Identifying ectomycorrhizal fungi

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Jun 22, 2007 - Extracting mycorrhizal tips from field samples . ..... 10.00 - 12.00: Compiling and analyzing molecular data. 12.00 - 13.00: Lunch ..... reagent (a aqueous solution of iodine, KI, and chloral hydrate) on a slide and mix in a small ...... Ordinate the data and graph the output as described in the instruction sheet.
Identifying ectomycorrhizal fungi - from environmental samples to DNA sequences A NordForsk - research training course

COURSE

COMPENDIUM

RASMUS KJØLLER JERI PARRENT ANDY TAYLOR TRUDE VRÅLSTAD

WITH CONTRIBUTIONS FROM:

ELLEN LARSON, URMAS KÕLJALG, IAN ALEXANDER & TROND SCHUMACHER

AUGUST 12TH – 24 TH 2007-06-22 HOSTED BY BIOLOGICAL INSTITUT, UNIVERSITY OF COPENHAGEN

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Content Preface and acknowledgements .............................................................................................. 4 Course at a glance .................................................................................................................... 5 Detailed time table.................................................................................................................... 6 Part 1 – Field course at Sorø Field Station .......................................................................... 10 Sunday August 12th ............................................................................................................. 10 Monday and Tuesday August 13 – 14th............................................................................... 10 General comments on collecting fungal sporocarps ........................................................ 10 Additional notes for collecting sporocarps of resupinate thelephoroid fungi .................. 11 Additional notes for collecting sporocarps of Ascomycetes............................................ 12 General comments for annotation of collections voucher specimens, DNA-samples and sequences.......................................................................................................................... 14 General comments for preparation of DNA samples ....................................................... 16 Student exercises: Describing sporocarps ........................................................................ 16 1: Agarics – general features and specific exercises .................................................... 17 1A: Describing Russula sporocarps ............................................................................. 17 1A: Describing Russula sporocarps ............................................................................. 18 1B: Describing Lactarius sporocarps........................................................................... 24 1C: Describing and identifying boletoid sporocarps.................................................... 27 2: Describing apothecial sporocarps (ascocarp)........................................................... 30 3: Describing resupinate thelephoroid sporocarps ....................................................... 32 Wednesday August 15th....................................................................................................... 42 Characterisation of ectomycorrhiza ................................................................................. 42 Background to ectomycorrhizal characterisation ......................................................... 42 1. Extracting mycorrhizal tips from field samples ....................................................... 44 2. Morphotyping........................................................................................................... 44 3. Mantle preparations.................................................................................................. 45 5. Microscopic features ................................................................................................ 46 6. Preparing tips as vouchers specimens and for molecular analysis........................... 47 Notes on Rhizomorphs ................................................................................................. 47 Notes on Emanating Hyphae........................................................................................ 48 Notes on Cystidia ......................................................................................................... 49 Notes on Mantle structures........................................................................................... 49 Thursday August 16 ............................................................................................................. 51 Sampling ectomycorrhizal communities along an environmental gradient. .................... 51 Sorting cores................................................................................................................. 51 Analysing the data........................................................................................................ 53 Friday August 17th............................................................................................................... 55 Virtual mycorrhizas – a sampling exercise ...................................................................... 55 Saturday August 18th ........................................................................................................... 55 Part 2 – Molecular lab course at University of Copenhagen.............................................. 56 Sunday August 19th ............................................................................................................. 56 DNA extraction using the CTAB-chloroform-isopropanol extraction method (adapted from Gardes et al. 1996) – description of the method...................................................... 56 Reagents and equipment needed for the CTAB extraction .............................................. 57 DNA extraction using the CTAB-chloroform-isopropanol extraction method - protocol59 Polymerase Chain Reaction (PCR) - Principles............................................................... 59 Primers for amplifying and sequencing fungal rDNA ITS region ................................... 61 PCR Protocol for a 30 µl setup ........................................................................................ 63 Monday August 19th............................................................................................................ 63

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Principles for electrophoresis ........................................................................................... 63 Student exercise: Electrophoresis - Checking the PCR result.......................................... 64 Protocol for detection of PCR-products ............................................................................... 65 Purifying PCR products and measuring the DNA content............................................... 66 Student exercise: Getting your PCR products ready for sequencing ............................... 66 Sequencing of PCR Products using MWG Biotech ......................................................... 66 Tuesday August 21th ............................................................................................................. 67 Principles for DNA sequencing........................................................................................ 67 Wednesday August 22th........................................................................................................ 70 Student exercise: Post – molecular work / computer lab ................................................. 70 Thursday August 23th .......................................................................................................... 77 Student exercise: Identification of sequences .................................................................. 79 Identification of sequences using UNITE .................................................................... 79 Identification using FasTa3.......................................................................................... 80 Relevant web-sites for sequence analysis ........................................................................ 81 Friday August 24th ................................................................................................................ 81

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Preface and acknowledgements The Nordic-Baltic NorForsk (previously NorFa) research network "Identification and Ecology of Ectomycorrhizal fungi" (http://www2.dpes.gu.se/project/unite/) was established in January 2003 and will end by the end of this year. The network was coordinated 2003-2006 by Susanne Erland (Lund University, Sweden) and in 2007 by Andy Taylor (SLU, Uppsala, Sweden). The network comprises scientists from Sweden, Finland, Denmark, Estonia, Norway and Scotland in the fields of taxonomy and ecology of ECM fungi. A major goal for the Network is to organize integrated and specialized Norforsk research training courses for PhD-students and Post docs in mycology, and especially for those with an interest in ectomycorrhizal topics. The current course "Identification of ectomycorrhizal fungi - from environmental samples to sequences" is the second research training course coming out of this initiative, the first was run in 2004 and was hosted by University of Oslo (Trude Vrålsatd). The present course is therefore the second specialized course in the Nordic region that offers practical and theoretical teaching in the field and lab on sampling approaches and analyses, morphological recognition of sporocarps and ECM as well as molecular techniques and analyses for ECM species identification. The organizing group of this course consists of Rasmus Kjøller and Thomas Læssøe (University of Coepnhagen, Denamrk), Andy Taylor and Jeri Parrent (Swedish University of Agricultural Sciences, Uppsala, Sweden) and Ian Alexander (University of Aberdeen, Scotland). Other people contributing to the course as teachers and lecturers include Reinhard Agerer (Ludwig-Maximilian-University, Munich, Germany), Urmas Kõljalg (University of Tartu, Estonia), Ian Anderson (The Macaulay Institute, Aberdeen, Scotland), Sari Timonen (University of Helsinki, Finland – not confirmed yet) and Taina Pennanen (Metla, Helsinki, Finland – not confirmed yet). All organizers, teachers and lecturers are gratefully thanked for their contribution. NorFa is financially supporting this course with 357 000 NOK. The Biological Intitute at the University of Copenhagen is hosting the course and allocating invaluable resources, facilities and equipment. We also thank Anette Hørdum Løth and Lis Mathorne for technical assistance and Helle Bek Mikaelsen for taking care of the economy. This course will briefly cross through some of the different ectomycorrhizal research fields, and we intend to offer you a taste from some of the methods applied. However, it will not be possible to touch or learn all aspects within these two intensive weeks. In the first part of the course, we will be focusing on basic principles for sampling of scientific collections, annotations and descriptions of sporocarps and ECM roots, rather than on species diversity and general ECM species knowledge. The second part will briefly touch DNA extraction, PCR, sequencing and molecular identification of ECM fungi. Instead of filling your baskets with all the ECM fungi and roots you can find, you should make a few thorough collections that will be your particular responsibility, and you should follow these up through all the processes from field to lab. On behalf of the organizers and teachers, I hope this course will provide you with valuable knowledge, friends and contacts that may help you in your further research development. Rasmus Kjøller, Copenhagen June 29th

19. DNA extraction PCR Gel electrophoresis

12. Arrival, installation and WELCOME

Sunday

14. Sampling of sporocarps Identification of sporocarps

21. Lectures, student presentations and sightseeing in Copenhagen

Identification of sporocarps

20. Gel electrophoresis Preparing for sequencing

Tuesday

13. Sampling of sporocarps

Monday

22. Student presentations, Computer lab.

Morphotyping

15. Sampling of root tips

Wednesday

23. Lectures, Computer lab. Social evening

16. Sampling of ectomycorrhizal communities Morphotyping and counting

Thursday

Course at a glance

24. Integrating molecular and morphotype based data END OF COURSE!

17. Compilation of community data Analysis of community data Truffle excursion PARTY !

Friday

25.

18. End of field part Packing and RELOCATING TO COPENHAGEN!

Saturday

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Detailed time table Teachers and lecturers on the course: Name (Abbr.) Institution Reinhard Agerer (RA) Ludwig-Maximilians-Universität München, Germany Ian J Alexander (IJA) University of Aberdeen, Scotland Ian Anderson (IA) The Macaulay Institute, Aberdeen, Scotland Rasmus Kjøller (RK) University of Copenhagen, Denmark Urmas Kõljalg (UK) University of Tartu, Estonia Thomas Læssøe (TL) University of Copenhagen, Denmark Jeri L Parrent (JLP) Swedish University of Agricultural Sciences, Uppsala, Sweden Taina Pennanen (TP) Finnish Forest Research Institute – not confirmed Andy F S Taylor (AT) Swedish University of Agricultural Sciences, Uppsala, Sweden Sari Timonen (ST) University of Helsinki, Finland – not confirmed

The field course - Sorø (part 1) Sunday August 12th 16.00: Arrival, registration, installation and accommodation at Sorø field station 17.00 - 18.00: Dinner and welcome (RK) 18.00 - 18.45: Introductory lecture: Ectomycorrhizal fungi: pivotal organisms in ecosystems (IA) Monday August 13th 08.00 - 08.30: Breakfast 08.30 - 09.00: Introduction to the day's activities (UK, TL, AT) 09.00 - 12.00: Fieldwork: Sampling of sporocarps from ectomycorrhizal fungi (UK, TL, AT) 12.00 - 13.00: Lunch 13.00 - 16.00: Labwork: Identification and preparation of sporocarps for mol. analysis (UK, TL, AT) 16.00 - 17.00: Free time 17.00 - 18.00: Dinner 18.00 - 18.45: Lecture: Phylogenetic diversity of ectomycorrhizal fungal lineages (TL) 19.00 - 19.45: Lecture on selected fungal groups I: Russulaceae (TL) 20.00 - 22.00: Labwork: Identification and preparation of sporocarps for mol. analysis (UK, TL, AT) Tuesday August 14th 08.00 - 08.30: Breakfast 08.30 - 09.00: Introduction to the day's activities (UK, TL, AT) 09.00 - 12.00: Fieldwork: Sampling of sporocarps from ectomycorrhizal fungi (UK, TL, AT) 12.00 - 13.00: Lunch 13.00 - 16.00: Labwork: Identification and preparation of collections for molecular analysis (UK, TL, AT) 16.00 - 17.00: Free time 17.00 - 18.00: Dinner

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18.00 - 18.45: Lecture on selected fungal groups II: Boletes (AT) 19.00 - 19.45: Lecture on selected fungal groups III: Tomentelloid (UK) 20.00 - 22.00: Labwork: Identification and preparation of collections for mol. analysis (UK, TL, AT) Wednesday August 15th 08.00 - 08.30: Breakfast 08.30 - 09.00: Introduction to the day's activities (RA) 09.00 - 12.00: Fieldwork: Collection of ectomycorrhizal root tips for morphotyping (RA) 12.00 - 13.00: Lunch 13.00 - 13.45: Lecture: Anatomical characters of ectomycorrhizas (RA) 14.00 - 16.00: Labwork: Extraction and analysis of root material (RA,UK, AT) 16.00 - 17.00: Free 17.00 - 18.00: Dinner 18.00-22.00: Labwork and lectures: Morphotyping and preparation of root tips for molecular analysis (RA, IJA, RK, AT) Thursday August 16th 08.00 - 09.00: Breakfast 09.00 - 09.45: Lecture: Sampling protocols, spatial & temporal variation (IJA) 09.45 - 10.00: Introduction to the day's activities (RK, AT) 10.15 - 12.00: Fieldwork: Sampling of ectomycorrhizal communities, root tips, mycelia and soil samples, upon return, prepare samples for labwork (RK, AT) 12.00 - 13.00: Lunch 13.00 - 16.30: Labwork: Extraction, morphotyping and counting ectomycorrhizas (RA, AT) 16.30 - 17.00: Free 17.00 - 18.00: Dinner 18.00 - 18.45: Lecture: Ectomycorrhizal communities: Root tips, sporocarps, mycelia and resistant propagules – four views of the ectomycorrhizal community (RK) 19.15 - 22.00: Labwork continues Friday August 17th 08.00 - 09.00: Breakfast 09.00 - 09.45: Lecture: Multivariate analysis (IJA) 10.00 - 12.00: Analysis of mycorrhizal community data I: Compilation of class data for analysis (IJA, AT) 12.00 - 13.00: Lunch 13.00 - 17.00: Analysis and interpretation of mycorrhizal community data II: Sampling exercise - virtual mycorrhizas (IJA, AT) 17.00 - 19.00: Excursion to either (a) the Danish CORE field site in Lille Bøgeskov, Sorø (measuring of forest carbon fluxes), or (b) Truffle demonstration 19.00 - 20.00: Dinner 20.00: Party Saturday August 18th 09.00 - 10.00: Breakfast 10.00 - 10.30: Student feedback on the field course

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11.00 - 12.00: Lecture: Introduction to molecular methods in mycorrhizal research (RK) 12.30 - 13.00: Lunch Afternoon: Packing of equipment and samples, closure of the lab. Departure from Sorö and arrival in Copenhagen, accommodation of students - Evening free ---------------------------------------------------------------------

The laboratory course – Copenhagen (part 2) Sunday August 19th 09.00 - 930: Welcome to the University of Copenhagen 09.30 - 10.30: Lab work: Begin DNA extractions from sporocarps, root-tips, mycelia and soil samples (RK, JLP) 10.45 - 11.15: Lecture: Practice and Principles of DNA extraction (JLP) 11.30 - 13.00: Lab work: DNA extractions, continued 13.00 - 14.00: Lunch 14.00 - 17.00: Lab work: Extractions continued 17.00 - 17.30: Lecture A practical introduction to PCR (JLP) 17.30 - 18.30: Dinner 18.30 - 21.00: Lab work: PCR from from sporocarps, root-tips, mycelia and soil samples (RK, JLP) Monday August 20th 09.00 - 12.30: Lab work: Gel electrophoresis of PCR products and selection of products for sequencing (RK, JLP) 12.30 - 13.30: Lunch 13.30 - 16.00: Preparing PCR products for sequencing (RK, JLP) 16.15 - 17.15: Free 17.30 - 18.30: Dinner 18.45 - 19.15: DNA sequencing – principles (RK) Tuesday August 21th 09.00 - 10.30: Molecular methods: identification of environmental samples- TRFLP, DGGE, Cloning - principles and practice (JLP) 10.45 - 11.45: Presentation of students own PhD/Post doc projects (4 projects) 12.00 - 16.00: Sightseeing in Copenhagen including lunch 16.30 - 17.30: Presentation of students own PhD/Post doc projects (4 projects) 17.45 - 18.45: Dinner 19.00 - 20.00: Presentation of students own PhD/Post doc projects (4 projects) Wednesday August 22th 09.00 - 09.45: Presentation of students own PhD/Post doc projects (4 projects) 10.00 - 12.30: Computer lab: sequence analysis 12.30 - 13.30: Lunch 13.30 - 17.30: Computer lab work continue 17.30 - 18.30: Dinner 18.45 - 20.30: Computer lab work continue

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Thursday August 23th 09.00 - 10.00: Lecture: Phylogenetic identification of mycorrhiza (UK) 10.15 - 12.30: Computer lab 12.30 - 13.30: Lunch 13.30 - 16.30: Computer lab 16.30 - 17.30: The Mycorrhizal Community: The Hubble Telescope Perspective (AT) 19.00: Social evening including dinner Friday August 24th 10.00 - 12.00: Compiling and analyzing molecular data. 12.00 - 13.00: Lunch 13.00 - 15.00: Final discussion and course evaluation 15.00: Departure

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Part 1 – Field course at Sorø Field Station Sunday August 12th The day of arrival is rather relaxing (in opposite to the rest of the course), but you should try to spend some time reading the course compendium. You will need to know some of what is written there before you are going out in the field!

Monday and Tuesday August 13 – 14th Our focus for Monday and Tuesday is on ectomycorrhizal sporocarps – recognition and collection in the field, and determination in the lab. You will have been allocated a work partner – it is a good idea to work with your partner in the field as well as the lab – two pairs of hands and two minds working together are usually more effective (and a lot more fun) than working on your own. For these two days, the lab will be split into two. On Monday, one half will deal with agarics and the other half will concentrate on thelephoroid fungi and ascomycetes. You will switch fungal groups on Tuesday.

General comments on collecting fungal sporocarps Your main objective is to obtain ‘collections’ of ECM fungi that are sufficiently complete for you to identify them in the laboratory. The term ‘collection’ refers to one or more sporocarps of a single species, with all sporocarps originating from a common mycelium. Obviously, it is impossible (without some detailed analyses) to say exactly which sporocarps have a common mycelium but, in general, those growing close together and which look similar are regarded as a single collection. If you are in doubt, it is best to treat the sporocarps as different collections. Your collections should ideally consist of sporocarps at all stages of development. This is because identification keys often ask about characteristics that are only found on very young or old sporocarps. In the field you will need: field notebook, basket, aluminium foil, small plastic boxes with lids, Eppendorf tubes, small ruler, pencil and paper, permanent marker pen, knife, hand lens. Optional – camera and tripod. IMPORTANT: 1) It is much better to have only a few good collections, each consisting of several sporocarps that have been photographed and well documented than having lots of collections comprising single sporocarps with poor notes. 2) Each collection must be assigned a unique code that can be used to distinguish it from all other collections. 3) It is essential that you collect entire sporocarps – pay particular attention to the base. Place a knife under the base of the sporocarp and gently remove the whole sporocarp from the substrate.

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4) Handle the sporocarps as little as possible – features essential for identification (cystidia on the stipe surface) are easily removed by touching and replaced by fingerprints! Lift sporocarps either by the soil-covered base or by placing your thumb on the base of the stipe and a finger or two on the cap. General comments 1) When you first find a potential collection, relax, take your time and spend a few moments looking around the immediate area where the ‘obvious’ sporocarps are situated. Try and locate ALL of the other potential members of the collection before you start working with the collection. Sporocarps can be very difficult to see – even those with bright colours. Remember: try to get all stages of development. 2) Once you have satisfied yourself that all sporocarps have been detected, decide which ones would be good to form the centrepiece of your photograph – if you don’t have a camera, try and get someone else to photograph your collection. Arrange your collection for the photograph. Remember: carefully remove each sporocarp you want to move and touch each one as little as possible. The photograph should illustrate as many features of the species as possible i.e. young and old cap colour, lamellae young and old, stipe young and old, flesh (cut one sporocarp in half lengthwise). If you are using a digital camera, do a ‘white balance correction’ using white paper before taking the image. It is a good idea to include a piece of paper with the collection identification code in one photograph. This will enable you to identify the collections in the photographs at a later date. 3) Once you have taken an image or two of your collection, carefully place all sporocarps in a sheet of aluminium foil. Write down details of the collection in your notebook. These must include comments on locality, the associated vegetation and date of collection. Note how many sporocarps are there and it is particularly important to record characters of the sporocarps that may change after collection: e.g. the presence and colour of veil remnants on the cap and stipe, the presence of liquid droplets, and the smell. Remember: all notes must be made under the correct collection identification code. 4) Then carefully wrap up your collection in the aluminium foil. Be careful doing this because you can easily damage your collection. Remember to include a piece of paper with the collection code on it, in with the sporocarps.

Additional notes for collecting sporocarps of resupinate thelephoroid fungi Where do you find sporocarps and how to collect them Resupinate thelephoroid fungi usually form thin (up to 1-2 mm thick) sporocarps on the underside of dead plant and soil debris. In order to find sporocarps you need to turn over logs, branches and twigs of (well) decayed wood that is lying on the ground. Sporocarps can also be found on the underside of stones, bark, leaves and any other material lying on the ground. Sometimes small sporocarps are also formed inside litter close to the tree roots or even directly on roots.

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Resupinate sporocarps of Tomentella terrestris (left) and Tomentella fibrosa (right). You will need to use a (sharp) knife to cut out the sporocarp along with some part of the substrate. It is important to study the margin of the sporocarp before removing it from substrate in order to ensure that you include in your sample any hyphal cords (rhizomorphs) that may be present in or on the substrate. Hyphal cords are important for identification purposes. The collected sporocarp and substrate can be wrapped in aluminium foil. Remember to include your identification code with the collection. How to collect sporocarps and ectomycorrhizas of the same specimen The mycelium forming sporocarps on the underside of decayed logs emerge either from the soil or from wood depending where the ectomycorrhizal roots are situated. Some species have bright coloured mycelium or hyphal cords and then it is comparatively easy to find and collect ectomycorrhizal root tips connected to the mycelium. The mantle colour of the ectomycorrhizas often corresponds to that of the sporocarp. However, many species have brown or dark coloured mycelium (and sporocarps!) and no hyphal cords. In these cases, it is not usually possible to track mycelium up to root tips except when mycelium and roots are on white rotting wood. Many resupinate thelephoroid species form ectomycorrhizas with dark (brown) coloured mantles (below left) but some have bright coloured ectomycorrhiza (right).

Additional notes for collecting sporocarps of Ascomycetes. Our knowledge on ectomycorrhizal ascomycetes is still rather limited. Important ECM forming genera are found among apothecial ascomycota (discomycetes) of the orders Pezizales (e.g. Geopora, Geopyxis, Morchella, Spaerosporella, Wilcoxinia,), Helotiales (Hymenoscyphus ericae aggregate including Cadophora (Phialophora) finlandia), and the well known anamorphic Cenococcum geophilum (Dothideomycetes). ECM forming members of Pezizales and Helotiales produce conspicuous to minute sporocarps in spring,

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high summer or autumn. Many are irregular in sporocarp production and need a “stimulatory” event of some kind of disturbance (e.g. forest fires). Where do you find sporocarps and how to collect them On this course, we will not observe many ECM forming ascomycetes as sporocarps, but they will certainly be present as ECM roots. When we find ascomycetes of the Pezizales and Helotiales related to well-known ECM formers, we will collect them and look at their characteristics. Ascomycetes of the orders Pezizales and Helotiales produce a range of different apothecial sporocarp forms. The larger ascomycete sporocarps should be easy to find, however, they may turn up in microsites where they are easily overlooked (let us see what we can learn about finding them!). The smaller apothecial ascocarps (a few millimetres) are often attached to rotten leaves, dying roots, or even on fresh roots. A couple of studies suggest that minute sporocarps of some species may even fruit below ground directly on the ECM root tips. The photos below (taken by Nina Wurzburger who reported this) illustrate this phenomenon, e.g. minute stipitate cup fungi (Hymenoscyphus-like) attached to lateral roots and ECM tips:

Finding such sporocarps is a challenge; they potentially represent new unknown ECM taxa, or ECM fungi that, until now, have been recognized only in asexual stages (e.g. in pure culture and as ECM morphotypes, which is the case for the common black and white Piceirhiza bicolorata morphotype formed by fungi of the H. ericae aggregate). Let us try to find them! A complete collection of an apothecial Ascomycota includes several mature, and even overmature, ascocarps. Quite often the hymenium is not fully developed before the apothecium is over-ripe (then looks irregular and "bad"), and details from the mature hymenium are necessary for microscopical determination. At least one ascocarp alone is needed to get enough DNA for sequencing, several sporocarps are needed to make sections for morphological descriptions, and some sporocarps should remain complete and untouched in the collection. Another suspected ECM forming ascomycete genus connected to coniferous forest which we will probably find, is Otidea (Pezizales). The picture below of some Otidea sporocarps with ECM roots underneath (photo Trude Vrålstad) is in fact indicative of an ectomycorrhizal relationship:

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Sporocarps of Otidea spp.

Otidea mycelium

Otidea ECM? In general, ascomycetes are not negatively affected by water. It is therefore a good idea to keep the collection moist; this will not harm the sporocarps, but keep them fresh for a while. If you find ascocarps of the larger type (more than 5 mm), the general method described above for basidiomycetes fungi is sufficient (i.e. wrap in Aluminium foil). However, don't forget to add a moist piece of cleaning paper or moss tufts to keep the sporocarps moist. Small plastic boxes with lids are perfect for collecting medium sized ascomycetes. Larger sporocarps, e.g. Morchella and Gyromitra are better to collect in aluminium foil, but still add something to keep them moist. Try to avoid the sampling of "the whole forest" when collecting small ascocarps embedded in sand. Try to isolate a few "clean" ascocarps (for DNA analyses) in the field. Use an eppendorf tube with moist paper. Collections of minute sporocarps will at best be covered with sand particles, and often “disappear” before you get to the lab, if not adequately cleaned and moistened when you are in the field. An eppendorf tube with moist paper, a small piece of moist moss or a drop of water is recommended. The eppendorf tube is easy to label with a collection code if you have a “permanent marker” pen. If you find sporocarps of ascomycetes that are known or suspected to form ECM, make sure to collect both sporocarps and the substrate including ECM roots below (the “substrate” may be soil, debris, rotten wood etc.). Wrap the collection carefully in aluminium foil. In the lab, you can search for possible hyphal connections between sporocarps and ECM roots and make collections of both.

General comments for annotation of collections voucher specimens, DNAsamples and sequences. In the lab, each good collection should be identified and preserved properly. That includes several steps: Sporocarps: 1) Correct species identification 2) Selection of good voucher specimens (for the herbarium and DNA work) from which a piece should be kept for DNA extraction (transfer a small clean piece of fungal material to an Eppendorf tube with lysis buffer – this will be provided). Each voucher specimen must have a unique code/ID different from collection code/ID. 3) Annotation of the collection and vouchers in an electronic XL-sheet (will be provided)

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4) Dry the collection. Very small voucher specimens should be dried separately in Eppendorf tubes or in envelopes labelled with voucher ID. 5) Mark an herbarium envelope with all required details and transfer the collection to this when completely dry ECM tips: 1) A collection should include several representatives of a specific morphotype within a sample (assuming there are sufficient numbers; these should preferably be adjacent to each other on the same main root). Transfer 4 single tips to separate Eppendorf tubes (for DNA work; transfer a clean single tip to an Eppendorf tube with lysis buffer; this will be provided). If there are sufficient numbers of tips, an additional 5 tips should also be saved in a separate Eppendorf tube. The rest of the collection should be used as a voucher and kept in a fixative (FAA or Gluteraldehyde). 2) Each single tip (for DNA extraction) must have a unique code/ID different from collection code/ID. The tube with the 5 tips and the vouchers should be marked with collection code. 3) Record the collection and vouchers in an electronic Excel-sheet (this will be provided). Good annotation is essential and should include the following elements: •

• • • • • • • •



Collection ID (for example RK00001 which refer to personal initials (Rasmus Kjøller) and the first collection (00001) or NF00001 (referring to the NordForsk course and the first collection). The next collection will then have RK00002 or NF00002 etc. If you use initials, each one of you can have your own system. If we decide to use NF as course ID, we have to coordinate the numbering of collections within the group. Important! Some public herbaria provide their own collection ID-s. If you are going to deposit your collections in such herbarium then it is useful to use such ID-s from the beginning. Collection date Locality: Country (Denmark), County (South West Zealand, SWZ), Municipality (Sorø), local name (e.g. Sorø Sønderskov), coordinates (optional) Vegetation type (forest type and details on the collecting site) Substrate (e.g. rotten wood, forest soil, leaf etc) Plant host (if recognized, e.g. Picea abies or Pinus sylvestris ) Leg (Collected by) : person that sampled the collection Det (Determined by) : person that identified/verified the species Voucher ID: If a collection includes many sporocarps then each sporocarp (or ECM tip) that is going to be cultivated and/or used for DNA-studies should be given a voucher number. Unique voucher numbers could for example be combinations from Collection ID like RK00001/1, RK00001/2 (personal initials) or NF00001/1, NF00001/2 (common system for the course). Dried voucher material (together with the rest of the collection) should be deposited in public herbaria. DNA-samples ID/Sequence ID: From each voucher you may take one or several samples. The sample number should refer to voucher number, and include information whether the sample is taken from sporocarp tissue, ECM root tissue, or cultures (if available). Numbering could be as follows: NF1/1.S1, NF1/1.S2, NF1/1.S3; referring to 3 samples of sporocarp tissue (S) from the voucher specimen NF00001/1. For roots, use collection ID combined with R and numbers, e.g. NF5/R1 (referring to first root-

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• • • •

tip sample in course collection 5). During this course, we will try to put the collections and vouchers into a common system (if possible) and use the system consistently. Photos (ID, reference to photos of the collection and possible vouchers) Sequences (when you get information here you can add sequences available for the, e.g. ITS1, 5.8S, ITS2 (complete, xx base pairs) LSU (partial, xx base pairs etc) Sequence primers (which primers has been used for sequencing, e.g. ITS1F and ITS4) Comments (extra things to note)

General comments for preparation of DNA samples During the field course, it is important that you take responsibility to make samples for DNA analysis. You will be provided with Eppendorf tubes containing lysis buffer. You will each be able to extract DNA from 10 DNA samples during the molecular part of the course. Make at least 10 samples, but not much more (max 20 – the tubes with buffer is a restricted resource)

How to make a sample from sporocarps and ECM tips: • •

• • • •

Choose a good collection and select a voucher specimen Use a sterile (flamed) scalpel and/or forceps to excise one or two small pieces of clean fungal tissue. For sporocarps of agarics, boletes and larger ascomycetes excise flesh inside the cap or stipe). Small ascomycetes (less than 3 mm) must be added as a whole piece (rinse in water). For resupinate thelephoroid fungi, use a clean forceps to select a (as clean as possible) piece of the fungus. For ECM tips, work under a stereo magnifier in distilled water: Excise a single tip from a collection of a specific morphotype and clean it in distilled water. Make sure to remove soil or sand particles from the tip. Do not touch the fungal piece (or ECM tip) if you can avoid it, and use gloves if you need to (to avoid that you transfer RNAses and DNAses on your skin to the sample – those may degrade your DNA). Transfer the clean piece of fungal sporocarp tissue or ECM tip to an Eppendorf tube containing 100 μl lysis buffer (this will be provided). Label the tube with DNA sample ID with a permanent marker pen Keep the sample cold (we will arrange Eppendorf boxes in the fridge)

Student exercises: Describing sporocarps This course is not intended to cover the huge diversity of ECM taxa. Instead we have chosen to focus upon a small number of species rich groups. Using these groups, you will learn how to work scientifically (thoroughly) with a few good collections that you will follow from sporocarp and/or ECM root tips to complete DNA sequences. The fungal groups we have chosen are the Russulaceae, Boletes, Ascomycetes and thelephoroid fungi. Below, you will find some general features for sporocarps and some detailed information on the determination of taxa from the chosen fungal groups. You will use

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this information to identify your collections. We will of course give you all the assistance that you require. 1: Agarics – general features and specific exercises Some general features of Agaricoid sporocarps (modified from Bon 1987)

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1A: Describing Russula sporocarps The objective of this exercise is for you to develop the skills to make detailed descriptions of your field collections. This will not only greatly enhance your chances of correctly identifying your collections but it will also allow others to check the identity of your collections after they have been dried. You are provided with pre-printed recording sheets where you can record the features of your collections. Make sure you write the collection number on the sheet. BUT BEFORE you start going down the list of features stated in the record sheet and filling in your observations, SET UP YOUR COLLECTIONS FOR OBTAINING SPORE PRINTS. Determining the spore colour of a Russula (you can use this method for nearly all agarics). The colour of Russula spores range from pure white to deep yellow – you are provided with a number of colour charts with which to determine spore colour. Although you can get some idea of the colour of the spores by examining the colour of the lamellae, it is essential that you determine the colour accurately on a reasonably thick layer of spores. In order to do this you need to collect a spore print, something best done on a piece of white paper. The time taken for sufficient spores to be deposited varies enormously (note that some collections will NOT give you a spore print no matter how long you wait). It is therefore best to set up the spore prints as soon as you get back to the lab. 1) If you have a number of sporocarps within your collection, choose a specimen (or better still choose several specimens) that is mature but not showing signs of decay. 2) Carefully cut off the stipe with a sharp knife just below the level of the gills. This stump of the stipe should form a small leg (2-3mm longer than the lamellae) on which the cap should rest. Place the cap onto a piece of white paper slightly larger that the cap diameter. Make sure that the lamellae are hanging straight down – the spores need to be able to fall onto the paper. If they are not vertical, change the level of the cap by either shaping the remains of the stipe or by wedging a small piece of material under the edge of the cap that needs to be elevated. Cover the cap with a glass or plastic dish as this prevent the spores being blown away and helps to slow down drying of the cap. You can also place a small dish of water under the dish, beside the cap, to increase the humidity. 3) If you only have a single specimen, cut the cap in half and proceed with one half as outlined above. The lamellae will be in direct contact with the paper if you are using only half a cap. 4) Make sure you mark the paper with the collection number associated with the rest of the collection. 5) Start looking after ca. 25 mins by carefully lifting up the edge of the cap. Remember that if you have a white-spored species it may be very hard to see if it is dropping spores. It is best to wait until you can see that you have a good layer of spores. Once you have this, remove the cap and return to your collection. Allow the spores on the paper to dry for 5-10 minutes. Take a microscope coverslip and VERY CAREFULLY scrape together sufficient spores to give a small heap. THE COVERSLIP IS VERY FRAGILE - DON’T BREAK IT AND CUT YOUR FINGERS. Once you have a small heap, scoop up the spores onto the coverslip and place then in the centre of another coverslip lying flat on the table. Then make a sandwich of the spores between the two coverslips. You can then place your spore sandwich onto the colour chart to compare the spores with the chart. Move the sandwich over the chart until you are satisfied that

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you have the best match between the colour chart and your spore deposit. This needs to be done in good light conditions. Record the colour of the spores (usually given as a Roman numeral and either a, b, c, d, or e - e.g. IIa) on your record sheet. If the colour of your spore deposit falls between two colour codes give both in your description (e.g. IIa, b). 6) Fold the paper so that the spores are on the inside and label with your collection code. This can then be added as a permanent feature to your dried sporocarp collection. Taste and smell Taste is an important factor in species determination within Russula. Although no species are considered to be highly poisonous, some of them contain compounds that make a very hot curry seem like a mild mint. So you need to be careful in tasting your collections. Choose a young clean specimen and remove a small piece of the stipe (pinch a piece out between two finger nails) and chew at the front of your mouth making sure your tongue comes in contact with the material. Some species are mild and remain so no matter how long you chew them. Others are mild at the start and only after 10-30 secs or so will an acrid taste be apparent. Some will, however, become very acrid immediately – as soon as you feel this - spit out the material. Unfortunately, it can be difficult to determine if some collections are acrid because it can be very weak and even lost in older specimens – so always try to use young and undamaged sporocarps. Once you have tried the stipe, you need to try the lamellae. This is because the flesh in some species is mild but the lamellae are acrid. Be careful here because the lamellae can be very acrid. Some species have other components to the taste that you should note. It may taste like a cedar wood pencil or it may be oily or disagreeable or even that your tongue feels cold and you taste menthol toothpaste. The smell is also important in determining species but it can be much more difficult to characterise. Some species do not smell, while others do so faintly. But some emit strong distinctive smells like stewed apples or gooseberry jam or like Pelargonium (Geranium). One group is characterised by having a very disagreeable smell like burned hair or horn! Another group smells like old herring or crabs. The smell of a Russula collection is best appreciated if your collection has been kept in a small container for some time. You can open the container lid a little bit (or open one part of foil storing your collection) and immediately sniff the air that is saturated with the aroma of the sporocarps. Try and relate the smell to something you know. There may be no smell! Macroscopic chemical reactions: 10% Iron sulphate (FeSO4) on the flesh of the stipe – this is a useful reaction that can be used to distinguish certain groups within the genus Russula. If you are lucky you will get a definite rapid colour change to bright salmon pink, or orange, or a strong green colour. If you are less lucky there will be little or no reaction or a rather dull grey-green-pinky coloration. You should note the immediate reaction and after 5-10 minutes as it can take sometime for the colour to develop. To get the best reaction with the iron sulphate it is necessary to slightly damage the surface of the stipe. One simple way of applying the reagent to the stipe is to use a pointer or needle. Simply dip the needle into the reagent and then scrape it onto the stipe surface. Structure of the cap cuticle

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This is a critical feature in the determination of Russula species. There are a number of features of the cap cuticle that you must determine in order to obtain the correct identification for your collections. a) Dermatocystidia (dcy): These are special hyphae in the cuticle of many Russula species that are differentiated from the normal hyphae of the cuticle. The dcy are often enlarged or swollen and may possess specialised apices (see Fig. 1). Carefully peel off a small piece (1mm2) of cuticle - this is best done by pinching the cuticle with your forceps and pulling towards the centre of the cap. Try to get as little of the underlying tissue as possible. Place the piece of cuticle on a drop of water – make sure that the surface is wet from the water. IT MUST BE TOP SIDE UP! Place a cover slip over the cuticle and examine under the compound microscope. Look also to see if there are encrusted hyphae (see part b) below. While you are looking at your preparation determined the shape of the apices of the normal cuticle hyphae. These are most commonly cylindrical but they can be capitate (with a small swollen end portion). With practice the dcy will be easily recognisable but they can be stained in order to make them very obvious. This is done using sulphovanillin: a mixture of sulphuric acid (H2SO4) EXERCISE EXTREME CARE!!! and Vanillin. Take another small piece of cuticle and place in a small drop of the reagent on a slide. It will probably curl up and you will need to use a needle to detach it from you forceps. Place a LARGE cover slip over the cuticle and examine under the compound microscope. If dcy are present they should stain black – occasionally they may only be grey or very infrequently they will not stain. Be sure to record the reaction of your dcy to the sulphovanillin. You will have to work quickly (but carefully) once you have the cuticle in the drop of reagent because it will dissolve. b) Encrusted hyphae: These are differentiated hyphae from the normal hyphae of the cuticle on which there are encrustations on the hyphal walls (Figure 2). With a good microscope with differential interference contrast you can see these without staining but you can stain them with Brilliant cresyl blue. Peel off a small piece of cuticle (as above) and place on a small drop of the reagent, cover with a coverslip and examine with the compound microscope. You may have to look VERY carefully to see the encrustations. Sometimes the hyphae are rather similar to the normal cuticle hyphae and lightly encrusted, other times they can be much enlarged and heavily encrusted and easy to see. You can even find dermatocystidia with encrustations!! Carefully record your observations. Spore ornamentation A general feature of Russula species is that the spores are distinctly ornamented and that within a species the spore ornamentation is relatively consistent. Fortunately for us, the ornamentation is readily stainable, making it easy to see. Place a small drop of Melzers reagent (a aqueous solution of iodine, KI, and chloral hydrate) on a slide and mix in a small quantity of spores from your spore print. Cover and observe with the compound microscope at highest magnification. It is best to use mature spores because immature spores (those still on the lamellae) have an under-developed ornamentation. The ornamentation can take the form of isolated spines, warts or small pustules. Low ridges may join these features or may themselves form strong ridges on the spores. You should note how prominent the ornamentation is and the amount of interconnections.

Figures: 1) Dermatocystidia (Dcy) in the cuticle of Russula atroglauca

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Dcy Cuticle hyphae

2) Encrusted hyphae in the cuticle of Russula turci stained with Brilliant Cresyl blue Encrusted hyphae

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Recording sheet for Russula collections 1. Collection code 2. Locality 3. Substrate 4. Forest type 5. Collected by 6. Collection date 7. Determined by (see back of sheet for possible character states) Sporocarp features Cap Shape and Dimensions: Colour: Surface: Edge: Stipe Shape and Dimensions: Colour: Texture: Lamellae Colour: Interveining: Flesh Colour: Changing: Taste: Smell: Fe reaction Cuticle Dcy (Size, shape, encrusted, staining): Encrusted hyphae (size, degree of encrusting): Cuticle hyphae (size, shape of end cells): Spores Drawings:

Colour: Ornamentation:

Size:

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Sporocarp characters and states for Russula sporocarps Cap shape and size: conical, plano-convex etc. Record maximum diameter Cap colour: can be very variable or uniformly unicolored. Generally describe from the edge towards the centre. There will often be a gradient of colour from the periphery towards the centre. Cap surface: smooth, dry, shiny, viscid (sticky), matt, pruinous (with a very fine dust on the surface). Cap edge: smooth or furrowed or tuberculate (very furrowed with the ridges having bumps) Peelable: grasp the very edge of the cap and peel the cap cuticle towards the centre of the cap. Record as either not peelable or ¼ - ½ or ½ to ¾. Stipe shape and dimensions: cylindrical or swollen towards the base. Length and breadth. Stipe texture: spongy or firm maybe even very firm. Stipe colour: Often white or wholly pink or red but if white then look carefully for faint hints of colour particularly pink or red. Look for brown spots near the base. Is the colour changing where you handle the specimen? Lamellae colour: This will vary with the degree of maturity of the spores, particularly if you have a specimen with darker coloured spores. Interveining between lamellae: seen as cross-connections between the lamellae Flesh taste and smell: Mild or acrid; any smell, faint distinct Flesh colour: Cut a specimen length ways and observe any colour reactions in the flesh. Some species go red when cut, others grey or black. Some initially go red or pink and then grey or black. Take notes over a period of time. Fe reaction: Colour reaction (e.g. pink, orange, green, no reaction, fast slow) Cuticle features: Presence, shape size of dcy or encrusted hyphae or both. Shape and size of cuticle hyphae Spores: Colour (e.g. Ib, IIa etc), size, ornamentation (high or low warts or spines, interconnected or free)

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1B: Describing Lactarius sporocarps. The objective of this exercise is for you to develop the skills to make detailed descriptions of your field collections. This will not only greatly enhance your chances of correctly identifying your collections but it will also allow others to check the identity of your collections after they have been dried. You are provided with pre-printed recording sheets where you can record the features of your collections. Make sure you write the collection number on the sheet. In addition to recording the general features of the sporocarps (see recording sheet) you also need to determine the following specific characters: Taste and smell Taste is an important factor in species determination within Lactarius. Like some species within the genus Russula, some Lactarii contain compounds that are very acrid to taste. So you need to be very careful in tasting your collections. Choose a young clean specimen and remove a very small piece of the lamellae and chew at the front of your mouth making sure your tongue comes in contact with the material. Some species are mild and remain so no matter how long you chew them. Others are mild at the start and only after 10-30 secs or so will an acrid taste be apparent. Some will, however, become very acrid immediately – as soon as you feel this - spit out the material. The smell is also important in determining species. Some species do not smell, while others do so faintly. Some emit strong distinctive smells like coconut (L. glyciosmus, L. mammosus) or curry powder (L. helvus). The smell is often best appreciated if your collection has been kept in a small container for some time. You can open the container lid a little bit (or open one part of foil storing your collection) and immediately sniff the air that is saturated with the aroma of the sporocarps. Try and relate the smell to something you know. There may be no smell! The milk The genus Lactarius is characterised by the presence of a liquid, the milk, which leaks from broken sporocarp tissue. The milk is contained within special hyphae called lactifers or laticiferous vessels. These are disrupted when you damage the sporocarp and the milk leaks out. The milk colour and taste are important characters in species determinations. Observe the colour of the milk when it leaks out and any changes that happen over time. In some species the milk is clear i.e. colourless. It may be white and change to yellow, or violet or grey-green. In others, initially orange milk can become red. You may see old dried spots on the lamellae that indicate what colour the milk can become. In some species the milk changes colour when in contact with the flesh of the sporocarp but not when it is isolated. Test this by soaking up a small amount of milk on white paper or a white tissue and check the colour after 5 to 10 mins. Spore ornamentation A general feature of the Russulaceae is that the spores are distinctly ornamented and that within a species the spore ornamentation is relatively consistent. Fortunately for us, the ornamentation is readily stainable, making it easy to see. Place a small drop of Melzers reagent (a aqueous solution of iodine, KI, and chloral hydrate) on a slide and place a small piece of lamella in the drop, cover and observe at highest magnification. The ornamentation usually takes the form of ridges. Note how prominent the ridges are and how organised the network is – i.e. are the spores netted or merely covered in a disorganised set of ridges.

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Recording sheet for Lactarius collections 1. Collection code 2. Locality 3. Substrate 4. Forest type 5. Collected by 6. Collection date 7. Determined by (use back of sheet for additional notes) Sporocarp features Cap Shape and Dimensions: Colour: Surface: Edge (smooth, hairy?): Stipe

Shape and Dimensions: Colour:

Lamellae

Texture:

Colour: Interveining:

Flesh Milk

Colour:

Changing:

Taste:

Smell:

Initial colour:

After 10 mins:

Colour change when isolated: Spores

Drawings:

Size:

Ornamentation:

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Sporocarp characters and states for Lactarius Cap shape and size: conical, plano-convex etc. Record maximum diameter Cap colour: can be very variable or uniformly unicolored. Generally describe from the edge towards the centre. There will often be a gradient of colour from the periphery towards the centre. Cap surface: smooth, dry, shiny, viscid (sticky), gelatinous (slimy), matt, pruinous (with a very fine dust on the surface). Cap edge: smooth or furrowed or tuberculate (very furrowed with the ridges having bumps), is there a fringe of hairs? Are these dry or slimy? You need to check the youngest specimens for this character. Stipe shape and dimensions: cylindrical or swollen towards the base. Length and breadth. Stipe texture: spongy or firm maybe even very firm. Is it hollow? Stipe colour and surface: Often follows the cap colour. Check for a pale ring at the stipe apex. In some species the stipes darken from the base upwards. Are there pits in the surface? Lamellae colour: This will vary with the degree of maturity of the spores, particularly if you have a specimen with darker coloured spores. Interveining between lamellae: seen as cross-connections between the lamellae Flesh taste and smell: Mild or acrid – quickly / slowly; any smell - faint / distinct. Flesh colour: Cut a specimen length ways and observe any colour reactions in the flesh. Any colour change is primarily due to the milk. Take notes over a period of time. Spores: Size, ornamentation (high or low warts or spines, interconnected or free)

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1C: Describing and identifying boletoid sporocarps You are provided with pre-printed recording sheets (containing the information below) where you can record the features of your collections. Make sure you write the collection number on the sheet. Your observations should, if possible, be made on sporocarps at all stages of development. Some of the terms used to describe the sporocarp features may be new to you and you can use the diagram of sporocarp features to get an explanation of the more common terms. The Boletales are a monophyletic order of homobasidiomycetes that includes ca. 1000 described species worldwide. Many members of the Boletales form mycorrhizas with a wide range of host species, but some are saprotrophs living on decaying wood, and a few are suspected to be mycoparasites. They are very diverse in both morphology and ecology. Fruiting bodies in the group include stipitate-pileate forms (with stalk and cap), crust-like resupinate forms, earthballs, and false truffles. In this course we will concentrate on those Boletoid taxa that have a stem and a cap. You may be most familiar with those taxa that bear their spores within tubes rather than on gills, but hopefully during this course you will get a chance to examine both poroid (with tubes) and lamellate forms (with gills) of boletoid fungi. The lamellate genera you are likely to encounter include: Paxillus, Chroogomphus, and Gomphidius The poroid genera you are likely to encounter include: Boletus, Chalciporus, Leccinum, Strobilomyces, Suillus, Tylopilus, and Xerocomus. You are provided with specialist keys to the more species rich genera of boletes. Below is a summary of some of the important characters you will have to observe in order to be able to identify collections of boletoid fungi. One character that can be very important for the identification of some bolete taxa is the host species. In a mixed forest, it is better to record all potential hosts within ca. 20 m, than to try and guess which particular tree is the host. Recording sheet for Boletoid fungi 1.

Collection code

2.

Locality

3.

Substrate

4.

Forest type

5.

Collected by

6.

Collection date

7.

Determined by Sporocarp features

(use back of sheet for additional notes)

Cap

Shape and Dimensions: Colour: Surface: Edge (smooth, hairy?):

Stipe

Shape and Dimensions: Colour:

Network:

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Lamellae/Tubes

Colour/depth:

Pores

Colour and shape:

Flesh

Colour:

Changing:

Taste:

Smell:

Size:

Ornamentation:

Spores

Notes:

Sporocarp characters and states. Cap shape and size: Hemispherical, plano-convex (see figure for terms and shapes). Record maximum diameter Cap colour: This can vary tremendously with the maturity of a sporocarp. It is therefore very important to determine colour on young and old material. Generally describe from the edge towards the centre. There will often be a gradient of colour from the periphery towards the centre. Cap surface: smooth, felty, dry, shiny, viscid (sticky) – in dry weather it may be difficult to tell if the cap surface should be viscid. One way of doing this is to touch your moist lips to the cap surface (the kissing test). If your lips stick slightly to the cap it is likely that the cap would normally be viscid. Cap edge: Is the cap margin overhanging the edge of the cap? In some Leccinum species, there are flaps of tissue 3-4mm in size, overhanging the edge of the cap. Tube or lamellae depth and attachment: How deep are the gills or tubes? How is the spore bearing structure attached to the stipe? Check the colour figure of sporocarp features for the possible character states. Tube and pore colour: In some species the tubes and pores are different colours. Make sure you check young and old specimens. Stipe shape and dimensions: The stem may be swollen slightly at the base making it slightly club shaped. It may be swollen like a bulb (bulbous). Record the length and breadth of the stipe in particular the width of the base if different from the rest. Does the stipe grow into the soil – is it rooting? Ring on the stipe: In some Suillus species there is a ring or a ring zone on the stipe. Be careful in determining this because in certain conditions (dry weather), the ring can become detached and be stuck on the edge of the cap! Stipe surface: The stipe surface can be an important character is distinguishing both between genera and species. For example, the stipe surface of Leccinum is adorned with numerous scales, which generally change colour as the sporocarp matures. In Boletus, many species

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have a net or reticulum on the stipe – note where this occurs, how strong it is and what colour it is. In Suillus, many species have glandular dots of the stipe- note what colour these are when fresh and when they are dry. Flesh taste and smell: Mild, or bitter. In general, smell is not an important character in separating boletes. Most either don’t smell of have a smell like rubber. However, Boletus satanas stinks like rotting cabbage - once smelt, never forgotten and Boletus legaliae smells of chicory as it dries. Try gently scraping the stipe with your fingernail or cut a specimen in half and check the smell of the cut flesh. Flesh colour: Cut a specimen length ways and observe the colour of the flesh. It is common that different parts of the flesh have different colours. Observe any colour reactions in the flesh. Some Boletus species stain blue or blue-black very easily when damaged and this can happen so fast that it is difficult to determine what the original colour was. In Leccinum and Xerocomus, the colour change can take some time (5-10 minutes) to develop– it is often a good idea to gently scrape your knife along one of the cut surfaces. This damaged surface will give a much stronger colour change than the unscraped surface and you can compare the two. Note where you get colour changes. Chemical reactions: place a very small drop of the reagent on the surface to be tested and observe any colour change. Spores: The size, shape and surface of spores are important characters used in the identification of boletoid taxa.

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2: Describing apothecial sporocarps (ascocarp) The objective of this exercise is for you to develop the skills to make detailed descriptions of your field collections. This will not only greatly enhance your chances of correctly identifying your collections but it will also allow others to check the identity of your collections after they have been dried. You are provided with pre-printed recording sheets where you can record the features of your collections. Make sure you write the collection number on the sheet. Ascocarp characteristics We will not have time to enter the conceptual framework and characteristics for all morphological features shared by apothecial fungi, neither among the known ECM-forming genera. However, checking a few general features in fresh ascocarps, according to the recording sheet below, will greatly help in identification work. Apothecial ascocarps may be differentiated in a cap or stipe (like Morchella, Gyromitra, Helvella, Hymenoscyphus), or they may be unstipitate and then regular to irregular discoid to pulvinate in shape. It is of utmost importance to recognize the shape and position of (a variety of) hairs on the apothecial outside (receptacle) and at the margin. Many genera, and even families, are recognized on the basis of presence of certain hair types. For example, the ECM forming genus Wilcoxina is characterized by bundles of stiff brown hairs on the receptacle, in contrast to the closely related Trichophaea (not yet recorded as a ECM former) with scattered, evenly distributed stiff hairs on the receptacle. The shape and opening of the ascus distinguish Pezizales and Helotiales, the operculate (opening by a lid), cylindrical ascus being typical of the former, and the inoperculate (with a pore in ascus apex), clavate ascus of the latter order. The bluing of the ascus tip in Melzer's reagent (an aqueous solution of iodine, KI, and chloral hydrate) is important to distinguish among genera and groups of taxa of the two respective orders. The paraphyses (sterile hyphae) also constitute an important diagnostic character on generic and species level. Here the colour (gives colour to the hymenium), type of pigmentation, e. g. crystalline granules or oil globules inside cells, or in some cases pigments deposited outside the cell walls, "gluing" the paraphysal ends together in a mazaedium, and shape are important to record in fresh specimens, as such characters more or less disappear in re-vived (in H2O) material. Whether the paraphyses are branched or not, distinctly enlarged at the tips, longer than asci, irregular in shape, and straight, curved or coiled in upper part are also important generic characters. The ascospores of the Pezizales are generally unicellular, large (> 10 um), broadly ellipsoid to globose, often with characteristic internal oil globules, and many representatives have ornamented spores that need to be observed with the compound microscope at highest magnification. Make a squash preparation mounted in Cotton Blue (methylene blue), and the spore wall (inclusive ornamentation) is strongly taking colour, making the various ornamentation patterns (spines, warts, reticulum etc.) clearly visible. Only mature spores have a fully developed ornamentation! OBS! many of the ECM forming genera of the Pyronemataceae known to date, e. g. Geopyxis, Wilcoxina, Tricharina, Sphaerosporella, all have smooth ascospores of different shape and "globulation". The ascospores of the Helotiales are unicellular or multi-cellular, generally small (< 10 um), and usually narrowly ellipsoid to allantoid to filiform in shape. The spores are, with very few exceptions, smooth. Literature needed for the course: Nordic Macromycetes, Vol 1., Fungi of Switzerland, Vol. 1.

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Recording sheet for apothecial ascocarp collections 1. Collection code 2. Locality 3. Substrate 4. Forest type 5. Collected by 6. Collection date 7. Determined by Ascocarp features Cap (Cup) Shape (cupulate, discoid, pulvinate etc.) and Dimensions: Colour: Surface (receptaculum): Margin (with/without hairs etc.): Stipe

Shape (unstipitate, stipitate) and Dimensions:

Hairs

Colour: Shape: Colour: Dimensions: Position (in tufts, scattered, at margin etc):

Asci

Shape (cylindical, clavate): Opening (pore, operculum): Melzer's reaction: bluing (J+, amyloid), not bluing (J-, inamyloid)

Paraphyses

Shape (enlarged/curved/irregular at tips,branched etc.) Pigment deposits (granules, oil globules, mazaedium)

Spores Drawings:

Colour: Size:

One-celled/multi-celled: Guttules: Ornamentation (Cotton blue):

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3: Describing resupinate thelephoroid sporocarps Character list #1. basidiocarps / 1. resupinate/ 2. with clavarioid processes at the margin, or effused-reflexed/

Fig. 1 (left). Basidiocarp of Tomentella radiosa with clavarioid processes at margin (TAA-152360). Fig. 2 (right). Basidiocarp of T. atramentaria with clavarioid processes at margin (TAA-90149). #2. basidiocarps / 1. separable from the substratum/ 2. adherent to the substratum/ #3. basidiocarps / 1. arachnoid/ 2. pelliculose/ 3. mucedinoid/ 4. crustose/ #4. hymenophore/ 1. white or whitish/ 2. red or reddish brown/ 3. brown/ 4. yellow or green/ 5. blue or grey/ 6. almost black/ #5. / #6. hymenophore/ 1. smooth/ 2. granulose/ 3. colliculose/ 4. hydnoid/ 5. poroid/ #7. hymenophore/ 1. turning paler than subiculum/ 2. concolorous with subiculum/ 3. turning darker than subiculum/ #8. subiculum/ 1. white or whitish/ 2. red or reddish brown/ 3. brown/ 4. yellow or green/ 5. blue or grey/ 6. almost black/ #9. sterile margin/ 1. determinate/ 2. indeterminate/

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#10. sterile margin/ 1. paler than hymenophore/ 2. concolorous with hymenophore/ 3. darker than hymenophore/ #11. sterile margin/ 1. white or whitish/ 2. red or reddish brown/ 3. brown/ 4. yellow or green/ 5. blue or grey/ 6. almost black/ #12. hyphal cords/ 1. absent/ 2. present only in subiculum/ 3. present in subiculum and margins / #13. hyphal cords / 1. monomitic/ 2. dimitic/

Fig. 3. Monomitic hyphal cord (from T. radiosa, TAA-111451).

Fig. 4. Dimitic hyphal cord (from T. ferruginea, TAA-159024). #14. monomitic hyphal cords / μm diam/ #15. monomitic hyphal cords / 1. hyaline/ 2. yellowish/ 3. pale brown / 4. dark brown/ 5. greenish / #16. individual hyphae / 1. clamped/ 2. simple-septate/ 3. skeletal/

Fig. 5. Clamped thin-walled (from T. radiosa, TAA-111451) and thick-walled (from T. atramentaria, TAA149211) hyphae.

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Fig. 6. Simple-septate thin-walled (from Pseudotomentella nigra, TAA-149827) and thick-walled (from P. tristis, TAA-149526) hyphae.

Fig. 7. Skeletal hyphae from the surface of hyphal cords (from P. griseopergamacea, TAA-152789). #17. individual hyphae / μm diam/ #18. individual hyphae / 1. hyaline/ 2. yellowish / 3. pale brown/ 4. dark brown/ 5. greenish / #19. individual hyphae / 1. thin-walled/ 2. thick-walled/ #20. dimitic hyphal cords / μm diam/ #21. dimitic hyphal cords / 1. yellowish/ 2. pale brown / 3. dark brown/ 4. bluish or greenish / #22. individual hyphae / 1. clamped/ 2. simple-septate/ 3. skeletal/ #23. clamped hyphae / μm diam/ #24. clamped hyphae / 1. hyaline/ 2. yellowish / 3. pale brown/ 4. dark brown/ #25. simple-septate hyphae / μm diam/ #26. simple-septate hyphae / 1. hyaline/ 2. yellowish / 3. pale brown/ 4. dark brown/ 5. greenish / #27. skeletal hyphae / μm diam/

Page 35 (81) #28. skeletal hyphae / 1. hyaline/ 2. yellowish / 3. pale brown/ 4. dark brown/ #29. subicular hyphae/ 1. skeletal/ 2. simple-septate/ 3. clamped/ #30. cross-shaped branching of subicular hyphae/ 1. common/ 2. rare/ 3. absent/

Fig. 8. Cross-shaped branching of hyphae of Tomentellopsis zygodesmoides (TAA-149543). #31. skeletal hyphae / μm diam/ #32. skeletal hyphae / 1. thin-walled/ 2. thick-walled/ #33. skeletal hyphae / 1. yellowish/ 2. pale brown/ #34. simple-septate hyphae / μm diam/ #35. simple-septate hyphae / 1. thin-walled/ 2. thick-walled/ #36. simple-septate hyphae / 1. encrusted/ 2. without encrustation/

Fig. 9. Encrusted simple-septate (from Pseudotomentella nigra, TAA-149827) and clamped (from Tomentella lapida, TAA-149608) hyphae. #37. simple-septate hyphae / 1. hyaline/ 2. yellowish/ 3. pale brown/ 4. dark brown/ 5. greenish / #38. clamped hyphae /

Page 36 (81) μm diam/ #39. clamped hyphae / 1. thin-walled/ 2. thick-walled/ #40. clamped hyphae / 1. encrusted/ 2. without encrustation/ #41. clamped hyphae / 1. hyaline/ 2. yellowish/ 3. pale brown/ 4. dark brown/ #42. hymenophoral trama / 1. absent/ 2. present/ #43. sterility of mature teeth / 1. with sterile apex/ 2. at least upper half sterile/

Fig. 10 (left). Section through tooth of T. fibrosa (TAA-159073). Fig. 11 (right). Section through tooth of T. crinalis (TAA-149492) #44. tramal hyphae/ 1. cystidia like/ 2. not cystidia like/ #45. tramal hyphae/ 1. skeletal/ 2. simple-septate/ 3. clamped/ #46. tramal hyphae / μm diam/ #47. tramal hyphae/ 1. thin-walled/ 2. thick-walled/ #48. tramal hyphae / 1. hyaline/ 2. yellowish/ 3. pale brown/ #49. subhymenial hyphae/ 1. simple-septate/ 2. clamped/

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#50. subhymenial hyphae / μm diam/ #51. subhymenial hyphae/ 1. thin-walled/ 2. thick-walled/ #52. subhymenial hyphae / 1. hyphal cells short and inflated/ 2. hyphal cells not short and inflated/

Fig. 12. Short and inflated hyphal cells of T. sublilacina (TAA-149847). #53. subhymenial hyphae / 1. hyaline/ 2. yellowish/ 3. pale brown/ 4. greenish / 5. bluish / 6. violet / #54. cystidia/ 1. absent/ 2. arising from subhymenial hyphae/ 3. arising from subicular hyphae/ #55. cystidia / μm long/ #56. cystidia / μm diam at base/ #57. cystidia / μm diam at apex/ #58. cystidia / 1. clavate/ 2. acuminate/ 3. capitate/ 4. hyphoid/

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Fig. 13.

Fig. 14.

Fig. 15.

Fig. 16.

#59. cystidia/ 1. projecting above the hymenium/ 2. embedded/ #60. cystidia/ 1. simple-septate along their length/ 2. aseptate along their length/ #61. cystidia / 1. encrusted/ 2. without encrustation/ #62. cystidia / 1. hyaline/ 2. pale brown/ #63. basidia / μm long/ #64. basidia / μm diam at apex/ #65. basidia/ 1. simple-septate at base/ 2. clamped at base/

Fig. 17 (left). Simple-septate, utriform and stalked basidia of Pseudotomentella mucidula (TAA-149719). Fig. 18 (right). Clamped, utriform and not stalked basidia of Tomentella bryophila (TAA-149203). #66. basidia / 1. clavate/ 2. utriform/

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Fig. 19. Clavate basidia of T. ferruginea (TAA-159024). #67. basidia/ 1. stalked/ 2. not stalked/ #68. basidia / 1. hyaline/ 2. yellowish/ 3. pale brown/ 4. greenish / 5. bluish / #69. basidia / 1. 2 sterigmata/ 2. 4 sterigmata/ 3. rarely 5-6 sterigmata/ #70. basidiospores / μm long/ #71. basidiospores / 1. slightly globose/ 2. ellipsoid/ 3. triangular with widened proximal part/ 4. lobed/

Fig. 20 (left). Slightly globose frontal face of basidiospores of T. stuposa (TAA-149749). Fig. 21 (right). Ellipsoid frontal face of basidiospores of T. subclavigera (TAA-106953).

Fig. 22 (left). Triangular frontal face of basidiospores of T. ferruginea (TAA-159024) Fig. 23 (right). Slightly lobed frontal face of basidiospores of T. sublilacina (TAA-159039). #72. basidiospores / 1. slightly globose/ 2. ellipsoid/

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Fig. 24 (left). Slightly globose lateral face of basidiospores of T. lapida (TAA-149608). Fig. 25 (right). Ellipsoid lateral face of basidiospores of T. ellisii (TAA-149037). #73. basidiospores / 1. smooth/ 2. verruculose/ 3. bi- and trifurcate/ 4. echinulate/

Fig. 26 (left). Smooth basidiospores of Amaurodon mustialaënsis (TAA-58379). Fig. 27 (right). Verruculose basidiospores of Tomentella calcicola (TAA-59642).

Fig. 28 (left). Bi- and trifurcate basidiospores of Pseudotomentella griseopergamacea (TAA-149851). Fig. 29 (right). Echinulate basidiospores of Tomentella bryophila (TAA-149203). #74. basidiospores / 1. hyaline/ 2. yellowish/ 3. pale brown / 4. dark brown>/ 5. reddish / 6. greenish / 7. violet / #75. chlamydospores/ 1. present / 2. absent/

Fig. 30. Chlamydospores of Pseudotomentella vepallidospora (TAA-149645). #76. chlamydospores / μm diam/ #77. chlamydospores/ 1. hyaline/ 2. dark brown/

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Recording sheet for resupinate thelephoroid collections 1. Collection code 2. Locality 3. Substrate 4. Forest type 5. Collected by 6. Collection date 7. Determined by Basidiocarp features Basidiocarp Characters 1 - 11 from the character list Hyphal cords Subicular hyphae Subhymenial hyphae Cystidia Basidia Basidiospores

Characters 12 - 28 from the character list Characters 29 - 48 from the character list Characters 49 - 53 from the character list

Chlamydospores

Characters 75 – 77 from the character list

Drawings:

Characters 54 - 62 from the character list Characters 63 - 69 from the character list Characters 70 – 74 from the character list

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Wednesday August 15th Characterisation of ectomycorrhiza Over the past two days you have examined in detail the sexual structures (sporocarps) produced by ECM fungi. You identified the sporocarps to species by determining the taxonomically relevant features present on the sporocarps. The overall aim of this next part of the course is to give you some experience in examining the vegetative features of ectomycorrhizal fungi that are taxonomically informative.

Background to ectomycorrhizal characterisation The use of the reproductive structures or sporocarps of ectomycorrhizal (ECM) fungi in taxonomic studies is well established, but few researchers are aware that the vegetative structures, the mantle, rhizomorphs and extra-radial mycelium (Fig. 1), can also provide valuable information on the taxonomic identity of ECM fungi. The preparation of detailed descriptions of these structures has been termed ectomycorrhizal characterisation and although most work on this subject has been carried out in the last 20 years, the earliest investigations are much older.

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Figure 1. Vegetative structures of the ectomycorrhizal association: (a,b) Mycorrhizal tips, (c) mantle and (b,d) rhizomorphs.

a

b

c

d

The first descriptions of the vegetative structures of ECM fungi were done by A.B. Frank (Fig. 2) in 1885. Frank, a German professor, was employed by the Kaiser to grow summer truffles (Tuber aestivum, an ECM ascomycete) and it was he who actually first described the mycorrhizal association and coined the term mycorrhiza. Figure 2. A.B. Frank and the drawings he made of mycorrhizal tips on European beech (Fagus sylvatica).

These early observations were very accurate and Frank observed most of the structures that are still used today in the characterisation of mycorrhizal fungi on roots. At the time neither Frank nor anyone else involved in mycorrhizal research realised the significance or usefulness of the vegetative features of ECM fungi in their identification. It was not until the 1950 and

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60s when Dominik (1969) and Chilvers (1968) attempted to classify ECM morphologies into recognisable categories, that interest was reawakened in these structures. However, it was only in the mid 1980’s that the characterisation of ectomycorrhizas was established as a discipline in its own right. The main driving force behind has been Professor Reinhard Agerer (Fig. 3), who is based in Munich in southern Germany. Figure 3. Professor Reinhard Agerer, who is the leading authority on ectomycorrhizal characterisation and who has been responsible for establishing the methodology involved in the field.

It is due to Prof. Agerer and his co-workers that we know so much about the diversity and complexity of ECM vegetative structures. During this course we will examine the ECM material using the principles and methodologies that he has established. In 1986, Prof Agerer published the Colour Atlas of Mycorrhizae, which contained colour and black and white photographs of ectomycorrhizas formed by a number of ECM species. At regular intervals, new sets of plates are added to this Atlas, which now contains 180 descriptions. This Atlas, along with the detailed descriptions published in the ‘Descriptions of Ectomycorrhizae’ (Agerer et a., 1996-2006), are the main reference literature for ECM characterisation. 1. Extracting mycorrhizal tips from field samples Aim: extract all live mycorrhizal tips with the minimum of disturbance A) Samples should first be soaked in water for at least ½ hour. This loosens up the soil making it easier to extract the root material and reducing the amount of damage caused to the mycorrhizal tips. Gently loosening the sample manually will speed up the soaking process. B) Pour the water and sample over a set of sieves. The best size combination of sieves is one at 2 mm over one with a mesh size of 250µm. Gently rinse your sample under a stream of water. Try to remove as much soil material as possible but with the minimum amount of disturbance to the root material. If there are large chunks of soil adhering to the roots, it is better to remove these from the sieve and place them in water in a large 15 cm Petri dish. Once you have removed all of the soil debris that you can, pick out all the root fragments visible to the naked eye and place them in a large Petri dish. C) Drain the water from the sieve and then place it under a dissection microscope and systematically scan the sieve for all live root fragments. Place these in the large Petri dish with the rest of the root material. 2. Morphotyping Aim: To group together all the morphologically similar mycorrhizal tips. A) Place a large Petri dish under the dissection microscope and partly fill with water. Then take one piece of root material from the Petri dish containing all of the material from the

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sample. Using fine forceps remove the attached debris from the root – if there are rhizomorphs on the tips try to leave these as intact as possible. Once you have removed the soil debris, examining the mycorrhizal tips for general growth form, hydrophobicity, colour, presence / absence of rhizomorphs and extra-radical mycelium development. Separate mycorrhizal tips that you think are morphologically distinct and place them in water in a cell of the multiwell dish. B) Repeat this process until you have worked through all of the root material from the sample. C) Finally, examine the multiwell dish under the dissection microscope. Go over each well and make sure that all of the tips in one well look the same. These groups of tips now represent your morphotypes. Makes notes on the characters of each morphotype that you used to distinguish between them. 3. Mantle preparations Aim: To confirm that your morphotypes are distinct and that within each morphotype, tips share the same anatomical features. A) Start with the most abundant morphotypes. Select a representative tip, remove from the well with forceps and place on a microscope slide. Only use the amount of water that came along with the tip. This will, however, dry up fairly quickly and you should prevent this by dipping closed forceps in water and adding this water to your tip. Using a small quantity of water gives you much greater control over the tip and the pieces of mantle that you remove. B) Hold the tip at the proximal end (the end at which it was attached to the main root). First, detach any rhizomorphs attached to the tip and move them a short distance away from the tip. Next, from a region about half way along your tip, try to remove pieces of the mantle with a sharp pointed dissection needle. If you have a very fine pointed pair of forceps, you can also use a single point of the forceps to remove the mantle. This is best done by pushing the pointed tip into the root and moving the needle towards the distal end of the root. Some mantles become detached very easily, others cannot be removed without cortical cells going with them. C) When you have managed to get some pieces of mantle, make sure that they are lying with the outside uppermost. You will be able to tell which way is up because of the curvature of the mantle. It is also important that you also have one piece which is upside down, so that the inner surface of the mantle is uppermost. D) Try and remove the mantle from the extreme tip of the root. This should come off as a small cap. Make sure this is sitting with the outside uppermost. E) Remove the remains of the root tip. Add a small quantity of water to your mantle preparation and make sure that the mantle and rhizomorphs pieces are near the centre of the drop in the correct orientation and the DROP a coverslip onto them. If you add the coverslip from one side all of the mantle pieces will end up at the end of your coverslip. F) Examine your preparation under 400x and then under 1000x. Determine the nature (see questions and notes below) of any extraradical mycelium extending out from the mantle, including any rhizomorphs. Add these characters to the notes you made on the morphotype.

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G) Focus on the upper most layer of the mantle and determine the shape of the cells. Then focus down into the mantle and determine the shape of the cells in deeper layers. Add these characters to the notes you made on the morphotype. H) Make preparations for at least another two tips from the same morphotype to determine if they have the same anatomy. I) Repeat this whole process for each morphotype. 5. Microscopic features Features to look for in your mantle preparation: Rhizomorphs (see notes below) Are there rhizomorphs present? Where were the rhizomorphs attached to the tip (distally / proximally)? Do the rhizomorphs have smooth or diffuse edges? Are these simple with all the hyphae the same diameter? Are some of the inner hyphae of a larger diameter? Are the septae of the enlarged hyphae dissolved or intact? Emanating hyphae (see notes below) Is there hyphae emanating from the mantle? Are there encrustations or warts on the surface? Are the hyphae clamped? Are there anastomoses between the emanating hyphae? Are these septate or open? Are the septae simple or clamped? If clamped, are they contact clamps? Are the walls coloured?

If so what colour are they?

Cystidia (See notes below) Are there short, determinate hyphae emanating from the mantle? What shape are these?

Are these clamped at the base?

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The Mantle (see notes below) Are the cells in the surface layer of the mantle linear or cellular? Does the mantle have the same structure all the way through? 6. Preparing tips as vouchers specimens and for molecular analysis A) Representative tips from each morphotype should be stored in a suitable fixative to act as voucher specimens for future reference. Many people use FAA (formaldehyde : ethanol 70% : acetic acid = 5 : 90 : 5), which can be stored at room temperature in air tight containers. B) If there is sufficient number of tips within a morphotype it is a good idea to place a representative tip in each of four separate Eppendorf tubes and then have a fifth tube with five tips. Tips can be placed in either 70% ethanol or in extraction buffer (e.g. CTAB). It is also possible to place the tips directly in the Eppendorf tubes and freeze them without placing them in any liquid. Notes on Rhizomorphs Many ECM fungal species form complex linear, multihyphal structure called rhizomorphs. These may comprise aggregations of simple hyphae (Fig 4A,B) or be more complex with differentiated inner hyphae surrounded by a layer of simple hyphae (Fig. 4C-E). The inner differentiated hyphae may be highly developed where the septae of connecting cells have been dissolved (Fig. 4E). The structure of the rhizomorphs may be difficult to see because the hyphae may be coloured or more commonly, the rhizomorphs are highly hydrophobic. When the rhizomorphs are hydrophobic, air trapped in the rhizomorphs will prevent you from seeing any of the internal structures. Mounting the rhizomorphs in 5% KOH will remove the hydrophobicity and allow you to view the structures.

Figure 4. Different rhizomorphs structures of ECM fungi. Left to right A-E (from Agerer 1991)

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Notes on Emanating Hyphae The development of the extraradical phase of ECM fungi differs greatly among species. Some genera (e.g. Cortinarius) are characterised by a very extensive development with numerous rhizomorphs and diffuse mycelia. Other genera (e.g. Russula and Lactarius), in general, form very little mycelium and it can be very difficult to find any hyphae emanating from the mycorrhizal tips. The hyphae of most ECM fungi are smooth and septate with very few distinguishing features. In some genera, the wall surface of the hyphae may be rough (e.g. Amphinema), have warts (e.g. Tylospora) or crystals (e.g. Piloderma, Hebeloma). The presence of clamped septae readily distinguishes the ECM fungus as a basidiomycete, but unfortunately not all basidiomycete fungi form clamps. If clamps are present, note whether all the septae are clamps. In some genera, (e.g. Hygrophorus) clamps only occur intermittently. It is sometimes possible to discern the presence of Woronin bodies in close proximity to the centre of septae. These are oval to spherical proteinaceous structures that are involved in plugging the central pore of the septum in the event of the hyphal cell disrupting. Woronin bodies are only found in ascomycete fungi. When the hyphae of many fungi grow in close proximity to each other, they form connections between individual hyphae called anastomoses (singular: anastomosis). These features can be very useful for distinguishing between morphotypes. Anastomoses can take several forms (Fig. 5). The simplest form is where the walls of the hyphae fuse together (Fig. 5b). The wall may break down and the hyphae are then joined by a bridge of varying length (Fig. 5A, C-F). The length of the bridge can be diagnostic. A septum may form on the bridge and this may be simple (Fig. 5E) or clamped (Fig. 5F). Within the genus Dermocybe a special anastomosis is found called a contact clamp where hyphae are joined directly by a clamped septum (Fig. 5C). The colour and thickness of the walls of emanating hyphae can also be very useful. Most hyphae are colourless or hyaline, some may appear yellow but a large number have melanised walls. The colour of this latter group range from lightly coloured walls that appear slightly brown (some Pseudotomentalla species) to heavily melanised walled that are black (Cenococcum geophilum and some Tomentella spp.). Figure 5. Different types of anastomoses between ectomycorrhizal fungal hyphae (from Agerer, 1991).

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Notes on Cystidia Attached to the surface layer of some mantles there are short, determinate length hyphae that have a specialised shape. These are termed cystidia and, when present, may be highly diagnostic of particular groups of fungi. The figure below shows the range of cystidia that have been observed on the mantles of ECM fungi. Figure 6. Types of cystidia that have been observed on the mantles of ectomycorrhizas (Partly from Agerer, 1991).

Notes on Mantle structures The structure of the mantle in plan view is the most informative character for distinguishing between ECM species. The arrangement and structure of the hyphae in the outermost layer is of special importance. Two basic patterns can be distinguished: in plectenchymatous mantles the linear nature of the hyphae is still visible, while in pseudoparenchymatous mantles the linear nature of the hyphae has been lost and the hyphal cells appear as short expanded cells. Figure 7A-H show a range of mantles that can be classified as plectenchymatous mantles and figs 7I-Q show pseudoparenchymatous mantles. The cellular nature of pseudoparenchymatous mantles usually changes to a linear form in the inner layers. A delicate outer linear network may also be found overlying a strongly developed pseudoparenchymatous layer (Figs. 7P & 7Q). The hyphae of the mantle may be embedded in a gelatinous matrix (Fig. 7C).

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Figure 7. Mantles structures of ectomycorrhizal mantles. (from Agerer, 1991).

References Agerer R. 1986-1998. Colour Atlas of Ectomycorrhizae. Schwäbisch-Gmünd: EinhornVerlag. Agerer R (1991) Characterization of Ectomycorrhiza. Methods in Microbiology 23: 25 – 73. Agerer R et al., (1996-2006) Descriptions of Ectomycorrhizae. Schwäbisch-Gmünd: EinhornVerlag. Chilvers GA (1968) Some distinctive types of Eucalypt mycorrhizas. Australian Journal of Botany. 26: 49-70. Dominik T (1969) Key to ectotrophic mycorrhizas. Folia Forestalia Polonica Seria A Lesnictwo 15, 309-328.

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Thursday August 16 Sampling ectomycorrhizal communities along an environmental gradient. Does EcM fungal community composition change in a predictable way along environmental gradients? One might expect this to be the case if different ectomycorrhizal fungi respond differently to environmental factors (eg pH, litter quality, moisture etc etc). The situation is further complicated because the availability of host species is itself one of the biotic factors that changes along environmental gradients, and which is likely to influence EcM communities. In this exercise we will lay out a transect across the transition between two host species or along an environmental gradient, and take core samples at defined locations to characterize the ECM community below ground. You will work in pairs and each pair will be responsible for providing data from a single point on the transect. These data will be used by the whole group. 1.

Once the transect has been laid out, you will be allocated a sampling location

2.

Record the plant species present in the ground vegetation at your sampling location in approx 1m2 centered on the transect. Record the identity of the five closest trees.

3.

Take a set of five cores, at right angles to the transect line, 15 cm apart with the central core on the transect line. Repeat the process 15 cm further down the transect – these are your backup cores in case the first 5 do not contain sufficient roots.

4.

Store each core separately in a polythene bag labeled in such a way that you know where each came from.

Sorting cores 1.

Soak each core in tap water for a minimum of 30mins before starting to sort. Always keep one core soaking while one core is being sorted.

2.

Empty the core and the soaking water into a 2mm sieve. Wash the core GENTLY in a water stream.

3.

Transfer the material on the sieve to a dish with some water.

4.

Transfer portions of the core to a fresh 9cm Petri dish and start teasing out the root fragments. Transfer these fragments to fresh water in a new dish and gently remove adhering soil fragments. Take care not to damage rhizomorphs or external mycelium, and try to keep clusters of similar mycorrhizas intact.

5.

Separate out the mycorrhizas on different host species - ask if you are in doubt.

6.

Amalgamate morphologically similar live mycorrhizas of each host species in water in wells in a multiwell plate. This is your initial attempt to define the morphotypes present in the sample. Live mycorrhizas are turgid over at least part of their length, and the cortex is appears light when the tip is broken.

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7.

When all the material from one core has been sorted revisit the material in each well and decide whether each well contains only one morphotype and whether any two wells contain the same morphotype, using the criteria you learned yesterday and described above.

8.

Remember that some morphotypes will be present on both host species and some may only be present on one.

9.

We need to ensure that each group uses the same classification for morphotypes, so that the data from the all cores are comparable. To do this we will photograph common morphotypes, project the images, and attempt to ensure that every group gives that morphotype the same name.

10.

Record the number of root apices of each morphotype.

11.

Complete 5 cores containing more than 10 apices.

12.

Once the morphotype classification for the whole group has been agreed, enter the data for each core on a common spreadsheet as instructed.

13.

We need to make voucher collections and select representative tips of each major morphotype. These tips will be processed to obtain sequences during the second week of the course. Four tips of each morphotype, each supported by a voucher collection, should be taken from different cores and, where appropriate, from different hosts and placed in buffer as previously described.

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Analysing the data Each group will be provided with a consolidated data set of morphotypes as a number of spreadsheets. The first two are Excel spreadsheets. The first spreadsheet has absolute abundances of mycorrhizas in each core Transect location 1

Core

Host

1

Pine Beech Pine Beech Pine Beech

2 3

Morphotype 1

Morphotype 2

etc

etc The second has relative abundances for each host species at each transect location Transect location 1 2 3

Host

Morphotype 1

Morphotype 2

etc

Pine Beech Pine Beech Pine Beech

The other two spreadsheets are Lotus spreadsheets for use with the ordination routines in PCORD. The first has the relative abundance data. This is called the main matrix. 1P = pine ectomycorrhizas at transect position 1, etc. xx xx 1P 2P 3P 1S 2S 3S etc

samples morphotypes Q Morphotype 1

Q Morphotype 2

etc

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The second is in the same format but contains other information about the samples. This is called the second matrix. xx xx 1P 2P 3P 1S 2S 3S etc

samples variables C Host 1 2 1 2 1 2

C Forest type 1 1 1 1 1 1

etc

1.

Tabulate the total number of morphotypes, the number of morphotypes common to both hosts, and the number of morphotypes restricted to each host.

2.

Plot the species richness of each host at each location on the transect as a bar chart.

3.

Calculate the overall relative abundance of each morphotype on each host for the whole community using EXCEL spreadsheet 1. Plot as a ranked bar chart.

4.

Calculate the frequency of each morphotype on each host using EXCEL spreadsheet 2. Plot as a ranked bar chart.

5.

Import the Lotus spreadsheets to PC-ORD and compute the EcM morphotype - area curve for each species as described in the instruction sheet.

6.

Ordinate the data and graph the output as described in the instruction sheet.

Additional analyses There are many other things that can be done with these data eg. 1. Examine the relationship between the number of tips in a core and the number of morphotypes recorded for that core. 2. Is the similarity between cores at a transect location greater than the similarity between cores at different locations? This tells us something about the scale of spatial patterns in the EcM community. 3. Test for specificity and preference of morphotypes between hosts by comparing presence/absence or relative abundance in cores where roots of both hosts are found. 4. Compare the axis scores of the ordination of the mycorrhizal data with the axis scores for the ordination of the ground vegetation data. Are the mycorrhizal fungi and the ground vegetation responding to the same gradients?

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Friday August 17th Much of this day will be continuing with the field samples and analysis from yesterday

Virtual mycorrhizas – a sampling exercise In order accurately to describe a community of mycorrhizas we need to be able to identify the species present, and to determine their relative abundance. We cannot count all the individuals, so some sampling scheme is necessary. The challenge is to deploy the available sampling effort (ie the number and size of the samples you can handle) in the most effective way. Sampling communities of ectomycorrhizas presents particular challenges. Because of the time consuming nature of the exercise, only a small proportion of the individuals in the community can be enumerated ie the sampling % is low. We might expect this to introduce uncertainty to the estimates of community composition and structure. The spatial arrangement of individual species differs. Some species will confined to discrete parts of the study area, others will be dispersed throughout. Some species will occur as individual tips others will be aggregated. This means that aggregation of individuals probably occurs at a nested series of scales – but we have no way of telling, when we look at a study site, what those scales are. Yet a sensible sampling scheme should take account of the spatial ecology of the individuals in the community. Some species will be present at low relative abundance (this does not mean few root tips!!), and individuals of these species may also be aggregated. A low sampling effort is likely to underestimate, or completely miss, these species. In this exercise we will look at some of these issues by sampling a virtual community of mycorrhizas, the species composition, species relative abundance and species spatial structure of which is known. A separate instruction sheet will be provided.

Saturday August 18th We will have a few lectures and then close the lab and leave Kristiansminde field station. Arrival to Copenhagen is estimated around 16.30-17.00. You will be accommodated and then you will have the evening off.

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Part 2 – Molecular lab course at University of Copenhagen

morphotypes -

sporocarps identification?

PCR RFLP

RFLP types identification?

identification?

DNA sequences sequencetypes – identification?

In the previous week we have looked at how to identify ectomycorrhizal fungi from their sporocarps and from their vegetative structures. This week we will continue with the same samples and apply molecular tools for the identification. We will extract DNA from sporocarps and ECM root tips and from this we generate DNA sequences. The RFLP part of the diagram above is included as many published studies used this method as the primary identification tool and because the basic principle is still used in T-RFLP. RFLP can also be a valuable tool for screening many samples before selecting samples to be sequenced e.g. in connection with cloning. In the present course we will though go directly from PCR to DNA sequencing.

Sunday August 19th Today, we are going to extract DNA from sporocarps and ECM root tips and set up and run PCR. Read the principles and procedures below before starting up the student exercise.

DNA extraction using the CTAB-chloroform-isopropanol extraction method (adapted from Gardes et al. 1996) – description of the method 1. First the material stored in 2 x CTAB is softened by alternating freezing and thawing 2. the samples. Alternatively samples can be softened by incubating them e.g. for 15 min at 65°C. This softening step is often a good idea with root tips and if using a pestle to

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3. 4. 5. 6. 7.

crush the sample. Using a “bead beater” like the Mixer Mill from Qiagen makes this step unnecessary. Samples are then incubated for ½-1 hour at 65°C to inactivate enzymes that may influence our extraction Samples are mixed with chloroform and a interface containing complex bound CTABpolysaccharides-proteins are formed and can be removed DNA is precipitated with isopropanol and can be pelleted after centrifugation Then the preparation is washed with ethanol, pelleted and dried Finally the DNA pellet is resuspended in water or in a weak buffer and is ready for PCR

The reason for using this rather simple DNA extraction method instead of some commercial kit is that the quality of the DNA extracted in our experience is fine for standard PCR targeting ribosomal genes. Additionally, with the relative few samples each group will have to process, we do not find it necessary to adapt any “high throughput procedures” an option that that some of the commercial kits offer. As an alternative to the CTAB method many different commercial kits are available for DNA extraction. The most commonly used are the ones offered by the company Qiagen. The chemistry behind most commercial kits relies on silica membranes (like Qiagens DNeasy kits) which binds DNA at high salt/high ethanol concentrations. While the DNA is bound to the membrane other inhibitory molecules can be washed away with high salt/high ethanol washing buffers. In the end, DNA is eluted by water or a low ionic strength buffer. Alternatively to the silica membranes, silica coated magnetic particles may be used (e.g. Qiagens MacAttract kits) but the chemistry is the same as for the membranes. Both types of kits are routinely used in the lab in Copenhagen and can be demonstrated in you are interested. Mycelia from sand filled in growth mesh bags are nicely extracted with the CTAB method but if you wish to extract DNA from soil you will have to adapt some appropriate soil DNA extraction kit.

Reagents and equipment needed for the CTAB extraction The below list of reagents and equipment should be assembled prior to DNA extraction and kept orderly and within reach in sufficient supplies before beginning the extractions. 1. Gloves 2. Clean Eppendorf tubes 3. Clean pestles 4. 2 x CTAB buffer 5. Chloroform (in fume hood) 6. Isopropanole, ice cold (stored at -20°C) 7. Ethanol, ice cold, 70% (stored at -20°C) 8. Pipettes and tips 9. Ice 10. Microcentrifuge 11. Vortex mixer 12. 65°C water bath

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!

Use gloves during DNA extraction - Our skin contains RNAses and DNAses that may contaminate and degrade our DNA samples. Additionally, we all have a fungal community associated to our skin and using gloves during the extraction step is therefore important to protect our DNA samples against both RNAses and DNAses as well as against contaminant fungal DNA. Before beginning the extractions you must always know the proper security handling information of the chemicals to be used in the protocol. In this protocol they are the following: • 2 x CTAB buffer: handle with gloves. Note that in our CTAB buffer we do not include any β-mercaptoethanol as many often does which make it a bit friendlier. • Chloroform: handle with gloves and within a fume hood • Isopropanol: handle with gloves and within a fume hood

!

Remember, all equipment is sensitive to contamination. If you are uncertain at any point if the equipment has been contaminated or not, tell someone! This is important for all work in the lab.

Recipe for 2 x CTAB Final concenentration 100 mM 1.4 M 20 mM 2% 0.2%

Stock solution 1M 5M 0.25 M

For 100 ml Tris pH 8.0 10 ml NaCl 28 ml EDTA 8 ml CTAB 2g ß-mercaptoethanol 200 μl DH2O 54 ml 1-3% SDS can be added. (ß-mercaptoethanol is optional and not used in the present course. If used it should be added the day of extraction)

Student exercise – DNA extraction Work in groups of 2. During the field course, you have added cleaned ECM root tips or pieces of fungal sporocarps to Eppendorf tubes containing 300 μl 2 x CTAB buffer. We are now going to extract DNA from these samples. Each group is going to extract DNA from 20 samples plus two extra negative controls (11 samples each). Make a list (1-22) with the sample name (voucher ID) and pre-mark your 22 Eppendorf tubes.

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DNA extraction using the CTAB-chloroform-isopropanol extraction method protocol 1. Put material in 300μl 2 x CTAB 2. To soften tissue: freeze-thaw sample three times or let sample incubate 15 min at 65°C before crushing (optional) 3. Crush sample with clean pestle 4. Incubate for 1 hour at 65°C 5. Meanwhile, label new 1.5 ml Eppendorf tubes 6. Add 300μl chloroform and vortex 7. Spin for 15 min at 13,000-20,000 g 8. Transfer upper face (200μl) to the labeled Eppies 9. If the upper face is not clear, go to step 5, otherwise go to step 10. 10. Add 400 μl -20˚C isopropanol 11. Invert tubes a few times and incubate for 1 hour at -20˚C (this could fit with your lunch break but you may also incubate them overnight) 12. Spin tubes for 10 min 13,000-20,000 g, discard supernatant 13. Wash pellet with -20˚C 70% ethanol 14. Spin for 5 min at 20,000 g, discard supernatant 15. Oven- (65°C) or air dry samples 16. Re-suspend DNA in 50μl TE buffer (THIS IS YOUR DNA STOCK) 17. Find the appropriate dilution for PCR (usually 5-100X will work)

Polymerase Chain Reaction (PCR) - Principles (Illustrations and explanations below is partly from the homepage of Andy Vierstraete with permission; http://allserv.rug.ac.be/~avierstr/) With PCR it is possible to make a large number of copies of a piece of a genome, which is a prerequisite to have sufficient starting template for sequencing. PCR consists of three major steps that are repeated for 30 - 40 times (cycles). This is done on an automated cycler (the PCR machine), which can rapidly heat and cool the tubes containing the reaction mixture. The reaction mixture contains DNA, primers, dNTPs, polymerase (enzyme) and a buffer. PCR is an exponential reaction: on DNA copy becomes two, which becomes 4 and so fourth. Try to calculate the numbers of DNA copies after 35 cycles!

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Figure 1: PCR - the three steps (Andy Vierstraete with permission). •

Step 1: Denaturation (94°C): Denaturation yields single stranded DNA (the double stranded DNA helix melts and separate) and enzymatic reactions terminate.



Step 2: Annealing (55°C): Primers that fit (match the template DNA) will attached (anneal) to the single stranded template. If the primers fit well, stable bound last long enough for the polymerase to attach. The polymerase will be able to start copying the template.



Step 3: Extension / Synthesis (72°C): The ideal working temperature for the polymerase is 72°C. During the extension (also referred to as DNA synthesis), the bases that are complementary to the template will be coupled to the primer on the 3' side (the polymerase adds dNTP's from 5' to 3', reading the template from 3' to 5' side, bases are added complementary to the template)

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Primers for amplifying and sequencing fungal rDNA ITS region For which primers to use you may visit the UNITE homepage (http://unite.ut.ee/index.php). For more extensive lists of fungal primers follow the links to Bruns- and Vilgarys labs from the AFTOL (Assempling the Fungal Tree of Life) homepage (http://aftol.org/). PCR primers: The most widely used primers for PCR-based identification of ectomycorrhizal (ECM) fungi are the primer-pairs ITS1-ITS4, ITS1F-ITS4 and ITS1FITS4B. However, using default annealing temperature of 55oC, we have met several difficulties using all of the primer pairs mentioned above. •

ITS1-ITS4 is a pair of universal primers which co-amplifies angiosperm DNA (but gymnosperm DNA is usually not amplified). In case the DNA is extracted from sporocarps, pure cultures, or ECM of conifers, this primer pair amplifies the highest amount of target (ca 600 bp). To our knowledge, ITS1-ITS4 can amplify DNA from all (ECM) fungi.



ITS1F-ITS4. ITS1F is a fungal specific primer. However, together with ITS4, ITS1F can result in weak amplification of highly concentrated pure angiosperm DNA. We have not observed plant bands on gel (slower) if DNA is extracted from ECM. A problem with ITS1F-ITS4 is gel smearing, which is the production of numerous hardly separable bands. Raising annealing temperature a few degrees might resolve this problem. ITS1F tends to result in less target-DNA compared to ITS1, but it has proved efficient in amplifying all ECM fungi and is the standard primer combination in most ECM community studies.



ITS1 or ITS1F together with LR21 or other reverse primers upstream in the LSU have the advantage that the, in comparison to the ITS region, more conservative LSU region can be sequenced. This may improve your possibility to identify your sequences later e.g. when using BLAST (see later).



On the course, we will use the primer combination ITS1F-ITS4.



Primer

ITS1 ITS1F ITS2 ITS3 ITS4 ITS4B LR21

Sequence 5` -> 3`

TCCGTAGGTGAACCTGCGG CTTGGTCATTTAGAGGAAGTAA GCTGCGTTCTTCATCGATGC GCATCGATGAAGAACGCAGC TCCTCCGCTTATTGATATGC CAGGAGACTTGTACACGGTCCAG ACTTCAAGCGTTTCCCTTT

Tm

57 55 57 57 53 67 54

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Figure 2: The central transcribed part of the nuclear ribosomal gene cluster. Approximate position of commonly used PCR and sequencing primers are indicated.

Student exercise – PCR: Continue to work in the groups of 2. You now have now 22 DNA extracts (hopefully two are without DNA) and you are now going to set up 24 PCR reactions (11 each plus one extra negative non-DNA control). Depending on how much PCR product you need for downstream processing you may adjust the PCR reaction volume between 10-100 μl here we will use a 30 μl setup. The following should be assembled prior to use and kept orderly and within reach in sufficient supplies before starting. 1. 2. 3. 4. 5. 6. 7. 8. 9.

DNA-template (10 times diluted work solution) Two primers, 10 μM Ice 0,2 ml tubes for reactions sterile sdH2O dNTPs (GATC) PCR buffer Taq polymerase enzyme Pipettes and filter tips

In order to avoid contaminating your reactions, work clean and carefully, use gloves and filter tips, always change tips between steps, and include a negative PCR control (a tube containing everything, but sterile water instead of DNA template).

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PCR Protocol for a 30 µl setup Prepare a master mix with the following ingredients per tube: • 3 μl PCR (TQ) buffer • 3 μl forward primer (10μM) • 3 μl reverse primer (10μM) • 7.5 μl H2O • 12 μl GATC mix • 0.15 μl Taq polymerase When calculating the total amount of master mix needed, remember to include a negative control (no DNA added) and some extra volume because you do not want to run dry on master mix before the last sample (one extra for each ten tubes works fine). • Distribute 28.5 μl of the master mix to the PCR tubes and add 1.5 μl DNA template to each tube • Run the PCR immediately after • Use the following PCR program: Step Denaturation Denaturation Annealing Extension Extension Soak •

Temperature (˚C) 95 ˚C 95 ˚C 55 ˚C 72 ˚C 72 ˚C 10 ˚C

Time (min:sec) 5:00 0:30 0:30 1:00 7:00 For ever

Cycles 1 35 1 1

Transfer the PCR tubes to the PCR machine and run the PCR program.

We have access to a number of PCR machines which we will start up independently as teams finishes their work. This will also give us some flexibility in the post PCR work as space is limited for electrophoresis When the PCR are finished you will check your PCR reactions on a 1.5% agarose gel and illuminate your PCR with UV light but this is tomorrow.

Monday August 19th Principles for electrophoresis Electrophoresis is a process where charged molecules move in a medium through an electric field. The movement of the molecule depends on its charge, shape and size. Electrophoresis of macromolecules is usually done by adding a certain amount of such molecules to a porous matrix. Different kinds of molecules will move with different speeds through the matrix. When the separation is complete, the molecules can be detected as bands at different positions in the matrix. The matrix, in this particular case, consists of an agarose gel. It has also a secondary separating effect, as the inner web like structure in the gel works as a sieve for different sizes of molecules - affecting their speed of movement.

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Normally, electrophoresis is a result of the total molecular charge in relation to the charge of the electrical field. A protein, which has a certain positive or negative charge because of its either acidic or alkaline elements, will within a certain time reach its isoelectric equilibrium. The isoelectric state can only be reached if the medium where the electrophoresis takes place has enough buffering capacity and the matrix is big enough, which is true for most proteins. But for a nucleic acid this is often not the case. Because of its very acidic composition it seldom reaches its isoelectric state in an electrophoresis medium with normal buffering capacity. It will continue to move beyond the limits of the matrix. Therefore, the electrophoresis has to have a time limit, and the result detected after that limit is reached. The electrophoresis solution must be of such a kind that it keeps its buffering capacity for the whole separation time. Constant formation of H+ and OH- at the anode and cathode will decrease the buffer capacity, which in turn affects the speed of separation and overall quality of the result. Agarose gels consist of highly refined polysaccharides from agar. They are available in a wide spectrum of melting temperatures, gel strength and transparency. Agarose most often comes as a powder, soluble in boiling liquid. Gels normally polymerize at about 40˚C, and once polymerized it can withstand temperatures below 100˚C without melting. Density is decided by adjusting the concentration of agarose in the gel. Higher concentration gives a denser matrix. Normally, the concentration is kept between 0.4% and 4%. In our case the concentration is 1.5%, which easily allows the migration of PCR products. Prior to loading the gel the DNA is mixed with a loading solution. This makes the samples sink readily in to the loading wells. Separated bands of nucleic acids are stained with ethidium bromide (extremely carcinogenic!). Ethidium bromide binds to the DNA making it fluorescent in UV-light. UV-light has a degenerating effect on DNA molecules. Some also run their extracted DNA on an agarose gel (then it is typically a low concentration gel because of the high molecular weight of the extracted DNA). This allows you to judge upon the quality of the extracted DNA which can be important if the extracted DNA is used for establishing a clone library etc. In the presents course we will not check our extracted DNA by electrophoresis (unless you have some extra time and would like to try this).

Student exercise: Electrophoresis - Checking the PCR result Continue to work in the groups of 2. You now have 24 PCR products, 12 each. These products shall only be worked with in the Post-PCR rooms and with Post-PCR pipettes. PCR products are concentrated DNA that represents a high contamination risk and should be kept strictly apart from rooms and equipment used for DNA-extraction and PCR setup. It is therefore strictly forbidden to bring PCR products into the PRE-PCR lab (“green lab – room 02-010) where we isolate DNA, and it is also strictly forbidden to use pre-PCR pipettes for PCR products. Post PCR pipettes are available in the Post PCR room. The Post-PCR electrophoresis lab is space limited, so we will have to work a few teams at a time. Depending on if you are in the fast or the slow track you will pour you own gel, share a gel with another group or we will put all your samples on common large gel.

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1.5% Agarose gel •



1

Prepare the electrophoresis tray and combs. We have three different sizes: A small system that may hold 12-36 samples (including the size standard), a medium system that can hold up to 72 samples (including the size standard) and a large system that holds 192 samples (excluding the size standard). Take out 1.5% agarose from the 65°C oven. This solution already contains ethidium bromide. Cast the gel and wait for about 20 min for it to solidify. EtBr (Etidium Bromide) is highly carcinogenic because it intercalates with the bases in the DNA strand. It is crucial that you are not in contact with EtBr.

Protocol for detection of PCR-products 1. Transfer the agarose gel to the electrophoresis system and fill it with 1xTBE buffer (running buffer) to cover the gel. Remove the combs. 2. Mix 2 µl loading solution and 5 µl PCR-product in a multi-well plate. 3. Load the mix of PCR product and loading buffer to the wells/slots in the agarose gel (note where the different samples are loaded in a separate sheet). Since the mix is slightly viscose it should be loaded slowly and carefully into the slot. In each lane on the gel, the last well should be loaded with 3 µl DNA length-marker (phiX). 4. Initiate electrophoresis on 160V for 15-20 minutes. The separation time is important to how well secondary bands can be detected. Longer separation time gives a more easily interpreted result. Shorter time gives a faster but probably less reliable result. 5. Transfer the gel to the UV-detector cabinet. Lock the cabinet door, view the results using the Kodak software on the computer next to the electrophoresis bench.

.

DNA fragments on a gel visualized with UV light

PCR products

DNA marker

Important – check that the negative PCR control is negative (no detected bands). If the DNA extraction control is positive, we have had contamination problems during the extraction procedure, if the negative PCR control is positive we have had contamination problems during the PCR setup. We are now going to evaluate the products and decide which ones can be submitted to further DNA sequencing. We will try to sequence all samples where amplification has been successful in both (forward and reverse) directions.

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Purifying PCR products and measuring the DNA content Before our PCR products are ready to send out for sequencing we need to purify the PCR products and measure and adjust the DNA concentrations. Purification of PCR products means that you will remove everything from the amplification tubes except DNA i.e. buffer, enzyme, nucleotides and primers should be removed. This can be done in several different ways. Some uses a similar chemistry as was explained under the DNA extractions section (e.g. Qiagens Qiaquick kit) but in Copenhagen we use a filter membrane system that simply allows buffer, enzyme, nucleotides and primers to be washed out but hold back the PCR product (we use a NucleoFast 96 plates from Macherey Nagel but many other similar kits are available from other manufactures e.g. from Millipore).

The ultra filtration method for purifying PCR products as used e.g. in the NucleoFast 96 system.

Student exercise: Getting your PCR products ready for sequencing 1. Make sure the Eppendorf spectrophotometer (in the Mol Bot lab) is turned on 2. In the Exp Myko lab, Add 50 µl TE buffer to each tube you are sequencing from (to wash down any DNA sticking to the sides of the tubes) 3. Transfer your DNA to your designated NucleoFast wells. Cover wells not used with transparent plate seals 4. Turn on the vacuum and wait until the membranes are dry (typically about 5 min but check – vacuum must not exceed 20 “in. Hg vac.”) 5. Add 50 µl water to each well and shake with plate shaker for 5 min at 600 rpm. 6. Transfer the “elute” to new tubes. THIS IS YOUR STOCK OF PURIFIED PCR PRODUCT 7. Make a 10 fold dilution of the elute (5 µl elute in 45 µl water) in new tubes. These we will use for measuring the DNA concentration with the Eppendorf spectrophotometer. 8. Remember to type in the dilution factor and start by running a blank against water (the same stock that was used to dilute samples). As a control, run water as the first and last sample. Do not change cuvete within a run.

Sequencing of PCR Products using MWG Biotech

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1. In a spreadsheet, input DNA concentration and length of PCR products and calculate the optimal amount of purified PCR product for sequencing. MWG recommend using 20 ng DNA (+/- 10 ng) per 100 bp. 2. Pipette the calculated amount to a new 1.5 ml Eppendorf tube and dry samples at room temperature or for approx. 1 hour at 65°C. Alternatively transfer the samples to a microtitter plate (if 48-96 samples). IF WE USE PLATES OR SINGLE TUBES DEPENDS ON HOW MANY SAMPLES THAT WERE SUCCESSFUL IN THE PCR 3. Go to your (Rasmus’) account at MWG 4. Choose “sequencing”, then click on “Manage Primers” and “Enclose primers” 5. For each primer, input name and concentration (use 10µM), barcode number and choose “value read” from the pull down menu. 6. After the last primer click “Done” and in the “Sequencing” menu you choose either “Value Read” or “Plate order” depending on the number of samples you plan to sequence. 7. For “Value Read” orders: input barcode label, sample ID and primer to be used. When done, click “save” and review your data and then go to exit. 8. Use your personal (Rasmus’) MWG barcode labels to label tubes and Primers and send the tubes or plates in a padded envelope (for the present course we will use an express mail service) together with 10-20 µl of each relevant primer (10 µM) to: MWG Biotech AG Fraunhoferstr. 22 82152 Martinsried GERMANY See http://ecom2.mwgdna.com/improve_results_seq/improve_results.html for many good suggestions for improving your results and for viewing your data

Tuesday August 21th It is not possible/wise/allowed to make sequencing a student exercise. Instead, we have sent our purified products away to the Germany based company MWG-BIOTECH (http://www.mwg-biotech.com/html/all/index.php) to be sequenced. This is therefore a day where we can allow a relaxing sightseeing for some hours after some lectures and student presentations. We also routinely send our PCR products to Korea to be sequenced by the company MACROGEN (http://www.macrogen.com/eng/sequencing/sequence_main.jsp).

Principles for DNA sequencing (Illustrations and explanations below is partly from the homepage of Andy Vierstrate with permission; http://allserv.rug.ac.be/~avierstr/) DNA sequencing is a method for determining the order of the nucleotides of a piece of DNA, in our case internal transcribed spacers (ITS1 and ITS2) between genes the small and the large ribosomal subunits. For sequencing, we need a large and pure amount of target DNA. Therefore, we cannot start from genomic DNA, but mostly from PCR generated fragments or cloned genes. Automated sequencing uses cyclic sequencing. To the sequencing reaction, both normal and labeled nucleotides are added.

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The sequencing reaction: Similar to PCR, there are three major steps in a sequencing reaction, and these steps are repeated for 30 or 40 cycles (for environmental samples like a ECM root tip, it is not recommended to exceed 35 cycles because that tend to amplify minor amount of other organisms/fungi present in the PCR product, which in turn yield unreadable sequences or background noise. 1. Denaturation at 95°C. 2. Annealing at 50°C : Only one primer is used and consequently only one strand is copied. 3. Extension / Synthesis (60°C): The ideal working temperature for the polymerase is 72°C. However, in the sequencing reaction, not only dNTPs but also chemically modified ddNTPs (dye labelled) are to be incorporated. The temperature is lowered to 60°C to enhance incorporation of the modified molecules (lowered temperature lower the specificity). During the extension the bases complementary to the template are coupled to the primer on the 3'side. The polymerase adds dNTP's or ddNTP's from 5' to 3'. Reading the template from 3' to 5' side, bases are added complementary to the template. When a ddNTP is incorporated, the extension reaction terminates because a ddNTP contains a Hatom on the 3rd carbon atom (dNTP's contain a OH-atom on that position). Since the ddNTP's are fluorescently labelled, and it is possible to detect the color of the last base of this fragment on an automated sequencer.

Figure 3: Cyclic sequencing - the three steps (Andy Vierstraete with permission).

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There is a linear increase of the number of copies of one strand of the gene since only one primer is used (one strand copied during sequencing). Hence, a large amount of copies of the gene/DNA fragment is required in the starting mixture for sequencing. Suppose there are 1000 copies of the desired DNA fragment in the start mix. After one cycle, there will be 2000 copies (1000 original templates and 1000 complementary strands with each one fluorescent label on the last base). 30 cycles yield 30 000 complementary strands etc.

Figure 4: The linear amplification of the gene in sequencing (Andy Vierstraete with permission). Sequencing at MWG-BIOTECH is performed on ABI PRISM 3700 capillary sequencers using Big Dye termination chemistry. Read more about both on Applied Biosystems homepage: https://products.appliedbiosystems.com/ab/en/US/adirect/ab?cmd=catNavigate2&catID=600522. Before the PCR products can be sequenced successfully, it is important to purify them. The products contain the amplified DNA, but also undesired left-over: primers, nucleotides, polymerases and salts. Separation of the molecules The sequencing product (the mixture of strands of different length and all ending on a fluorescently labeled ddNTP) are separated using capillary electrophoresis. Capillary array electrophoresis uses narrow-bore capillaries filled with a separation matrix to resolve DNA sequencing fragments. An electric field is used to cause the DNA fragments to migrate into (electrokinetic injection) and through (electrophoresis) the capillaries. Similar to gel electrophoresis, the DNA fragments are separated by size, with the shorter fragments moving faster (peaks earlier in the electropherogram) than the longer fragments (peaks later in the electropherogram). The large surface-to-volume ratio of the capillary efficiently removes the heat generated during electrophoresis. Each capillary has a clear detection window located at a fixed distance from the sample loading point through which the samples are scanned with a laser.

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Wednesday August 22th Student exercise: Post – molecular work / computer lab Based on the text and illustration on the next pages, you will be able to go through all sequences of your responsibility and do as described below: o Evaluate chromatograms o Make reliable consensus sequences o Make two sets of files in BioEdit: One with the two original strands + consensus (original data) and one with the confirmed sequence (the hopefully reliable consensus sequence) alone. Save the latter file in BioEdit and use the sequence ID as file name. o Make also a multisequence file (alignment) containing all your sequences o Annotate the sequence o Exchange sequences with the other students (will be organized). o Collect all sequences in one file and make your own sequence database. o Conduct local BLAST for all sequences of your responsibility to find potential similar and identical sequences. o Is there any match between sequences from sporocarps and ECM roots? Read the information below before proceeding. We will receive the sequences as individual *.scf files that can be viewed, evaluated and edited in different programs (e.g. Sequencher, CromaPro and BioEdit). We will use BioEdit (free ware program) on this course.

BioEdit - biological sequence alignment BioEdit is a biological sequence alignment editor written for Windows 95/98/NT/2000/XP (made by Tom Hall). It is one of the best multialigning programs available for PC. For Mac, other programs are available. BioEdit provides an intuitive multiple document interface with convenient features that makes alignment and manipulation of sequences relatively easy on your desktop computer. Several sequence manipulation and analysis options and links to external analysis programs facilitate a working environment that allows you to view and manipulate sequences with simple point-and-click operations. You can download the program for free at http://www.mbio.ncsu.edu/BioEdit/bioedit.html and also read more about it here. Interpretation and evaluation of sequence chromatogram in BioEdit When you open an *.scf file of the sequence in BioEdit you get the sequence and the chromatogram:

Unreliable sequence - exclude

Reliable sequence - include

As shown on the figure, the beginning of the sequence is always messy (dubious) and must be excluded. If the sequencing has been successful, the sequence turns better quickly and is hopefully of good quality and reliable. Each base must be checked and confirmed.

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Above: This is a perfect sequence. All curves are clear and regular and the sequence is reliable and can safely be confirmed. Below: Towards the end of the sequence, the quality is reduced, and it is not easy to determine the base order anymore:

This area is unreliable It is now required to have a sequence of the opposite strand (sequenced in the opposite direction) to determine the actual sequence. Comparing forward and reverse sequence – how to make a consensus sequence 1. Copy forward and reverse sequence into a common BioEdit file as shown below: 2.

3. Chose the “Accessory Application” menu and “CAP contig assembly program” (as shown below) and accept “run application”. Push “enter” once to finish the analysis. 4.

5. You will now receive a contig sequence (consensus sequence) where both strands (sequences) are taken into account.

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6.

7. 8. 9. 10. 11. 12. 13. Consensus sequence Disagreement between the two sequences, these bases must be confirmed manually 14. 15. 16. The consensus sequence must be checked base for base (when disagreement between the two strands) and corrected manually. The two chromatograms must be compared simultaneously. This part is not automatically done in BioEDIT (Sequencher is much better in this respect: www.sequencher.com). However, by manually arranging the different windows in BioEdit, the chromatograms can be compared and consensus sequence corrected. a. To compare the sequences, the reverse chromatogram (representing the reverse strand) most be converted to “forward” sequence by choosing “reverse – complement” from the “view” menu. The sequence is then reversed (the start becomes the end and vice versa) and the bases are converted to complementary bases (A→G, G→A, C→T and T→C) b. To find a certain motifs, use “find” from the “edit” menu (see below): 17. 18. 19. 20. 21. 22. 23. 24. 25. 26. 27. 28. 29. 30. 31. To correct the sequence it can be a good idea to make a copy of the consensus (contig) sequence and rename (use “copy sequence” and “paste sequence” from the edit menu, and double click on the name to rename). The new sequence can be edited manually based on evaluation of the chromatograms (to edit, chose “edit” mode and “insert” mode). Below, an example:

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32.

Incorrect bases removed in the corrected contig sequence, reverse strand is trusted here

Unreliable, A incorrect

Reliable

Unreliable

Reliable

Confirmed and reliable sequences should be saved in a new file containing all your confirmed sequences. Sequence annotation in BioEdit It is possible include a lot of information about the sequence in BioEdit, and in this way store important annotation together with the sequence (in addition to the XL-file made for each collection). Double-click in a sequence title. In the window you receive there is a red arrow. Click on the arrow to view and edit the annotation form for each sequence. But behold that if you at some point choose to copy your sequence as a FASTA file all information except the sequences title and the sequences is lost! Local BLAST in BioEdit You can make your own sequence database in BioEdit where you can perform local BLAST searches. This is very useful if you have a large dataset of unpublished sequences and would like to find closest matching sequence in your own dataset. To create a local protein or nucleotide database for BLAST searching, you need only have a Fasta-format file containing all of the sequences you want in the database. Nucleotide and protein sequences cannot be mixed within the same database. From the “Accessory Application” menu, choose “BLAST”, then “Create a local ... database file”. You will be prompted for the input Fasta file. The rest is automatic. The database will be placed in to the \database folder of the BioEdit install directory. The new database should appear in the appropriate database list box of the local BLAST interface form. To use local BLAST from within BioEdit, highlight the title of the query sequence from within a BioEdit document. Next, choose “Local Blast” from the “Blast” menu under the “Accessory apps” menu. Don’t worry about gaps, these will be removed automatically. You may also choose several sequences at once if you want to for a batch job. Choose the

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program you would like to use, then the database to search (the window will appear as below):

In the upper right of the form, there will be a drop-down list for both nucleotide and protein databases. Choose the one you want for the appropriate type, and don’t worry about the other type (a selected choice of nucleotide database will be ignored when doing a protein search and vice versa). You may choose whether to save the output to a user-named file, or simply have BioEdit create a temp file that is automatically opened when the search is done

Sequence alignment in BioEdit There may not be time to do alignments, but below we describe the principles. To compare two or more sequences, it is necessary to align the conserved and unconserved residues across all the sequences (identification of locations of insertions and deletions that have occurred since the divergence of a common ancestor). These residues form a pattern from which the relationship between sequences can be determined with phylogenetic programs. When the sequences are aligned, it is possible to identify locations of insertions or deletions since their divergence from their common ancestor. There are three possibilities: •

The bases match: this means that there is no change since their divergence.



The bases mismatch: this means that there is a substitution since their divergence.



There is a base in one sequence, no base in the other: there is an insertion or a deletion since their divergence.

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Figure 8: The comparison of sequences. A good alignment is important for the construction of phylogenetic trees (will not be covered in detail on this course). The alignment will affect the distances between 2 different species and this will influence the inferred phylogeny. There are several programs available on the net for aligning sequences. These are all based on different mathematical models to compare two or more sequences with the most optimal score for matching bases with a minimum number of gaps inserted (because you can insert a huge amount of gaps, so every base will match an other). In BioEdit it is possible to conduct alignment of huge datasets (multiple alignments). Automatic multiple alignment using the program ClustalW is integrated in BioEdit and is good to use in combination your own brain (manual adjustments are always needed after an automatic aligning in ClustalW). Below, there is an unaligned sequence matrix of closely related genotypes:

Above: unaligned sequence matrix of closely related fungal genotypes.

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Chose ClustalW multiple alignment for a first automated aligning of the whole sequence matrix.

ClustalW aligns the sequence matrix quite well, but manual adjustments are always needed.

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After manual adjustments, you have a “perfect” alignment ready for submission to Phylogenetic analysis. Read more about phylogenetic analysis on the PAUP homepage (http://paup.csit.fsu.edu/index.html).

Thursday August 23th Molecular Identification using public sequence databases During this day you should spend time on molecular identification using public sequence and identification databases: UNITE (Userfriendly Nordic ITS Ectomycorrhiza-base). UNITE (http://unite.ut.ee/index.php) is a rDNA ITS sequence database focused on ectomycorrhizal asco- and basidiomycetes. The sequences are generated from sporocarps collected and identified by specialists and deposited in public herbaria; type specimens are used whenever possible. In the future we also wish to include sequences from environmental samples so it e.g. will be possible to “ask” the database give me all sequences associated with Norway spruce from the database. Selected species also have full descriptions and illustrations linked to the sequences. The purpose of the database is to facilitate identification of environmental samples of fungal DNA. Sequence-similarity searches against UNITE are performed using the BLAST software (“Run Analysis”). BLAST outputs have links to the full database records including information on the fruit body from which the DNA was isolated (details on collection, information on identifier, location of deposition, contact person(s), and so on). When you perform your BLAST search you may choose to tick the UNITE+INSD box. This means that you will search both against UNITE and sequences from INSD - the International Nucleotide Sequence Database (GenBank, EMBL, DDBJ). When receiving the BLAST results sequences from UNITE and INSD will be in different colors so you will be able to tell them apart. THE “UNITE+INSD” OBTION SHOULD BE YOUR STARDARD OBTION WHEN IDENTIFYING YOUR OWN SEQUENCES. It is also possible to query the database

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by taxonomy (“Search Pages”). Currently the UNITE database holds 2511 ITS sequences of 1046 species from 118 genera. The database covers most genera/clades of ECM fungi pretty good but currently the database is biased against sporocarp derived sequences. Therefore ECM clades not forming or forming very inconspicuous sporocarps are ill represented. Specific problematic groups of ECM fungi are: Thelephoroid fungi, Sebacina, Helotialian fungi, Cantharelloid fungi and probably more. In addition to BLAST, alternative search tools based on phylogenetic inference have been developed: The UNITE galaxieBLAST interface. Employing automated phylogenetic analysis, this facility is meant to provide an alternative view on the identity of your query sequence. It is not a replacement of but rather an addition to the UNITE BLAST facility; it should be seen as an attempt to utilize the powerful properties of phylogenetic analysis to address sequence relatedness rather than as a quick way to generate phylogenetic trees for publication, the latter being a task for which it is poorly suited. FasTa3 (http://www.ebi.ac.uk/fasta33/nucleotide.html) provides sequence similarity and homology searching against nucleotide and protein databases using the Fasta programs. Fasta can be very specific when identifying long regions of low similarity especially for highly diverged sequences. You can also conduct sequence similarity and homology searching against complete proteome or genome databases using the Fasta programs. The Fasta platform provide a great possibility for fast identification of unknown fungal DNA sequences based on all fungal sequences available in public databases. The drawback is that the reliability of the sequences is not always good. UNITE is therefore more reliable, but provide a smaller selection of ECM fungal sequences. You can also do BLAST searches against NBCI/GenBank directly from BioEdit. Simply highlight the title of the query sequence from within a BioEdit document. Next, choose “NBCI Blast over the internet” from the “Blast” menu under the “Accessory apps” menu.

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Student exercise: Identification of sequences Identification of sequences using UNITE Go to the UNITE website (http://unite.ut.ee/index.php) and follow the instructions there.

Try to identify your sequence using some of the different methods (BLAST and galaxie). UNITE is under development and do not cover all groups of ECM fungi. You may also try the FasTa search tool.

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Identification using FasTa3 Go to the webpage (http://www.ebi.ac.uk/fasta33/nucleotide.html), make sure to choose “Nucleic Acid” and “Fungi” under “Database”, and ask for “interactive” results (See the window below):

Name your sequence with sequence ID under “search title”. Copy your unknown sequence and paste it in the window directly. The sequence should be without gaps. The FasTa program requires a > in front of the sequence (see illustration). You can also upload sequences directly form a file, e.g. a BioEdit file. Select “Run Fasta3” and wait for the results. The results can be viewed with or without sequence annotation (“show/hide annotation”), as pair wise alignments (“Fasta results”), as a multi alignment (“MView”) etc. You can also identify the sequences directly from BioEdit with BLAST searches in NBCI/GenBank. Play with the different tools, identify all your sequences and make a table of species identified (closest mach to known fungus in %) If we have time, we will briefly demonstrate phylogenetic analysis using PAUP.

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Relevant web-sites for sequence analysis There are several good and relevant sites online that provide more or less useful services and tools. Below you find a list a few relevant web addresses. •

UNITE (http://unite.ut.ee/)



FASTA (http://www.ebi.ac.uk/fasta33/nucleotide.html)



EMBL-EBI (European Bioinformatic Institute): http://www.ebi.ac.uk/Information/. Of particular relevance here is o “Services” (http://www.ebi.ac.uk/services/) o “Toolbox” (http://www.ebi.ac.uk/Tools/). Fasta3 is one of several services and tools provided by EMBL-EBI.



ExPASy (The ExPASy - Expert Protein Analysis System proteomics server): http://us.expasy.org/



SRS – data and tool integration: http://srs.csc.fi/ (for accessing public sequences by accession number or taxonomy)



NBCI (National Center of Biotechnology Information): http://www.ncbi.nlm.nih.gov/ This page include among others o GeneBank (http://www.ncbi.nlm.nih.gov/Genbank/GenbankOverview.html) o BLAST (http://www.ncbi.nlm.nih.gov/BLAST/)



PAUP: http://paup.csit.fsu.edu/index.html (phylogenetic program package by Dawid Swofford including tools for inferring and interpreeting phylogenetoc trees)



BioEdit: http://www.mbio.ncsu.edu/BioEdit/bioedit.html (a biological sequence alignment editor written for Windows 95/98/NT/2000/XP). The program is freeware.

• •

Homepage of Andy Vierstraete: http://allserv.rug.ac.be/~avierstr/ (provide a pedagogic and nicely illustrated introduction to DNA-extraction, PCR, and sequencing) Sequencher: http://www.sequencher.com/ (a very good program for producing and editing contigs of two or more sequence chromatograms)

Friday August 24th This day will be used for compiling a common molecular-morhotype based dataset from our field sampled tips. Further more, we will sum up and evaluate the course etc. Departure to the airport will be arranged / explained (basically the trains leaves every 20 min from Nørreport station to the airport).