFPuv protein. (a) Scaffolding of GFPuv protein according to a secondary structure prediction [13]. Mutations are indicated by open circles for positions 66 and 145, producing a blue fluorescent protein, and a closed circle for a deletion in position 64, producing a nonfluorescent protein. (b) Amino acid primary sequence of GFPuv. The three SYG residues generating the chromophore are in bold. The positions that had been mutated are underlined.
properties of an enzyme or the introduction or removal of restriction sites tend not to be properly evaluated by the students on an undergraduate course. Here, we present a laboratory exercise intended to solve these problems. Site-directed mutagenesis is carried out using a non-PCR thermocycling amplification followed by the DpnI digestion. For this purpose, instead of a commercial kit, we use recombinant Pfu polymerase and DpnI restriction enzyme [5]. The system is robust enough for undergraduate students and most, if not all, of the students are able to obtain successful mutagenic reactions that can be verified by agarose gel electrophoresis (Fig. 2a). Also, this electrophoresis helps the students to notice that the DnpI digestion removes the plasmid template and reduces the background of this mutagenesis technique. For the transformation of bacteria with the mutated plasmids, we use competent cells obtained in our laboratory by the method of Nishimura et al. [6]. These competent cells are inexpensive to obtain compared with commercial sources and still have enough transformation efficiency to be used in the site-directed mutagenesis
technique, because, with the transformation of an aliquot of the mutagenesis reaction, the students usually obtain 50–100 colonies per plate. The selection of GFPuv protein as template for the site-directed mutagenesis allows the students to appreciate the power of the technique. Students are provided with the plasmid pGFPCR that encodes for the cycle 3 variant of GFP (F99S, M153T, and V163A) described by Crameri et al. [3] or GFPuv (Fig. 1) and is ideal for detection using a 365 nm transilluminator. The GFPuv monomer has 238 amino acids. The chromophore responsible for the autofluorescence is autocatalytically formed from residues Ser 65, Tyr 66, and Gly 67. The protein has a maximum excitation peak at 380–395 nm, whereas the emission wavelength is 508 nm [1]. Alternatively, students are provided with a plasmid that carries a deletion in Thr 64 (Fig. 1). This deletion produces a protein lacking any fluorescence. The restoration of the Thr at position 64 or the introduction of mutations Y66H or Y66H/Y145F has a dramatic effect on the fluorescent properties of the mutated protein because green fluorescence is restored, or, more importantly, green fluorescence is shifted to
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FIG. 2. Mutagenesis of plasmid pGFPCR coding for GFPuv protein. (a) Electrophoresis on 0.8% agarose of the PCR mutagenic reaction. PCR reaction was carried out using plasmid pGFPCR (60 ng) as a template and forward and reverse oligonucleotides corresponding to the mutation Y66H, as described in the Material and Methods section. Lane 1, plasmid template used for the PCR reaction; lane 2, PCR reaction; lane 3, PCR reaction after digestion with DpnI enzyme. Arrows mark the bands corresponding to the amplified mutated product (a), plasmid template (b), and free oligonucleotides (c). (b) Restriction analysis of plasmid isolated from individual colonies obtained after transformation of competent XL1-blue cells with the product of mutagenesis reactions. The digestion of the mutant pGFPCR Y66H should produce a 325 bp band, whereas the wild-type pGFPCR generates a 354 bp band. In the double mutant, pGFPCR Y66H/Y145F, the digestion with HindIII produces a 481 bp, whereas in plasmids lacking the Y145F mutation, the digestion with HindIII linealizes the plasmid. In the two examples obtained by students, sample 1 corresponds to a wild-type plasmid and samples 2–7 correspond to plasmids with the desired mutation.
blue [12]. Additionally, the mutation Y145F produces a significant increase in the blue fluorescence [1]. All these changes can be detected in the cell colonies (Fig. 3a). In fact, students are readily able to distinguish background colonies transformed with the template plasmid from those transformed with the mutated plasmid by the brightness or fluorescence color. Thus, it would not be necessary to introduce silent mutations that incorporate or remove a restriction site in the mutagenic oligonucleotides to detect mutations (Table II, Fig. 2b). However, we consider that for the student, the design of oligonucleotides containing silent mutations that alter the restriction pattern has a clear didactic purpose and should therefore be included. This also allows students to carry out plasmid minipreps, digest plasmidic DNA, and identify the mutated plasmid by agarose gel electrophoresis. The
new ‘‘fast digest’’ enzymes (Fermentas), which perform 15-min digestions, enable students to easily follow the course timetable. GFP has a very strong secondary and tertiary structure, which allows the protein to retain fluorescence in the presence of high concentrations of SDS if it is not heated. This property permits the analysis of the mutants by SDS-PAGE (Fig. 3b). Furthermore, comparison of the fluorescence and mobility of boiled and not boiled samples (Fig. 3b) enables students to learn concepts related to protein denaturation and stability. Finally, for the characterization of the mutated proteins, excitation and emission fluorescence scans of bacterial crude extracts are carried out (Fig. 3c). By using these fluorescence determinations, students become familiar with basic fluorescence spectroscopy and thus begin to comprehend concepts
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BAMBED, Vol. 39, No. 4, pp. 309–315, 2011 that highlight the relevance that changes in protein structure have on protein properties. Finally, the experiments are complemented by the study of the structures of the mutated proteins. The pdb files corresponding to these mutated proteins are available [9–11], and the students can familiarize themselves with secondary structure prediction software [13] and the analysis of three-dimensional structures using the SwissPdbViewer/DeepView software [8]. The relevance of the influence of the mutations on the ionization state of the chromophore and protein folding (Fig. 4) are discussed with the students to encourage them to relate structure and functions in this versatile protein. LABORATORY EXERCISES ASSESSMENTS
FIG. 3. Expression of GFPuv mutants. (a) Bacterial colony fluorescence of transformed bacteria harboring mutated pGFPCR plasmids. Competent XL1-blue cells were transformed with wild type or mutated pGFPCR plasmids and allowed to grow overnight at 37 8C on selective LB agar plates. Colonies were visualized using a 365 nm UV transilluminator. Plates were photographed using a UV filter, maintaining constant exposure settings. (1) Mutant pGFPCR DT64; (2) Wild-type pGFPCR; (3) Mutant pGFPCR Y66H; (4) Mutant pGFPCR Y66H/Y145F. (b) SDS-Poliacrylamide gel electrophoresis of crude extracts from transformed bacteria harboring mutated pGFPCR plasmids. Samples were electrophoresed on 12% SDS-PAGE and photographed under 365 nm UV light (left panel) and Coomassie-blue stained afterward (right panel). (1) Mutant pGFPCR DT64; (2) Wild-type pGFPCR; (3) Mutant pGFPCR Y66H; (4) Mutant pGFPCR Y66H/Y145F. (5) Sample corresponding to wildtype pGFPCR plasmid that had been boiled before loading. Arrows a and b show the mobility shift of the band corresponding to GFPuv due to the partial (no boiling) or complete (boiling) denaturalization of the protein. (c) Fluorescence spectra of bacterial crude extracts containing GFPuv mutant proteins. Cells were resuspended in PBS and disrupted by sonication. Cells debris was removed by centrifugation and crude extract spectra were recorded in a Shimadzu RF-5301PC spectrofluorophotometer. Emission spectra (solid lines) were recorded using an excitation wavelength of 380 nm. Excitation spectra (dotted lines) were measured using the maximum emission wavelength for each mutant.
These laboratory exercises have been carried out during the past 5 years by a total of 136 students. As stated in the course design section, at the end of the laboratory exercises, students were evaluated according to their ability to perform the exercises presented in the course, the results obtained and their responses to relevant questions. Also, at the end of the practical course, the students responded to a questionnaire to assess their laboratory skills, the utility of the laboratory practices and their evaluation of the tasks covered by these exercises. Finally, the students were asked to suggest improvements to the laboratory exercises. Our results show that 94% of the students who enrolled on the course were able to perform the laboratory exercises correctly. In the skill survey carried out at the end of the course, 77% of the students declared that they considered themselves capable of performing basic site-directed mutagenesis and protein engineering experiments independently. This result represents a very significant increase in student perception of their confidence to carry out these studies by themselves, because, at the beginning of the course, only 37% of the students considered themselves equipped to carry out these assignments independently. Asked about the most relevant aspects of the laboratory course, all the students remarked on the straightforward analysis of structure–function relationship in GFP due to the changes in the phenotype of the mutated proteins. Furthermore, most of the students (97%) were impressed by their ability to restore green fluorescence in the plasmid that carries a deletion in Thr 64 by simple mutagenesis techniques. Another highly valued aspect (78% of the students) was the technically uncomplicated mutagenesis method used, which showed them the power of this technique to manipulate protein structure. In addition, the fact that mutated proteins retained fluorescence in the SDS-PAGE was valued positively as a learning tool by practically all the students. Finally, improvements to the laboratory exercises proposed by the students include dedicating more time to the analysis of crystal structure models of the mutated proteins (proposed by 59% of the students). Also, 44% proposed to include the purification by chromatography and further characterization of the mutated proteins produced in the exercises. Unfortunately, time constraints in the course have prevented us from incorporating these interesting suggestions.
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FIG. 4. Crystal structure of GFPuv mutants. Models corresponding to the (a) wild-type, (b) Y66H, and (c) Y66H/Y145F GFPuv structures are shown. These models are based on the 1B9C, 2EMD,’ and 1BFP PDB models, respectively, and are depicted using the SwissPdbViewer/DeepView software [8]. Side chains of residues 145 and 148, as well as the chromophore group in the crystal structures, are shown to highlight the relevance of these residues in the protein fluorescence properties. (d) Relative position of the chromophore and amino acids 145 and 148 in the pGFPuv crystals models: wild-type, green, GFPuv Y66H, yellow, and GFPuv Y66H/Y145F blue.
We conclude that based on both the students’ questionnaires and our own teaching experience, the incorporation of dramatic changes in the phenotype of the mutated proteins, the trouble-free mutagenesis techniques, the well known three-dimensional structure of GFP and its mutants, as well as the chemical stability of the GFP, are the most relevant aspects of the laboratory exercises proposed. Acknowledgments— We thank Javier Lo´pez-Jaramillo for expert assistance with the three-dimensional protein models and for helpful discussions. We also want to thanks all our students from the Genetic Engineering applied to drug design course at the School of Pharmacy, University of Granada, Spain. We thank Robert Abrahams, BSc, for checking the manuscript. REFERENCES [1] R. Heim, R. Y. Tsien (1995) Engineering green fluorescent protein for improved brightness, longer wavelengths and fluorescence resonance energy transfer, Curr. Biol. 6, 178–182. [2] J. D. Pe´delacq, S. Cabantous, T. Tran, T. C. Terwilliger, G. S. Waldo (2006) Engineering and characterization of a superfolder green fluorescent protein, Nat. Biotechnol. 24, 79–88. [3] A. Crameri, E. A. Whitehorn, E. Tate, W. P. Stemmer (1996) Improved green fluorescent protein by molecular evolution using DNA shuffling, Nat. Biotechnol. 14, 315–319.
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