DNA Barcoding Methods for Land Plants

Chapter 11 DNA Barcoding Methods for Land Plants Aron J. Fazekas, Maria L. Kuzmina, Steven G. Newmaster, and Peter M. Hollingsworth Abstract DNA barcoding in the land plants presents a number of challenges compared to DNA barcoding in many animal clades. The CO1 animal DNA barcode is not effective for plants. Plant species hybridize frequently, and there are many cases of recent speciation via mechanisms, such as polyploidy and breeding system transitions. Additionally, there are many life-history trait combinations, which combine to reduce the likelihood of a small number of markers effectively tracking plant species boundaries. Recent results, however, from the two chosen core plant DNA barcode regions rbcL and matK plus two supplementary regions trnH–psbA and internal transcribed spacer (ITS) (or ITS2) have demonstrated reasonable levels of species discrimination in both floristic and taxonomically focused studies. We describe sampling techniques, extraction protocols, and PCR methods for each of these two core and two supplementary plant DNA barcode regions, with extensive notes supporting their implementation for both low- and high-throughput facilities. Key words: DNA barcoding, Plant field collecting, Plant DNA extraction, PCR amplification, Cycle sequencing, rbcL, matK, trnH–psbA, Internal transcribed spacer

1. Introduction The land plants encompass an enormous diversity of form and function. They consist of the seed plants (angiosperms and gymnosperms), along with the bryophytes (mosses, hornworts, and liverworts), ferns, and fern allies. Estimates of total species numbers vary greatly among authors (1–3), but a recent estimate has suggested that there are approximately 380,000 species of land plants, comprising ca. 352,000 species of angiosperms, ca. 1,300 species of gymnosperms, and ca. 13,000 species each of bryophytes and ferns/fern allies (4). The standard animal DNA barcode comprising a portion of the mitochondrial gene CO1 evolves too slowly in plants to serve as a

W. John Kress and David L. Erickson (eds.), DNA Barcodes: Methods and Protocols, Methods in Molecular Biology, vol. 858, DOI 10.1007/978-1-61779-591-6_11, © Springer Science+Business Media, LLC 2012

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useful DNA barcode (5). This has led to the search for an equivalent DNA barcode for land plants. The primary focus of this search has been the plastid genome, with many authors recognizing that multiple regions are required (5–12). Selecting a standard plant DNA barcode has been difficult, as all of the various candidate loci have different strengths and weaknesses, with no clear-cut front runners. In a community-authored paper, the combination of portions of the plastid regions rbcL and matK was suggested as the core DNA barcode for land plants (13) and subsequently provisionally adopted by the Consortium for the Barcode of Life. In addition to this core DNA barcode, other loci are often required to increase the levels of species resolution. At the 2009 International Barcode of Life Conference in Mexico City, it was recommended that the community continue to gather data from additional DNA barcoding loci to establish whether other loci should be formally incorporated into the plant DNA barcode. The two most widely used supplementary loci are the plastid intergenic spacer trnH–psbA (one of the leading contenders for the core plant DNA barcode) and the nuclear ribosomal internal transcribed spacers (ITS). The nuclear ribosomal ITS regions had previously been discounted as a standard DNA barcode due to concerns over paralogy and the presence of putative pseudogenes which lead to sequencing difficulties in many plant groups (e.g., refs. 14–18). However, the increased resolution of ITS over plastid DNA barcodes in many studies (e.g., ref. 19) suggests that it should continue to be explored as part of the plant DNA barcode, and some authors have noted that just using a subset of the ribosomal cassette (ITS2) can lead to greater amplification and sequencing success compared to the entire ITS region (20). We, therefore, include methods for all four of these regions [rbcL, matK, trnH–psbA, and ITS (including ITS2)] to provide the maximum utility to users of plant DNA barcoding. Details of other loci that have been used in plant DNA barcoding studies can be found elsewhere (e.g., refs. 5, 11, 13, 21). It should be noted that levels of species discrimination in plants with standard DNA barcoding loci are in general lower than those obtained by CO1 in many animal groups (22). This is in part due to the lower rate of nucleotide substitution in the plastid genome, but also due to other reasons, including hybridization, polyploidy, speciation via breeding system transitions, species defined on very narrow taxon concepts, large ancestral population sizes, and low levels of intraspecific gene flow for plastid markers (23, 24). These issues are not evenly distributed among all plant groups; therefore, it is expected that resolution at the species level will be reasonably good in some groups and quite poor in others. In floristic contexts where geographical limitation usually restricts the number of closely related species, rates of species discrimination are expected to be greater (e.g., refs. 25, 26). Methods are invariably open to improvement from a variety of sources, and there are often many ways to achieve the same result.

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For example, the reader may have a different way of drying plant samples or prefer to do PCR in larger reaction volumes. Where multiple methods are commonly in use, we attempt to provide details for each. The notes provided in the last section illuminate some of the principles that the methods we have provided aim to achieve. Some of the methods provided have been optimized to be cost-efficient, and are those currently in use at the Canadian Centre for DNA Barcoding (http://www.ccdb.ca/pa/ge/ research/protocols).

2. Materials 2.1. Field Collecting

1. Field press with blotting paper and spacers for voucher preparation. 2. Jewelry tags for labeling. 3. Silica gel (with 10–30% indicating silica beads). 4. Waterproof markers or pens. 5. Container(s) for silica drying of tissue, e.g., 20-ml scintillation vials, sealable plastic whirl-packs, zip-lock bags, or coin envelopes/tea bags that can be placed in a sealable container.

2.2. Tissue Sample Storage

1. Use of a climate-controlled facility if available or airtight containers filled with silica gel desiccant to archive tissue samples.

2.3. Tissue Subsampling for DNA Extraction

1. Grinding beads: for example, stainless steel 440C 3.17 mm beads. 2. Small forceps. 3. Latex or nitrile disposable gloves. 4. Ethanol: 100%. 5. ELIMINase®, DNA AWAY®, or a similar product. 6. Alcohol burner. 7. For single tube-based extractions: 2-ml screw-cap tubes with O-ring seals that are strong enough to withstand the homogenization process without breaking. 8. For plate-based extractions: Racked sterile mini tube strips with cap strips (e.g., PROgene® Mini Tube System 1.1 ml 8 Strip Pre-sterilized Mini Tube and sterile cap strips).

2.4. DNA Extraction: Single Sample-Based Extraction: Commercial Kits

1. Equipment for tissue grinding: for example, FastPrep® or TissueLyser with tube adaptor. 2. Microcentrifuge with a rotor for 2-ml tubes. 3. Vortex.

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4. Ethanol: 100%. 5. Heating block/incubator capable of heating to 70°C. 6. Pipettes and pipette tips. 7. 1.5- or 2-ml microcentrifuge tubes. 8. Individual tube-based DNA extraction kit. 9. Latex or nitrile disposable gloves. 10. ELIMINase®, DNA AWAY®, or a similar product. 2.5. DNA Extraction: Single Sample Extraction: Non-kitBased Method (Adapted from Ref. 26)

1. ELIMINase®, DNA AWAY®, or a similar product. 2. Silica-membrane spin columns (e.g., EconoSpin® mini spin columns, Epoch Life Science Inc.). 3. Equipment for tissue grinding: FastPrep® or TissueLyser with tube adaptor. 4. Microcentrifuge with a rotor for 2-ml tubes. 5. Vortex. 6. Ethanol: 100%. 7. Molecular biology grade water. 8. Heating block/incubator capable of heating to 70°C. 9. Pipettes and pipette tips. 10. Latex or nitrile disposable gloves. 11. 1.5- and 2-ml microcentrifuge tubes. 12. CTAB lysis buffer: 2% cetyltrimethylammonium bromide (CTAB), 100 mM Tris–HCl pH 8.0, 20 mM EDTA, and 1.4 M NaCl. 13. Binding buffer: 5 M guanidine thiocyanate, 20 mM EDTA pH 8.0, 10 mM Tris–HCl pH 6.4, and 4% Triton® X-100. 14. First wash buffer: 50% ethanol, 3 M GuSCN, 10 mM EDTA pH 8.0, 5 mM Tris–HCl pH 6.4, and 2% Triton® X-100. 15. Second wash buffer: 60% ethanol, 50 mM NaCl, 10 mM Tris– HCl pH 7.4, and 0.5 mM EDTA pH 8.0.

2.6. DNA Extraction: Plate-Based Extraction (96 Samples): Commercial Kits

1. Equipment for tissue grinding (e.g., TissueLyser with plate adaptor). 2. Centrifuge with a deep-well swinging bucket rotor capable of achieving 5,600–6,000 × g force. 3. Ethanol: 100%. 4. Incubator capable of heating to 70°C. 5. Pipettes and pipette tips. 6. Latex or nitrile disposable gloves. 7. ELIMINase®, DNA AWAY®, or a similar product.

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8. 96-well microplate. 9. Reagent reservoirs (100 ml). 10. Plate-based DNA extraction kit. 2.7. DNA Extraction: Plate-Based Extraction (96 samples): Non-kitBased Method (Adapted from Ref. 26)

1. 96-well microplate. 2. AcroPrep™ 96 1 ml filter plate with 1.0 μm Glass Fiber media (PALL Life Sciences). 3. Equipment for tissue grinding: FastPrep® or TissueLyser with tube adaptor. 4. Centrifuge with a deep-well swinging bucket rotor capable of achieving 5,600–6,000 × g force. 5. Vortex. 6. Orbital Shaker for microplates. 7. Laboratory tape. 8. Molecular biology grade water. 9. Ethanol: 100%. 10. Incubator capable of heating to 70°C. 11. Pipettes and pipette tips. 12. Latex or nitrile disposable gloves. 13. ELIMINase®, DNA AWAY®, or a similar product. 14. CTAB lysis buffer: 2% cetyltrimethylammonium bromide (CTAB), 100 mM Tris–HCl pH 8.0, 20 mM EDTA, and 1.4 M NaCl. 15. Binding buffer: 5 M guanidine thiocyanate, 20 mM EDTA pH 8.0, 10 mM Tris–HCl pH 6.4, and 4% Triton® X-100. 16. First wash buffer: 50% ethanol, 3 M GuSCN, 10 mM EDTA pH 8.0, 5 mM Tris–HCl pH 6.4, and 2% Triton® X-100. 17. Second wash buffer: 60% ethanol, 50 mM NaCl, 10 mM Tris– HCl pH 7.4, and 0.5 mM EDTA pH 8.0. 18. Square-well block PALL collar (PALL Life Sciences). 19. Square-well block.

2.8. PCR

1. D-(+)-Trehalose dehydrate: 10 and 20% solutions. 2. 10× Polymerase Chain Reaction (PCR) Buffer, without Mg (Invitrogen). 3. Magnesium chloride: 50 mM solution. 4. Molecular biology grade water. 5. Latex or nitrile disposable gloves. 6. Pipettes and pipette tips. 7. Deoxynucleotide solution mix: 10 mM. 8. Oligonucleotide primers (Table 1).

a

Trehalose buffer

10× Buffer

MgCl2

dNTPs

Forward primer

Reverse primer

Polymerase

Second

Third

Fourth

Fifth

Sixth

Seventh

Eighth

Total volume of reaction

DNA (30–50 ng/μl)

Recommended amount to mix for a 96-well plate

Last

Molecular-grade water

First

Total volume of PCR mix

Component

Order to add PCR components

0.05 mM 0.1 μM 0.1 μM 0.025 U/μl

10 μM 10 μM 5 U/μl

2.5 mM

50 mM 10 mM

1×

5%

Final concentration

10×

10%

Stock concentration

Table 1 General PCR mix for rbc L, ITS, ITS2, and trn H–psb A

12.5

2

10.5

0.0625

0.125

0.125

0.0625

0.625

1.25

6.25

2

Volume for 1 reaction (ml)

1,050

6.25

12.5

12.5

6.25

62.5

125

625

200

Volume for 100 reactionsa (ml)

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9. Platinum Taq DNA Polymerase (Invitrogen). 10. PCR 96-well microplate. 11. Aluminum Sealing Film (Axygene Scientific, VWR). 12. Clear Sealing Film (Axygene Scientific, VWR). 13. Thermocycler. 14. Microcentrifuge. 15. Centrifuge with a swinging bucket rotor for microplates. 16. PCR workstation. 2.9. PCR Product Determination: Precast E-gel Method

1. Precast agarose gel (e.g., 2% E-gel, Invitrogen). 2. E-Base. 3. Reagent reservoir. 4. Molecular biology grade water. 5. Latex or nitrile disposable gloves. 6. Pipette and pipette tips. 7. Gel imaging system.

2.10. PCR Product Determination: Routine Agarose Gels

1. Gel rig and combs. 2. Agarose. 3. Latex or nitrile disposable gloves. 4. Pipette and pipette tips. 5. Gel imaging system. 6. 1× TBE buffer: 90 mM Tris base, 90 mM boric acid, 2 mM EDTA. 7. DNA stain: Ethidium bromide or equivalent (e.g., SYBR® Safe DNA gel stain, Invitrogen). 8. Gel loading solution (e.g., Gel loading solution Sigma G7654) * if not already in the PCR mixture. 9. Size standard (e.g., 1 kb DNA ladder). 10. Power supply.

2.11. Cycle Sequencing

1. D-(+)-Trehalose dehydrate: 10% solution. 2. 5× Sequencing Buffer: 400 mM Tris–HCl pH 9.0, 10 mM MgCl2. 3. Molecular biology grade water. 4. Latex or nitrile disposable gloves. 5. Pipette and pipette tips. 6. 96-well PCR microplate. 7. Aluminum sealing foil.

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8. Clear sealing film. 9. Microcentrifuge. 10. Thermocycler. 11. Centrifuge with a swinging bucket rotor for microplates. 12. PCR workstation. 13. Oligonucleotide primer: 10 μM. 14. BigDye™ Terminator v3.1 Cycle Sequencing Kit (Applied Biosystems). 2.12. Cycle Sequencing Reaction Cleanup and Processing for an ABI 3730xl Capillary Sequencer

1. Sephadex® G50 (Sigma). 2. Acroprep™ 96 Filter plate, 0.45 μM GHP (PALL Corporation Catalog No. 5030). 3. Molecular biology grade water. 4. Latex or nitrile disposable gloves. 5. Pipette and pipette tips. 6. Septum (Applied Biosystems). 7. Black plate base (Applied Biosystems). 8. White plate retainer (Applied Biosystems). 9. Pop-7™ Polymer for 3730xl DNA Analyzers (Applied Biosystems). 10. 3730xl DNA Analyzer Capillary Array, 50 cm (Applied Biosystems). 11. 10× Running buffer for 3730xl DNA Analyzers (Applied Biosystems). 12. MicroAmp 96-well reaction plate (Applied Biosystems).

3. Methods 3.1. Field Collecting

1. Prior to going to the field, dispense the silica gel into scintillation vials (~2/3–3/4 full), plastic bags (~15 ml of silica), or a 1-L container (~15% full) for coin envelopes or tea bags. 2. Harvest the plant: whole plant if small, or a branch with leaves from woody shrubs or trees. 3. Place the voucher in the field press such that identifying features (flowers, fruits, both sides of leaves) can be easily inspected when dried. 4. Identify the voucher with a unique collecting number, either with a jewelry tag attached to the voucher or by writing on the paper the sample is pressed in.

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5. Take a small amount of leaf tissue (3–10 cm2; see Notes 1–6), and place in either: the scintillation vial containing silica gel, the plastic bag containing silica gel, or a coin envelope/tea bag, which is placed in the 1-L container with silica gel. 6. Label the container or coin envelope with the same collection number as the voucher. 3.2. Sample Storage

1. Store the tissue samples in a dry location or retain in silica until ready to subsample for DNA extraction (see Note 7).

3.3. Tissue Subsampling: For DNA Extraction Using Single Tubes

1. Clean the bench working area with ELIMINase®, DNA AWAY®, or a similar product. 2. With clean gloves and forceps, add one clean grinding bead to each tube and recap tubes. 3. Sterilize the forceps by dipping them in alcohol and flaming them. 4. Open a container with the sample, break off a piece of leaf or find a piece of the right size (see Notes 8–13), and insert it into a tube. 5. Label the tube with the collection number. 6. Clean the forceps by dipping them in alcohol and flaming, and then repeat step 4 for the remaining samples. 7. Change gloves often (or any time, you feel that they may have become contaminated).

3.4. Tissue Subsampling: For DNA Extraction Using 96-Well Plate Format

1. At a computer, organize the sample names in a spreadsheet in the plate format (8 rows × 12 columns). A good practice is to organize samples such that different genera are in adjacent wells. This facilitates the detection of cross contamination. 2. In the lab, clean the bench working area with ELIMINase®, DNA AWAY®, or a similar product. 3. With clean gloves and forceps, add one clean grinding bead to each tube in the plate, and add the strip caps to the tubes. 4. Organize the physical tissue samples in silica gel (vials, bags, or coin envelopes) on the bench, in columns and rows corresponding with the spreadsheet created in step 1. 5. Work with one strip of eight tubes (each corresponding to a numbered column) at a time. Remove one set of eight tubes to a new holder to physically separate the eight tubes being filled from the others. 6. Remove the lids from the strip of eight tubes and put them somewhere where they will not be contaminated by any flying plant material (e.g., between two kimwipes or on a kimwipe covered with a plastic lid).

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7. Sterilize the forceps by dipping them in alcohol and flaming them. 8. Open the container with the sample, break off a piece of leaf or find a piece of the right size (see Notes 8–13), and insert it into the correct tube. Pieces of plant tissue that are linear in shape (e.g., grass leaves and stems, conifer needles) need to be broken into smaller pieces to achieve proper homogenization using the grinding beads. 9. Clean the forceps by dipping them in alcohol and flaming, and wipe the gloves with a kimwipe moistened with ethanol in order to remove any plant tissue. 10. Repeat steps 8 and 9 for the remaining seven samples in the column. 11. Once the column of eight tubes is loaded, discard the gloves and put on new ones. 12. Attach the clean strip cap to the tubes, making sure that the lids are on tightly (they may pop off if not pushed all the way on). 13. Repeat the process from step 5, changing gloves after each set of eight tubes (or any time, you feel that they may have become contaminated). 3.5. Tissue Disruption

1. Homogenize the plant material with the grinding bead using a FastPrep®, TissueLyser, or a similar instrument: for the TissueLyser, apply 28 Hz for 30 s, then rotate the adaptors, and repeat once (or a maximum of two more times if necessary to obtain good disruption) (see Note 14). 2. Briefly centrifuge the tubes or the plate of strip tubes after homogenization to limit the amount of material stuck to the cap (see Note 15).

3.6. DNA Extraction: Kit-Based Protocols

1. For kit-based instructions.

methods,

follow

the

manufacturer’s

3.7. DNA Extraction: For Non-kit, Single Sample-Based Methods (Adapted from Ref. 26)

1. Carefully remove the screw caps from each tube and discard the caps. Powderized plant tissue will be adhered to the cap and will easily dislodge if the caps are not handled carefully (see Note 15). 2. Dispense 200 μl of CTAB lysis buffer to each tube and recap the tubes with new caps. 3. Gently invert each tube in order to mix the powderized plant material with the lysis buffer, and briefly centrifuge the tubes for 1,000 × g force for 1 min to collect the sample to the bottom. 4. Incubate the samples for 1 h at 65°C with occasional mixing by inversion.

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5. Centrifuge the tubes at 1,500 × g force for 1 min. 6. Remove the caps and transfer 50 μl of lysate from each sample to a new 1.5-ml microcentrifuge tube (see Note 16). 7. Add 100 μl of binding buffer to each tube with lysate. 8. Immediately after addition of the binding buffer, carefully and slowly mix three to four times by aspirating and dispensing 100 μl. 9. Transfer 150 μl of each lysate into a spin column, placed in a 1.5-ml microcentrifuge tube, and close the cap on the spin column. 10. Centrifuge at 5,000 × g force for 5 min to bind the DNA to the membrane of the spin column. 11. Add 200 μl of the first wash buffer to each spin column. 12. Centrifuge at 5,000 × g force for 2 min. 13. Remove the spin column from the tube, discard the flow through, and replace the spin column in the tube. 14. Add 500 μl of the second wash buffer to the spin column. 15. Centrifuge at 5,000 × g force for 5 min. 16. Remove the spin column from the tube and discard the tube and contents. 17. Open the cap of the spin column, place the spin column on the lid of a tip box, and incubate at 56°C for 30 min to evaporate residual ethanol. 18. Place the spin column in a new 1.5-ml microcentrifuge tube. 19. Add 50 μl of ddH2O (at 56°C) to the center of the spin column. 20. Incubate at room temperature for 1 min. 21. Centrifuge at 5,000 × g force for 5 min to collect the DNA eluate. 22. Remove the spin column and discard it. 23. Store the DNA at 4°C for short-term storage or at −20°C (preferably at −80°C) for long-term storage. 3.8. DNA Extraction: For Non-kit, PlateBased Methods (Adapted from Ref. 26)

1. Remove one set of strip tubes to a separate holder for cap removal and addition of CTAB lysis buffer. 2. Carefully remove the strip of caps using each individual cap tab to pull the cap off the tube, and discard the strip caps. Powderized plant tissue will be adhered to the cap and will easily dislodge if the caps are not handled carefully (see Note 15). 3. Dispense 200–350 μl of CTAB lysis buffer to each tube (depending on the amount of sample) and recap the tubes with a new strip cap.

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4. Repeat steps 2 and 3 for the remaining 11 sets of strip tubes. 5. Use tape to tightly seal the caps on the tubes (which may otherwise pop off during incubation). 6. Gently invert the rack of tubes once to mix the powderized plant material with the lysis buffer. 7. Briefly centrifuge the tubes at 1,000 × g force for 1 min to collect the sample to the bottom. 8. Incubate the samples for 1 h at 65°C using shaker (80–100 rpm). Do not invert rack. 9. Centrifuge the plate at 1,500 × g force for 1 min. 10. Remove the strip caps and transfer 50 μl of lysate from each sample to the corresponding position of a 96-well microplate (see Note 16). 11. Add 100 μl of binding buffer to each well. 12. Immediately after addition of the binding buffer, carefully and slowly mix three to four times by aspirating and dispensing 100 μl. 13. Transfer 150 μl of each lysate into a well in a 1 ml Acroprep™ 96-well glass fiber plate, placed on a 2-ml square-well block (see Note 17). 14. Seal the glass fiber plate with clear PCR film. 15. Centrifuge at 5,000 × g force for 5 min to bind the DNA to the glass fiber membrane. 16. Remove the PCR film and add 200 μl of the first wash buffer to each well of the glass fiber plate. 17. Seal the plate with clear PCR film and centrifuge at 5,000 × g force for 2 min. 18. Remove the PCR film and add 750 μl of the second wash buffer to each well of the glass fiber plate. 19. Seal the plate with clear PCR film and centrifuge at 5,000 × g force for 5 min. 20. Remove the seal, place the glass fiber plate on the lid of a tip box, and incubate at 56°C for 30 min to evaporate residual ethanol. 21. Position a collar on the collection microplate (optional) and place the glass fiber plate on top. 22. Add 50 μl of ddH2O (at 56°C) to each well of the glass fiber plate. 23. Seal the glass fiber plate with clear PCR film. 24. Incubate at room temperature for 1 min. 25. Place the assembled glass fiber plate and microplate on top of a square-well block to prevent cracking of the collection plate and centrifuge at 5,000 × g force for 5 min to collect the DNA eluate.

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26. Remove the glass fiber plate and retain it at −20°C as a backup until the extraction is determined to be successful, after which it can be discarded. 27. Cover the DNA plate with aluminum sealing film and store at 4°C for short-term storage or at −20°C (preferably at −80°C) for long-term storage. 3.9. PCR: PCR Mixture

1. Prepare and label a 1.5-ml microcentrifuge tube for the PCR cocktail of 100 reactions (Table 1). This number of reactions is recommended when using a 96-well plate to accommodate pipetting error. 2. Defrost all components of the cocktail at room temperature, except the polymerase which has to be kept at −20°C at all times prior to use. 3. Prepare the PCR cocktail adding the components in order listed in Tables 1–3 (see Notes 18–21). See also Table 4 for the standard primers for amplification of rbcL, matK, ITS, ITS2, and trnH–psbA. 4. Vortex the mix and centrifuge at 1,000 × g force briefly. 5. Dispense 10.5 μl of the PCR cocktail in each well using the same tip [replace tip occasionally (every 16 wells) to reduce pipetting error]. 6. Add 2 μl of the sample DNA (30–50 ng/μl) to each well. Leave one or two wells blank as a negative control. Use a fresh tip for each DNA sample. 7. Seal the plate tightly with aluminum foil (using a roller to seal) or thermo-seal cover (apply heat to seal) (see Note 22). 8. Centrifuge the plate at 1,000 × g force for 1 min (see Note 23). 9. Place the plate into the thermo-cycling block, close it, and apply the appropriate PCR program.

3.10. PCR Thermal Cycling Programs

1. rbcL, trnH–psbA (see Notes 24 and 25): 94°C for 4 min; 35 cycles of 94°C for 30 s, 55°C for 30 s, 72°C for 1 min; final extension 72°C for 10 min. 2. trnH–psbA for ferns and allies, and bryophytes (see Note 25): 94°C for 4 min; 2 cycles of 94°C for 45 s, 50°C for 45 s, 72°C for 1 min; 35 cycles of 94°C for 45 s, 45°C for 45 s, 72°C for 1 min; final extension 72°C for 10 min. 3. trnH–psbA using Phusion polymerase (see Note 26, Table 3): 98°C for 45 s; 35 cycles of 98°C for 10 s, 64°C for 30 s, 72°C for 40 s; final extension 72°C for 10 min. 4. matK first round (matK-KIM1R/matK-KIM3F) (see Note 27): 94°C for 1 min; 35 cycles of 94°C for 30 s, 52°C for 20 s, 72°C for 50 s; final extension 72°C for 5 min.

Polymerase

Eighth

0.5 μM 0.5 μM 0.1 U/μl

10 μM 10 μM 5 U/μl

0.15

0.375

0.375

0.15

7.5

Reverse primer

Seventh

0.2 mM

10 mM

0.225


Forward primer

Sixth

Recommended amount to mix for a 96-well plate

a

dNTPs

Fifth

1.5 mM

50 mM

0.75

1.875

1

MgCl2

Fourth

1×

5%

10×

20%

2.60


DNA (3–5 ng/μl)

10× Buffer

Third

Last

Trehalose buffer

Second

Final concentration

6.5


First

Stock concentration


Component


Table 2 PCR mix for mat K

650

15.0

37.5

37.5

15.0

22.5

75

187.5

260



Polymerase

Seventh

0.025 U/μl

2 U/μl

b

Recommended amount to mix for a 96-well plate Note that in limited trials HF buffer does not appear to be compatible with trehalose

a

0.1 μM

10 μM

0.125

0.1

0.1

10

Reverse primer

Sixth

0.1 μM

10 μM

0.056

2


Forward primer

Fifth

0.056 mM

10 mM

1×

1

dNTPs

Fourth

5×

6.32

0.3


DNA (30–50 ng/μl)

HF buffer (containing 1.5 mM MgCl2)b

Third

Last


Second

3%

Final concentration

9

DMSO

First

Stock concentration


Component


Table 3 PCR mix for use with Phusion polymerase

900

12.5

10

10

5.6

200

632

30


11 DNA Barcoding Methods for Land Plants 237

AB101 AB102

psbAF trnH2 psbA trnH(GUG) psbA501f

ITS

trnH–psbA

GTTATGCATGAACGTAATGCTC CGCGCATGGTGGATTCACAATCC CGAAGCTCCATCTACAAATGG ACTGCCTTGATCCACTTGGC TTTCTCAGACGGTATGCC

ACGAATTCATGGTCCGGTGAAGTGTTCG TAGAATTCCCCGGTTCGCTCGCCGTTAC

ATGCGATACTTGGTGTGAAT TCCTCCGCTTATTGATATGC

R F R F R

F R

F R

F R F R F R F R

R

R

F

Direction

Sang et al. (29) Tate and Simpson (28) Hamilton (30) Hamilton (30) Cox et al. (31)

Sun et al. (39) Sun et al. (39)

Chen et al. (20) White et al. (38)

Ki-Joong Kim, personal communication Ki-Joong Kim, personal communication Cuenoud et al. (37) Cuenoud et al. (37) Damon Little, personal communication Damon Little, personal communication Fazekas et al. (5) Fazekas et al. (5)

Fazekas et al. (5)

Levin et al. (35), modified from Soltis et al. (34) Kress and Erickson (24), modified from Fofana et al. (36)

References

See Notes 24–27 for primer usage and alternatives. As different authors use different conventions as to what constitutes “forward” and what constitutes “reverse” primers, the notation of F and R on primer names can mean different things. This is particularly problematic for matK and trnH–psbA. The “Direction” column indicates primer orientation with reference to the direction of the reading frame of rbcL and matK and following the convention of clockwise orientation for trnH–psbA

ITS-S2F ITS4

ITS2

rbcLajf634R ACCCAGTCCATCTGGAAATCTTGGTTC CGTACAGTACTTTTGTGTTTACGAG CGATCTATTCATTCAATATTTC TCTAGCACACGAAAGTCGAAGT CTGGATYCAAGATGCTCCTT GGTCTTTGAGAAGAACGGAGA CCCTATTCTATTCAYCCNGA CGTATCGTGCTTTTRTGYTT

GAAACGGTCTCTCCAACGCAT

rbcLa-R

matK-KIM1R matK-KIM3F matK-390f matK-1326r NY552F NY1150R matKpkF4 matKpkR1

GTAAAATCAAGTCCACCRCG

rbcLa-F

rbcL

matK

ATGTCACCACAAACAGAGACTAAAGC

Primer name

Region

Sequence (5′–3′)

Table 4 Primers commonly used for DNA barcoding in plants


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5. matK second-round failure tracking (matK-390f/matK-1326r) (see Note 27): 94°C for 1 min; 35 cycles of 94°C for 30 s, 50°C for 40 s, 72°C for 40 s; final extension 72°C for 5 min. 6. ITS (AB101/AB102) (see Note 28): 94°C for 5 min; 30 cycles of 94°C for 1 min, 55°C for 1 min, 72°C for 1 min, 45 s; final extension 72°C for 10 min. 7. ITS2 (ITS-S2F/ITS4): 94°C for 5 min; 35 cycles of 94°C for 30 s, 56°C for 30 s, 72°C for 45 s; final extension 72°C for 10 min. See Note 29 in situations, where results from PCR are unsuccessful or poor. 3.11. PCR Product Determination: Electrophoresis with Precast E-gels

1. Open the package with precast agarose gel (see Note 30), remove the plastic comb, and place the gel on the mother E-base. 2. Set the mother E-base at “EG” program and a runtime of 4 min. 3. Load 14 μl of molecular-grade water into each well of the 96-well precast agarose gel. 4. Load 3–4 μl of each PCR product into the corresponding E-gel well. 5. Slide E-gel into electrode connections of mother E-base and start electrophoresis. A green light indicates the beginning of run. A red light and beeping indicate the end of run. Stop the current by pressing pwr/prg button. 6. Remove E-gel from base and capture a digital image with the imaging documentation system.

3.12. PCR Product Determination: Electrophoresis with Routine Agarose Gels

There is a large selection of gel combs and trays on the market designed to accommodate different numbers of samples. Please refer to the manufacturer’s notes for the recommended volume of agarose to be used. 1. Select the appropriate gel tray and combs for the number of samples to be run (leaving an appropriate number of wells free for size standards). Seal the ends of the tray with masking tape or use a gel-forming cassette. 2. Weigh out the agarose and place in a glass conical flask. To check PCR success, a 1% agarose gel is used; 1% agarose gel = 1 g of agarose per 100 ml of 1× TBE buffer. 3. Add the appropriate volume of 1× TBE buffer to the agarose and gently swirl. 4. Heat the solution in a microwave on maximum heat setting for approx. 30 s, remove flask from the microwave, and gently swirl to mix. Continue to heat, mixing occasionally. Carefully

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remove the solution from the microwave, gently swirl, and check that all the agarose has dissolved. 5. Place the gel on the bench and leave to cool (or cool under cold running water), until it is comfortable to touch the side of the conical flask. 6. Add the appropriate volume of DNA gel stain (see Note 30). For Sybrsafe, this is 1 μl/10 ml of agarose gel; for ethidium bromide, this should be to a final concentration of 0.5 μg/ml of agarose gel. Gently swirl the solution to mix. 7. Pour the gel into the gel tray and leave to set for approx. 30 min. 8. If gel-loading solution is not already in the PCR mixture, prepare your samples for gel electrophoresis by mixing the gelloading solution with the PCR product (3 μl gel-loading solution plus 5 μl PCR product). 9. Carefully remove the masking tape or undo the clamp of the gel-forming cassette and gently remove the comb. 10. Place the agarose gel in the electrophoresis tank containing 1× TBE buffer, making sure that the gel is totally immersed in buffer. The buffer should just be covering the surface of the gel. 11. Load the recommended volume of size standard into the assigned lanes (typically, 0.1 μg of standard per millimeter lane width). Then, load the samples into the subsequent wells. 12. Run gel for 30 min to 1 h at 80 V. 13. Transfer the gel to the imaging documentation system and capture a digital image. 3.13. Cycle Sequencing

1. Dilute the PCR product: (a) For rbcL, ITS, ITS2, trnH–psbA: one part of PCR product/two parts of water. (b) For matK: one part of PCR product/nine parts of water. 2. Cover the plate with plastic seal, and spin at 1,000 × g force for 1 min. 3. Defrost sequencing reagents (Table 5) at room temperature. Keep BigDye™ away from light exposure prior to use. 4. Prepare sequencing mix adding components in the order listed in Table 5 (see Note 31). After adding BigDye™, mix components gently by inverting the tube several times. Do not vortex. Add one primer. Mix gently with tip. Note that separate reactions are carried out using the forward or reverse primers. 5. Dispense 9.0 μl of sequencing mix into each well of 96-well plate.

Primer

Fifth

Last

BigDye™

Fourth

11


9.0

1

0.25

1.875

0.875

5


2

10 μM

5×

Final concentration

Diluted PCR product

Total volume of sequencing mix

Sequencing buffer

Third

10%

Stock concentration

b

Recommended amount to mix for a 96-well plate Sequencing buffer: for 50 ml: 20 ml of 1 M Tris–HCl pH 9, 500 μl of 1 M MgCl2, 29.5 ml of molecular-grade water

a


Second b

Trehalose (Sigma-Aldrich, No. T9531-100 g)

Component

First

Order to add components

Table 5 General cycle-sequencing mix

936

104

26

195

91

520


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6. Add 2 μl of diluted PCR product to each well (use fresh tip for each PCR product). 7. Place aluminum foil or heat-seal cover over the top of the 96-well plate. Apply heat for heat-seal cover, and use roller to close the plate tightly (see Note 22). 8. Spin the plate using centrifuge at 1,000 × g force for 1 min (see Note 23). 9. Place the plate into the thermocycler block and apply the program (see Note 32): 96°C for 2 min; 30 cycles of 96°C for 30 s, 55°C for 15 s, 60°C for 4 min; hold at 4°C. 10. After cycle sequencing reaction is complete, keep the plate in a dark box at 4°C to avoid degradation of BigDye™. 3.14. Cycle Sequencing Cleanup and Processing for an ABI 3730xl Capillary Sequencer

1. Measure dry Sephadex G-50 (Sigma-Aldrich, Cat. No. G5080500 g) with the MultiScreen Column Loader (Millipore, Cat. No. MACL09645) into the Acroprep 96 Filter plate with 0.45 μm GHP membrane (PALL, Cat. No. PN5030). This loader adds the specific amount of Sephadex required (see Note 33). 2. Hydrate each well with 300 μl of molecular-grade water using a pipette. 3. Let the Sephadex hydrate overnight at 4°C or for 3–4 h at room temperature before use. 4. Assemble the Sephadex plate onto the collection plate and secure with two rubber bands. 5. Centrifuge at 750 × g force for 3 min to drain the water from wells. Discard water from the collection plate (when centrifuging two plates, make sure that both sets have equal weight which can be achieved by using additional rubber bands). The collection plate can be reused without autoclaving. 6. Add the entire volume of the sequencing reaction to the centre of the Sephadex columns using a pipette. 7. Add 25 μl of 0.1 mM EDTA to each well of the Sephadex plate. 8. Elute clean sequencing reaction by attaching a 96-well plate to the bottom of Sephadex plate and secure with rubber bands. 9. To balance two plates, attach additional rubber bands as needed. 10. Centrifuge at 750 × g force for 3 min. Remove Sephadex plate. 11. Cover the top of the collection plate with a septum. 12. Place 96-well plate into black plate bases and attach white plate retainer.

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13. Stack assembled plate in ABI 3730xl capillary sequencer and import plate record using Plate manager module of the Data Collection software (Applied Biosystems). 14. Begin sequencing run with Run Scheduler. 3.15. Sequence Editing

Careful and consistent editing of the raw sequence data is a critical component of generating a high-quality dataset. There are a number of software programs (e.g., Sequencher, CodonCode Aligner, etc.) that allow the import of raw trace files and include a variety of editing features. Since each sequence editing program is different, we cannot include a software-specific detailed editing procedure. We present instead the chain of events involved in going from the output of the sequencer to a useable sequence. 1. Retrieve electropherogram trace files from sequencer. 2. Import trace files into a sequence editing software package. 3. Generate sequence-quality scores for individual trace files. 4. Trim primer sequences from the sequences. 5. Trim sequences from both ends based upon minimum quality threshold (e.g., mean QV > 20 and no more than 2 bp QV < 20 in any 20-bp window). 6. Assemble forward and reverse sequence traces for each individual sample to create a sequence contig. 7. Manually edit individual sequences: pay particular attention to bases with low-quality scores or ambiguous calls (see Notes 34–37). 8. Acquire sequence-quality statistics for individual forward and reverse sequences (e.g., length of read, proportion of bases with QV > 20). 9. Generate consensus sequence. 10. Acquire consensus sequence quality statistics (e.g., length of consensus, percentage of bidirectional coverage, proportion of bases with QV > 20 for unidirectional and bidirectional portions of the consensus). 11. Export consensus sequence for downstream analysis.

4. Notes 1. Properly collected plant tissue is essential for maximizing PCR and sequencing success. Key to this process is that material from which DNA is extracted must be dried as quickly as possible to prevent the degradation of the DNA. Field collections

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of specimens must be immediately split into two components: (a) the voucher and (b) a portion of the voucher (typically, leaf tissue) which is placed in a container with silica gel or similar drying agent. It is important that the portion taken for DNA is put into silica gel as rapidly as possible after harvesting from the field. This should ideally be done immediately, but if impractical, it should be done no later than at the end of the collecting day. Delays to drying material in silica gel can result in samples with reduced DNA quality and lower PCR success. 2. It is important to keep the freshly collected tissue samples in separate containers. Pooling different samples into a single Ziploc bag, for example, increases the chances of cross contamination. 3. We describe three types of containers that we have used in various settings, each with relative advantages and disadvantages. (a) Scintillation vials provide a separate enclosed environment for each sample. This can be useful in humid conditions, in which coin envelopes may absorb some moisture from the air, slowing the tissue drying process, or for tissue that has a high water content and dries more slowly inside a coin envelope rather than when in direct contact with the silica. (b) Coin envelopes are probably the simplest medium for sampling plant tissue. It is easier to insert a sample into an envelope than into the narrow opening of a scintillation vial. Multiple coin envelopes can be stored in an airtight container with silica gel, requiring less space than scintillation vials. The envelopes also keep the silica gel separate from the tissue, facilitating tissue subsampling. Tea bags can also be used in place of envelopes; they are more porous, facilitating the drying process, but are also slightly more fragile. (c) Small (~10 × 15 cm) plastic bags with silica can work well in the field, but are prone to punctures from thorns or prickles, and are somewhat permeable which does exhaust the silica over time. When the samples are dry, the plastic bags need to be handled carefully to prevent excessive breakage of the plant tissue. 4. In the case of specimens that are likely to take a long time to dry (such as samples with waxy leaves), tear the leaf sample into smaller fragments or chop with a sterile blade to increase the surface area available for contact with the silica gel. 5. The best samples for plant DNA extraction typically come from actively growing plant tissues; senescing, damaged, or infected tissues should be avoided. The usual choice of plant tissue is leaves, but shoot tips or flower buds or petals can also be used. For canopy tree species in which reaching leaf or flower material

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is logistically challenging, an alternative approach is to use a leather punch to obtain samples of cambium tissue which avoids the need for tree climbing (27). 6. Sampling herbarium material for DNA extraction can be successful, but success is often variable and unpredictable. The quality of the extraction is most likely a function of the age of the specimen, the species in question, and the speed with which samples have been dried, which is often unknowable. The priority should be given to samples not much greater than 10 years old. However, the most critical criterion is that the samples should still be green in color. Brown coloration of the herbarium sample indicates that the tissue quickly oxidized after collection or was infected by mold, indicating that the DNA is most likely degraded and/or contaminated by fungal DNA. 7. Tissue samples from which DNA extractions are made should be prevented from rehydrating from the atmosphere. This can be achieved through a climate-controlled facility or in airtight containers (refresh the silica as necessary). Long-term experiments are still needed to provide empirical data on optimum storage procedures for tissue samples. 8. The sampling of silica-dried material into tubes for DNA extraction and the extraction process are probably the most important steps in the process of generating good-quality DNA barcode data. It is the step that is the easiest for contamination or sample mix-up to occur. Thus, it is very important to follow the steps outlined in Subheading 3.3 to prevent this. A poor-quality extraction will result in inefficient or failed PCR reactions. 9. The appropriate amount of plant material to sample for DNA extraction is 10–15 mg dry tissue. In this case, more is not better; using more than this amount of tissue will result in a poorly ground sample, overwhelm the buffers used in the extraction process, and result in low-yield or poor-quality DNA. This amount usually corresponds to ~0.5 cm2, but may be smaller depending on the leaf thickness. Plastic materials (such as sampling tubes) often have a static charge that will attract small particles of plant tissue. Fragments of plant material literally jump from one well to another, so care must be exercised when placing bits of leaves into the tubes. 10. Plant tissues that are linear in shape (e.g., grass leaves and stems, conifer needles) need to be broken into smaller pieces to achieve proper pulverization using the grinding beads. 11. When sampling plant tissue from herbarium samples in areas where an alcohol burner is prohibited, it is good practice to wipe the forceps after each sample with a kimwipe moistened with ethanol.

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12. It is important to keep freshly collected silica-dried material and older herbarium-sampled tissue in separate extraction plates, as they may require different extraction protocols. 13. A note on bryophytes: Extreme care is required when sampling bryophytes due to the common occurrence of mixed species samples being collected from the field. Tissue subsampling is best done at the same time as determinations are made. 14. A frequency higher than 28 Hz can destroy the tubes. We do not recommend homogenization for longer than a total of 1 min, with the exception of samples with very tough tissue in which case an additional run of 30 s can be applied. 15. After the plant tissue is ground to a fine powder, the tubes require careful handling. Centrifuging does not help significantly in removing powderized plant tissue from the lids or caps as the static charge is strong enough to keep them adhered to the interior surface of the tube’s walls and caps. Opening the caps should be done with extreme care to avoid cross contamination prior to addition of the lysis buffer. 16. In the non-kit-based protocols are provided, the entire volume of the CTAB lysate is not used. Unused lysate can be stored at −20°C as a backup until the extraction is determined as being successful as indicated by the results of first PCR reaction. The lysate can also be used as a source for additional extractions if more testing of the DNA is necessary. 17. The square-well blocks that are specified in the protocol have enough volume to collect all the wash buffers without needing to discard between washes. However, if a block with a smaller volume is used, it may be necessary to discard the wash buffer between steps 16 and 17 of Subheading 3.8. 18. Trehalose (which is also a potent PCR enhancer) acts as a cryoprotectant for Taq polymerase when PCR mixes are prepared in large volume batches and frozen for future use. 19. Many available PCR protocols for matK include 4% DMSO. Experiments based on several hundred reactions have demonstrated that a 5% Trehalose solution can replace DMSO without any significant difference in PCR success or sequence quality. 20. After DNA extraction, it is recommended to begin the first round of PCR for the rbcL DNA barcoding marker using the nearly universal primers rbcLa-F/rbcLa-R; a greater degree of PCR success and quality is obtained in bryophytes with the reverse primer rbcLajf634R. These primers generate a high rate of PCR success with DNA of good quality. Hence, this first PCR for rbcL acts as a test for DNA quality for a broad variety of taxa among angiosperms, gymnosperms, ferns, and mosses.

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21. PCR cleanup is both expensive and time consuming, but can be avoided through use of the low concentrations of primers and dNTPs in the PCR mix and the subsequent dilution of the PCR product prior to cycle sequencing reaction. This protocol provides a high success rate for PCR and sequences for regions that are amplified by universal, highly conserved primers (plastid rbcL, trnH–psbA, and nuclear ribosomal ITS2). In contrast, the matK DNA barcoding region needs distinct conditions for successful PCR amplification. For matK, the concentration of the primers, dNTPs, and Taq polymerase cannot be significantly reduced. Based on experiments optimizing the PCR conditions for matK, we recommend a protocol with diluted DNA (0.3–0.5 ng/μl) and a smaller PCR reaction volume (7.5 μl). These conditions have yielded a higher rate of PCR success and increased sequence quality over the general PCR mix. 22. The volumes of the PCR and cycle sequencing reactions recommended here are very small. Thus, it is very important to follow the instructions in Subheadings 3.9 and 3.14 carefully. The foil or thermal-seal cover should be placed evenly and tightly over the PCR plate without wrinkles or holes to prevent evaporation during PCR cycling. 23. Centrifuging is required to collect the PCR components at the bottom of the well and eliminate any air bubbles that might have been trapped. It also aids in mixing the PCR components with the DNA sample, or cycle sequencing mix with PCR product. 24. Although rbcL is present in the vast majority of land plants, there are some groups, such as holoparasites, that no longer have a functioning copy of this gene. As a result, the primers most commonly used typically do not work in these groups. 25. The primers most widely used for PCR amplification of the plastid trnH–psbA intergenic spacer for DNA barcoding are those recommended by Kress et al. (7) or Kress and Erickson (10) (Table 4). They are, respectively, trnH2 (originally from ref. 28) and psbAF (originally from ref. 29) or trnH(GUG) and psbA (originally from ref. 30). In bryophytes, this region is often short (