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
223
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A.J. Fazekas et al.
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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225
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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A.J. Fazekas et al.
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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A.J. Fazekas et al.
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DNA Barcoding Methods for Land Plants
229
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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233
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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A.J. Fazekas et al.
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
Total volume of reaction
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
Volume for 1
reaction (ml)
DNA (3–5 ng/μl)
10× Buffer
Third
Last
Trehalose buffer
Second
Final concentration
6.5
Molecular-grade water
First
Stock concentration
Total volume of PCR mix
Component
Order to add PCR
components
Table 2
PCR mix for mat K
650
15.0
37.5
37.5
15.0
22.5
75
187.5
260
Volume for 100
reactionsa (ml)
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A.J. Fazekas et al.
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
Total volume of reaction
Forward primer
Fifth
0.056 mM
10 mM
1×
1
dNTPs
Fourth
5×
6.32
0.3
Volume for 1
reaction (ml)
DNA (30–50 ng/μl)
HF buffer (containing
1.5 mM MgCl2)b
Third
Last
Molecular-grade water
Second
3%
Final concentration
9
DMSO
First
Stock concentration
Total volume of PCR mix
Component
Order to add PCR
components
Table 3
PCR mix for use with Phusion polymerase
900
12.5
10
10
5.6
200
632
30
Volume for 100
reactionsa (ml)
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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
238
A.J. Fazekas et al.
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DNA Barcoding Methods for Land Plants
239
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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A.J. Fazekas et al.
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
Total volume of reaction
9.0
1
0.25
1.875
0.875
5
Volume for 1
reaction (ml)
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
Molecular-grade water
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
Volume for 104
reactionsa (ml)
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A.J. Fazekas et al.
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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DNA Barcoding Methods for Land Plants
243
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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A.J. Fazekas et al.
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
11
DNA Barcoding Methods for Land Plants
245
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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A.J. Fazekas et al.
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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DNA Barcoding Methods for Land Plants
247
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 (<200 bp) and an alternative primer, psbA501f
(31) located further back in the psbA gene can be used to
obtain additional characters in combination with the primer
trnH2.
26. The trnH–psbA region often contains homopolymer runs that
are known to reduce sequence quality after the run. Fazekas
et al. (32) have shown that the use of alternative polymerases,
such as Phusion (Finnzymes) or Herculase II Fusion (Agilent),
can improve quality for runs of up to 13–14 bases.
248
A.J. Fazekas et al.
27. The matK gene region is more difficult to amplify and sequence
than rbcL for a number of reasons. First, it is approximately
300 bp longer than rbcL, and thus more sensitive to DNA
degradation. Second, the presence of mononucleotide repeats
in some groups dramatically affects the quality of the sequence
reaction, resulting in a contig that is primarily supported by
two unidirectional sequences with only a small amount of overlap. Finally, matK requires different primer combinations for
different taxonomic groups. Therefore, it is important to estimate the taxonomic composition of the plate prior to amplification. If the plate contains angiosperms from different genera,
families, and even orders, the combination of matK-KIM1R/
matK-KIM3F is the optimal first choice. This primer combination was recommended as the first choice by the CBOL Plant
Working Group (13), and has been confirmed in thousands of
PCRs from floristic projects in biodiversity hot spots. Those
samples, which failed in the first round, are collected into a
new plate, and subjected to a second round of PCR using the
primers matK390f/matK1326r (failure tracking). Usually, the
combination of these two sets of matK primers yields around
80% successful sequences in floristic projects focusing on angiosperms. However, if the project is represented by one specific
group like a genus or family that does not work well with any
of these primer combinations, it is best to search for appropriate primers for this group. A selection of order-specific primers
has been published by Dunning and Savolainen (33). Two
alternate primer pairs for gymnosperms (NY552F/NY1150R
and matKpkF4/matKpkR1) are given in Table 4. Routine
amplification of this region is difficult in non-seed plants and
the development of primers for ferns/fern allies and bryophytes is currently underway.
28. ITS and ITS2 offer higher levels of species discrimination in
some groups. However, one risk with ITS is that of fungal
contamination. Even the cleanest leaf sample will likely have
fungal hyphae associated with it, and in some groups this can
be a serious source of contamination. For the entire ITS region,
the use of the angiosperm-specific primers AB101 and AB102
can reduce this problem for flowering plants.
29. In situations where PCR is unsuccessful or patchy in its success,
some optimization of PCR conditions can improve success. An
often-successful first approach is to dilute the DNA ten times.
DNA is often not limiting in PCR, but the extraction process
occasionally does not remove all PCR inhibitors sufficiently.
A dilution often reduces inhibitors to the point, where PCR
can succeed. Other steps to improve success are as follows: (a)
for faint or absent PCR products: a decrease in primer-annealing
temperatures, an increase in primer concentration, an increase
11
DNA Barcoding Methods for Land Plants
249
in MgCl2, or an increase in the number of cycles and (b) when
multibanded PCR products are produced: an increase in
primer-annealing temperatures, a decrease in primer concentration, or a decrease in MgCl2. These various approaches of
course depend on the initial PCR conditions one starts with;
there are minimums and maximums, which, if exceeded, will
usually result in failed PCR.
30. Ethidium bromide is highly toxic and a mutagen. Handling
agarose gels and/or buffers containing ethidium bromide
requires a designated work area isolated from other space in
the lab, dedicated pipettes, and water reservoirs. Nitrile gloves
should be worn when handling any components, and discarded
when the PCR check is completed to avoid contamination by
ethidium bromide. There are numerous alternatives to ethidium bromide that are less toxic. SYBR® Safe DNA gel stain, for
example, is not classified as hazardous waste under the US federal regulations but has a comparable sensitivity to ethidium
bromide.
31. The use of 5.5% trehalose in the sequencing reaction mix allows
sequencing mixes to be premade in primer specific batches,
aliquoted directly into plates, covered with PCR film, and frozen for future use. The mix with the primer can be stored at
−20°C for up to 3 months. However, one must avoid unnecessary thawing of the mix before use, which can cause degradation of the Dye Terminator. The mix should be thawed only
just prior to use.
32. The annealing temperature of the sequencing reaction can also
be adjusted to the specific primer conditions. For example, for
those primers which require a 50°C annealing temperature
during PCR, the annealing temperature in the cycle sequencing program also can be set at 50°C. However, it is not recommended to use an annealing temperature for the sequencing
reaction lower than 50°C.
33. For higher throughput, the semiautomated AutoDTR™
method from EdgeBio® (http://www.edgebio.com/catalog/
dye-terminator-removalproducts-AutoDTR™-96-c-28_1005.
html) has been used at the Canadian Centre for DNA
Barcoding. This sequencing cleanup method is less sensitive to
PCR product concentration and allows longer high-quality
reads and a further reduction of BigDye™ in the sequencing
reaction due to increased sensitivity.
34. Check and maintain the proper orientation of the sequences.
Sequences that are in the reverse complement orientation disrupt proper alignment, causing subsequent analytical problems
if not corrected.
35. Look for odd gaps or insertions in the sequence, especially at
the ends of the read. The trace file you view is a result of an
250
A.J. Fazekas et al.
algorithm that interprets the fluorescence pattern and compression artifacts often result at the ends of a read. These are
particularly problematic with multiple repeats, where the algorithm may have difficulty distinguishing whether there are 4 or
5 Adenine nucleotides in a row for example.
36. Check for an open reading frame (ORF) in coding regions,
such as rbcL and matK. This is an important first step in identifying stop codons and errors in editing. In the correct reading frame, there should be no internal stop codons present in
coding regions. There are two potential reasons for observing
stop codons in the sequence. First, it may be a real stop codon
indicating that the region that has been sequenced is no longer
functional, and likely a pseudogene. These cases are quite rare
in both rbcL and matK genes, and usually observed in only a
few specific groups of plants. Second, it can be evidence of
errors due to editing, e.g., miscalls, or a frameshift of the ORF
due to missed or extra base calls. Each instance needs to be
carefully investigated and addressed.
37. Check for chimeric sequences (a result of nonspecific PCR
amplification from multiple templates), bacterial, virus, fungal,
and algal contaminations, and remove them from the consensus sequence.
Acknowledgments
We are grateful to Michelle Hollingsworth, Alan Forrest, David
Erickson, and John Kress for helpful comments on this
manuscript.
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