Curated Experiment: DNA Barcoding

Identifying Taxa through DNA barocode sequences

Summary: Identifying species by sequencing "DNA Barcodes" and compare sequences to a reference library.

Protocol information

Protocol Credits

• Author(s)
  • Anna Feitzinger, Cold Spring Harbor Laboratory
  • Jeffry Petracca, Cold Spring Harbor Laboratory
• Maintainer/contact: Anna Feitzinger, Cold Spring Harbor Laboratory: email
• Last updated: March, 2025
• Source materials and references
  • Barcoding 101 Lab Resources
  • Rapid barcoding protocol

Nanopore Sequencing

• Sequencing format: Flongle
• Sequencing kit: Rapid Barcoding Kit 24 v14 (SQK-RBK114.24)
• Oxford Nanopore Sequencing protocol: Rapid Barcoding Kit (SQK-RBK114.24)
• Indexed/Barcoded: Yes, 24 indicies
• Samples per run: 24 samples

Computer and Bioinformatics

• Analysis tools
  • EPI2ME
    • EPI2ME amplicon workflow
• Software to download or install:
  • Oxford Nanopore Software Downloads Oxford Nanopore account and login required:
    • MinKNOW
    • EPI2ME Desktop Application
  • Other software:
    • Docker
• Analysis difficulty: Easier
• Command line needed: No
• GPU/Super-high accuracy basecalling required: No

Reagents

Sample collection and prep

• See resources and recipies at barcoding101 Sample Collections

DNA extraction

For rapid DNA extraction only:

• Lysis solution (6 M Guanidine Hydrochloride GuHCl) (120 µL)
• Wash buffer (480 µL)
• TE buffer (75 µL)
• Specimen tissue sample(s)
• 2 Sterile plastic pestles
• Whatman No.1 Chromatography paper discs (2, 3-mm diameter)

For chelex DNA extraction only:

• 10% Chelex Solution (2 tubes of 100 µL)
• 2 Sterile plastic pestles
• 6 Sterile toothpicks or pipette tips
• Tissue sample(s) (from Part I)
• 2 Microcentrifuge tubes (1.5mL)

PCR

• For Ready-To-Go PCR Bead Amplification:
  • 2 Ready-To-Go PCR Beads in 0.2- or 0.5-mL PCR tubes
  • Appropriate primer/loading dye mix (50 µL; 23 µL per reaction)*

Barcoding Primers

Plant Primers:

• rbcLa Forward 5’- TGTAAAACGACGGCCAGTATGTCACCACAAACAGAGACTAAAGC-3’
• rbcLa reverse 5’- CAGGAAACAGCTATGACGTAAAATCAAGTCCACCRCG-3’

Invertebrate Primers:

• LCO1490 5'-TGTAAAACGACGGCCAGTGGTCAACAAATCATAAAGATATTGG-3'
• HC02198 5'-CAGGAAACAGCTATGACTAAACTTCAGGGTGACCAAAAAATCA-3'

Fungi Primers:

• ITS1F 5'-TGTAAAACGACGGCCAGTTCCGTAGGTGAACCTGCGG-3'
• ITS4 5'-CAGGAAACAGCTATGACTCCTCCGCTTATTGATATGC-3'

DNA prep, library creation, and sequencing

• Rapid Barcoding Kit (SQK-RBK114.24)
• Flongle Expansion Kit (EXP-FLP002)
• Flongle Flow Cell (FLG-FLO114)

Equipment and consumables

Lab equipment

• Micropipettes and tips (2-1000 μL)
• Magnetic microfuge tube racks
• Microfuge tube rack
• Microcentrifuge
• Thermal cycler
• Qubit™ tubes

Consumables

• 1.5 mL microfuge tubes
• PCR tubes
• Qubit™ fluorometer, Nanodrop™, QIAxpert®, or other method of DNA quantification

Nanopore sequencing equipment

• MinION Flongle Adapter
• MinION Mk1B or Mk1D sequencer

Computer equipment

• Desktop or laptop with MinKNOW software

Estimated timings

• Sample Collection: Variable
• DNA extraction: 60-75 minutes
• PCR, DNA quality control, and library prep:
  • PCR: ~2 hours
  • Quality control: 10 min (Qubit); 60-75 min (electrophoresis)
  • Library prep: 30-60 min
• Sequencing: Atleast 60 min
• Data analysis: 30-60 min

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Background

Identifying organisms has grown in importance as we monitor the biological effects of global climate change and attempt to preserve species diversity in the face of accelerating habitat destruction. We know very little about the diversity of plants and animals—let alone microbes—living in many unique ecosystems on earth. Less than two million of the estimated 5-50 million plant and animal species have been identified. Scientists agree that the yearly rate of extinction has increased from about one species per million to 100-1,000 per million. This means that thousands of plants and animals are lost each year. Most of these have not yet been identified.

Classical taxonomy falls short in this race to catalog biological diversity before it disappears. Specimens must be carefully collected and handled to preserve their distinguishing features. Differentiating subtle anatomical differences between closely related species requires the subjective judgment of a highly trained specialist – and few are being produced in colleges today.

DNA barcodes allow non-experts to objectively identify species – even from small, damaged, or industrially processed material. Just as the unique pattern of bars in a universal product code (UPC) identifies each consumer product, a “DNA barcode” is a unique pattern of DNA sequence that can potentially identify each living thing. Short DNA barcodes, about 700 nucleotides in length, can be quickly processed from thousands of specimens and unambiguously analyzed by computer programs.

DNA amplicons are the copied sections of DNA produced by an amplification process that replicates a particμLar target region of a DNA molecule. For example, DNA amplicons are commonly generated via the polymerase chain reaction (PCR).

In DNA barcoding, a DNA amplicon might represent a region of DNA that can be used to determine the species of an organism. In medical research, amplicon DNA sequences might be used to detect pathogens, search for inherited diseases, or identify single nucleotide polymorphisms (SNPs) that are associated with a particular disease phenotype. More broadly, amplicon sequencing can be used to assess and analyze genetic variation at particular positions in a genome.

Explanation

Because the regions of DNA used for identification are approximately ~70% conversved, differences can be used to distinguish between species but allows the same primer pair to be use across many species.

• rbcL or matK located on the chloroplast are used to identify plants
• CO1 located on the mitochondria is used for animals, including invertebrates.
• ITS located in nuclear ribosomal DNA (rDNA) is used for fungi

DNA sequencing can be carried out in a number of different ways depending on the needs of an experiment. The ONT MinION sequencing device offers a portable, versatile, efficient, and relatively inexpensive way to sequence DNA in a classroom setting. In order to sequence DNA amplicons on this platform, they must first be prepared using an ONT kit. One efficient option for amplicons is the Rapid Barcoding Kit, which we used in this protocol.

Example Projects

DNA Barcoding Student Project Examples

Example of DNA Barcoding student projects can be found for the DNALC's barcoding programs below:

• Urban Barcode Research Project
• Barcode Long Island
• Urban Barcode Research Project

Additional Reading

1. Hebert P.D., Cywinska A., Ball S.L., deWaard J.R. (2003). Biological identifications through DNA barcodes. Proceedings of the Royal Society B: Biological Sciences 270(1512): 313-21.
2. Hebert P.D.N., Penton E.H., Burns J.M., Janzen D.H., Hallwachs W. (2004). Ten species in one: DNA barcoding reveals cryptic species in the neotropical skipper butterfly Astraptes fulgerator. Proc Natl Acad Sci USA. 101(41):14812-7.
3. Hollingsworth P.M. et al (2009). A DNA barcode for land plants. Proc Natl Acad Sci USA 106(31): 12794-7.
4. Ratnasingham, S., Hebert, P.D.N (2007). BOLD: The Barcode of Life Data System. Molecular Ecology Notes 7(3): 355-64.
5. Stoeckle M. (2003). Taxonomy, DNA, and the Bar Code of Life. BioScience 53(9): 2-3.
6. Van Den Berg C., Higgins W.E., Dressler R.L., Whitten W.M., Soto-Arenas M.A., Chase M.W. (2009) A phylogenetic study of Laeliinae (Orchidaceae) based on combined nuclear and plastid DNA sequences. Annals of Botany 104(3): 417-30.
7. Benson D.A., Cavanaugh M., Clark K., Karsch-Mizrachi I, Lipman D.J., Ostell J., Sayers E.W. (2013). Nucleic Acids Res. GenBank. 41(D1): D36–D42.

Sample Collection

Goal: Collect samples for DNA Barcoding

Instruction tip

Each student (up to 24) can prepare their own sample.

Summary

Your collection of specimens may support a census of life in a specific area or habitat, an evaluation of products purchased in restaurants or supermarkets, or may contribute to a larger “campaign” to assess biodiversity across large areas. It may make sense for you to use sampling techniques from ecology. For example, a quadrat samples the plant and/or animal life in one square meter (or ¼ square meter) of habitat, while a transect collects samples along a fixed path through a habitat. A “Hula Hoop” can be used as an acceptable substitute for a quadrat.Do not take more sample than you need. Only a small amount of tissue is needed for DNA extraction—a piece of plant leaf about ⅛- to ¼-inch diameter or a piece of animal or fungal the size of a grain of rice.

Minimize damage to living plants by collecting a single leaf or bud, or several needles. When possible, use young, fresh leaves or buds. Flexible, non-waxy leaves work best. Tougher materials, such as pine needles or holly leaves, can work if the sample is kept small and is ground well. Dormant leaf buds can often be obtained from bushes and trees that have dropped leaves. Fresh, frozen leaves work well. Dried leaves and herbarium samples are variable.

Avoid twigs or bark. If woody material must be used, select flexible twigs with soft pith inside. As a last resort, scrape a small sample of the softer, growing cambium just beneath the bark. Roots and tubers are a poor choice, because high concentrations of storage starches and other sugars can interfere with DNA extraction.

DNA Extraction

Goal: Extract DNA from tissue samples

Summary

The Chelex, Silica, and Rapid protocol are all simple and efficient DNA extraction methods suitable for the classroom.

• The Rapid DNA extraction protocol works extremely well for plants and for select groups of terrestrial invertebrates. This method eliminates the need for equipment such as water baths and centrifuges, and students can complete the DNA isolation and have DNA ready for PCR in <30 min.
• The Chelex DNA extraction uses chelex resin which binds to substances that can inhibit PCR (polymerase chain reaction, a common DNA analysis technique), like metal ions. The DNA is then released from the sample by heating.
• The Silica DNA extraction method is inexpensive and has the advantage of working reproducibly with almost any kind of plant, fungus, or animal specimen.

More information about the percentage of success of theses DNA extraction methods for different organism types can be found under the "Isolation DNA" section in here.

Pre-isolation preparation: If sample has been stored in 95%+ EtOH prior to DNA isolation, remove the sample from the EtOH using a sterile toothpick and allow it to dry for >10 min on a clean surface. Proceed when sample no longer smells of EtOH. Ethanol carried through the DNA isolation can inhibit DNA amplification later.

Rapid ExtractionChelexSilica

Steps:

1. Obtain tissue ~10 mg or ⅛- to ¼-inch diameter in size by removing a piece of the tissue with a razor blade or sterile tweezer. Be sure to preserve remainder of the organism, as well as additional collected specimens, at -20°C, in 95%+ EtOH, or both. Place tissue in a clean 1.5-mL tube labeled with a sample identification number.
2. Add 50 µL of lysis solution to each tube.

Explanation

Lysis solution dissolves membrane-bound organelles including the nucleus, mitochondria, and chloroplast..

1. Twist a clean plastic pestle against the inner surface of 1.5-mL tube to forcefully grind the tissue for at least 2 minutes. Use a clean pestle for each sample. Ensure the sample is ground into fine particles.

Explanation

Grinding breaks up cell walls and other tough material. Once ground, the sample should be liquid, but there may be some particulate matter remaining.

1. For each sample, use a separate sterile tweezer to add one 3-mm diameter disc of Whatman No. 1 Chromatography paper to the lysed extract. Tap or flick the tube gently to ensure the disc is fully submerged in the extract. Allow the disc to soak in the extract for 1 minute.

Explanation

Whatman chromatography paper binds the DNA, helping separate DNA from contaminants.

1. While the disc is soaking, add 200 µL of wash buffer to a clean 1.5-mL tube labeled with the sample identification number. Wash buffer will remove contaminants that can inhibit PCR while the DNA remains bound to the paper.
2. Remove the disc from the extract using a sterile tweezer or pipette tip and transfer the disc into the fresh tube containing wash buffer. Tap or flick the tube to mix for 5 seconds, then allow the disc to sit in the wash buffer for 1 minute. Discard/set-aside the tweezer following Step 7. Use of the tweezer to transfer the disc in future steps will contaminate the disc with impurities that may affect PCR.
3. Use a sterile pipette tip to gently drag the disc out of the wash buffer and up the tube wall to dry at the top of the tube. Ensure that little to no debris is attached to the disk. Allow the disc to air dry for 2 minutes to evaporate the ethanol on the disc.

Explanation

Ethanol in the wash buffer can inhibit PCR, so drying the paper after the wash step is required.

1. While the disc is air-drying, add 30 µL of TE to a clean 1.5-mL tube labeled with the sample identification number.
2. Once dry, carefully transfer the disc using a sterile tweezer or pipette tip into the fresh tube containing 30 µL of TE. Allow the disc to soak for a minimum of 15 minutes at ambient temperature (soaking the disc overnight at 4° C is optimal) to elute the purified DNA
3. The disc in TE can be stored at 4° C temporarily or frozen at -20° C for long-term storage until ready to begin Part III; ensure that the disc has incubated at ambient temperature for at least 15 minutes before storage at 4° C or -20° C. In Part III, you will use 2 μL of DNA for each PCR reaction. This is a crude DNA extract and contains nucleases that will eventually fragment the DNA at room temperature. Keep the sample cold to limit this activity.

Steps:

1. Obtain tissue ~10 mg or ⅛- to ¼-inch diameter in size by removing a piece of the tissue with a razor blade or sterile tweezer. Some organisms or samples will be small enough that the entire specimen should be used. If you are working with more than one sample, be careful not to cross-contaminate specimens. Be sure to preserve remainder of the organism, as well as additional collected specimens, at -20°C, in 95%+ EtOH, or both.

Explanation

Tissue should be no larger than a grain of rice. Using more than the recommended amount can inhibit amplification.

1. Gently tap 10% Chelex solution tube on a hard surface to ensure the solution is at the bottom. Place tissue into Chelex tube labeled with a sample identification number.

Explanation

Chelex binds positively charged contaminants that could be inhibitors of DNA amplification and binds magnesium ions that are cofactors of DNA nucleases.

1. Twist a clean plastic pestle against the inner surface of Chelex tube to forcefully grind the tissue for at least 2 minutes. Use a clean pestle for each sample. Ensure the sample is ground into fine particles. Securely close the cap of the tube.

Explanation

Grinding breaks up cell walls and other tough material. Once ground, the sample should be liquid, but there may be some particulate matter remaining.

1. Fill a mug nearly to the top with boiling water and cover with aluminum foil. Punch small guide holes in the foil for the number of samples you are processing. Prevent the tube from opening in the following step by using a cap lock to secure the cap to the rim of the tube. Be sure that both the tube rim and cap are held within the cap lock so that steam can’t force the cap open. Place tubes through foil so that the Chelex and sample mixture is fully submerged, but do not submerge the top of the tube.

Explanation

The heat will help to lyse cell and organelle membranes, releasing the DNA.

1. Incubate tubes for 10 minutes. It is not necessary to keep water boiling. Alternatively, you can use a water bath or thermocycler to incubate tubes at 95°C for 10 minutes. Discard water from mug and return the tubes in foil to the mug. Allow Chelex to settle for an additional 10 minutes. Alternatively, place tubes in a balanced configuration in a microcentrifuge, with cap hinges pointing outward. Centrifuge for 30 seconds at maximum speed to pellet Chelex. Once boiled, the DNA and Chelex mixture can be stored at 4° C temporarily, frozen at -20° C for long-term storage, or shipped at ambient temperature.

2. Carefully transfer ~30 µL of supernatant from the Chelex tube, avoiding the Chelex, into a clean 1.5mL microcentrifuge tube labelled with the sample identification number. This tube can be stored at 4° C temporarily or frozen at -20° C for long-term storage until ready to begin Part III. Alternatively, 2 µL of the supernatant from the Chelex tube can be used directly for future PCR reactions. Do not transfer any of the white Chelex resin along with the supernatant. This is a crude DNA extract and contains nucleases that will eventually fragment the DNA at room temperature. Keep the sample cold to limit this activity. Chelex inhibits DNA amplification by PCR.

Steps:

1. Obtain plant, fungal, or animal tissue ~10 mg or ⅛- to ¼-inch diameter by removing a piece of the tissue with a razor blade, clean tweezers, scissors, or back of a 10-µL pipette tip to enable efficient lysis. If you are working with more than one sample, be careful not to cross-contaminate specimens. (If you only have one specimen, make a balance tube with the appropriate volume of water for centrifugation steps.)

Explanation

Tissue should be no larger than a grain of rice. Using more than the recommended amount can affect amplification.

1. Place sample in a clean 1.5-mL tube labeled with an identification number.
2. Add 300 µL of lysis solution to each tube.

Explanation

Lysis solution dissolves membrane-bound organelles including the nucleus, mitochondria, and chloroplast.

1. Twist a clean plastic pestle against the inner surface of the 1.5-mL tube to forcefully grind the tissue for 2 minutes. Use a clean pestle for each tube if you are doing more than one sample.

Explanation

Grinding the tissue breaks up cell walls and other tough material. Once fully ground, the sample should be liquid, but there may be some particulate matter remaining.

1. Incubate the tube in a water bath or heat block at 65° C for 10 minutes.
2. Place your tube and those of other groups in a balanced configuration in a microcentrifuge, with cap hinges pointing outward. Centrifuge for one minute at maximum speed to pellet debris.
3. Label a clean 1.5-mL tube with your sample number. Transfer 150 μL of the supernatant (clear solution above pellet at bottom of tube) to the fresh tube. Be careful not to disturb the debris pellet when transferring the supernatant. Discard old tube containing the debris.
4. Add 3 μL of silica resin to tube; ensure silica resin is mixed and homogenous. Close tube and mix well by flicking or vortexing (solution will turn cloudy, but silica will settle shortly after). Close and incubate the tube for 5 minutes in a water bath or heat block at 57° C.

Explanation

Silica resin is a DNA binding matrix that is white. In the presence of the lysis solution the silica resin binds readily to nucleic acids.

1. Place your tube and those of other groups in a balanced configuration in a microcentrifuge, with cap hinges pointing outward. Centrifuge for 30 seconds at maximum speed to pellet the resin. Use a micropipette with fresh tip to remove all supernatant, being careful not to disrupt the white silica resin pellet at the bottom of the tube.

Explanation

Centrifugation pellets the silica resin, which is now bound to nucleic acid. The pellet will appear as a tiny teardrop-shaped smear or particles on the bottom side of the tube underneath the hinge.

1. Add 500 μL of ice cold wash buffer to the pellet. Mix well by pipetting up and down (or by closing the tube and flicking or vortexing) to resuspend the silica resin.

Explanation

Wash buffer removes contaminants from the sample while nucleic acids remain bound to the resin. The silica resin is not soluble in the wash buffer. The silica resin may stay as a pellet or break up during the washing.

1. Place your tube and those of other groups in a balanced configuration in a microcentrifuge, with cap hinges pointing outward. Centrifuge for 30 seconds at maximum speed to pellet the resin. Use a micropipette with fresh tip to remove all supernatant, being careful not to disrupt the white silica resin pellet at the bottom of the tube.
2. Once again, add 500 μL of ice cold wash buffer to the pellet. Close tube and mix well by vortexing or by pipetting up and down to resuspend the silica resin.

Explanation

Washing twice is much more effective than washing once with twice the volume.

1. Place your tube and those of other groups in a balanced configuration in a microcentrifuge, with cap hinges pointing outward. Centrifuge for 30 seconds at maximum speed to pellet the silica resin.
2. There will be approximately 50 μL of supernatant remaining after the brief spin to be removed. In the presence of water or TE buffer, nucleic acids are eluted from the silica resin.
3. Use a micropipette with fresh tip to remove the supernatant, being careful not to disrupt the white pellet at the bottom of the tube. Spin the tube again for ~15 seconds to collect any drops of supernatant and then remove these with a micropipette.
4. Add 100 μL of distilled water (or TE buffer) to the silica resin and mix well by vortexing or by pipetting up and down. Incubate the mixture at 57° C for 5 minutes.

5. For long-term storage it is recommended DNA samples be stored in TE buffer (Tris/EDTA). Tris provides a pH 8.0 environment to keep DNA and RNA nucleases less active. EDTA further inactivates nucleases by binding cations required by nucleases.

6. Place your tube and those of other groups in a balanced configuration in a microcentrifuge, with cap hinges pointing outward. Centrifuge for 30 seconds at maximum speed to pellet the silica resin.

7. Label a clean 1.5-mL tube with your sample number. Transfer 50 μL of the supernatant (clear solution) to the fresh tube. Be careful not to disturb the pellet when transferring the supernatant. Discard old tube containing the silica resin.

Explanation

Transferring silica resin to the PCR reaction in Part III can inhibit the PCR amplification.

Store your sample on ice or at -20° C until you are ready to begin the PCR.

PCR

Goal: Extract DNA from tissue samples

Summary

Steps:

1. Obtain PCR tube containing Ready-To-Go PCR Bead containing dehydrated Taq polymerase, nucleotides, and buffer. Label the tube with your identification number.
2. Use a micropipette with a fresh tip to add 23 µL of the appropriate primer/loading dye mix to each tube (refer to primer table below). Allow the beads to dissolve for 1 minute at ambient temperature.
3. Place the PCR tubes on ice to prevent premature replication of unwanted primer dimers.
4. Use a micropipette with fresh tip to add 2 µL of your DNA (from Part II) directly into PCR tube with primer and polymerase mixture. Ensure that no DNA remains in the tip after pipetting.

Tip

If the reagents become splattered on the wall of the tube, pool them by briefly spinning the sample in a microcentrifuge (with tube adapters) or by sharply tapping the tube bottom on the lab bench.

If your DNA was extracted using Chelex, allow the tubes containing DNA to sit upright for 10 minutes (or centrifuge for 30 seconds) to ensure that any residual Chelex settles on the bottom of the tubes. When removing DNA for PCR, be careful to only pipet from the very top of the liquid to avoid transferring Chelex into the PCR tube as Chelex inhibits PCR.

1. Store your sample on ice until your class is ready to begin thermal cycling.
2. Place your PCR tube, along with those of the other students, in a thermal cycler that has been programmed with the appropriate PCR protocol.
3. Amplification from some templates, such as the COI barcode region, may be improved by transferring PCR tubes directly from ice into a hot thermal cycler that has been temporarily paused at the beginning of the first 95°C denaturation step. This limits the formation of undesirable primer dimers. Resume the program when all of the PCR tubes are in the thermal cycler.

RbClCOIITS

Primers: (rbcLaF / rbcLa rev)

Temperature	Time	Cyles
94 °C	1:00 minutes	1
94 °C	15 seconds	35
54 °C	15 seconds	35
72 °C	30 seconds	35
4 °C	ad infinitum	1

Primers: (LCO1490 / HC02198)

Temperature	Time	Cyles
94 °C	1:00 minutes	1
94 °C	30 seconds	35
50 °C	30 seconds	35
72 °C	45 seconds	35
4 °C	ad infinitum	1

Primers: (ITS1F / ITS4)

Temperature	Time	Cyles
94 °C	1:00 minutes	1
94 °C	1 minute	35
55 °C	1 seconds	35
72 °C	2 minutes	35
4 °C	ad infinitum	1

AMPure XP Solid Phase Reversible Immobilization (SPRI) Bead PCR Clean-Up (optional)

Summary

This protocol is modified from Beckman Coulter Life Sciences’ protocol for typical PCR cleanup of DNA barcoding PCR products using solid phase reversible immobilization (SPRI) beads (i.e., AMPure XP beads). This protocol will suitably remove the remnants of PCR, selecting for PCR products in the range of 400-800 base pairs, which is consistent with most standard DNA barcode marker regions (e.g., rbcL, COI, etc.). The intent of this protocol is to prepare DNA amplicons for rapid sequencing with ONT’s Rapid Barcoding Kit. Note that it is “optional,” because once all samples are pooled, bead clean-up will still need to be conducted on the pooled amplicon library.

Steps:

1. Re-suspend AMPure XP beads by using the pipette to mix.
2. Add the entire PCR reaction (~20 μL) directly to a labeled 1.5 mL microfuge tube containing 36 μL AMPure XP beads. Use the pipette to carefμLly mix the beads and DNA.
3. Incubate for 5 minutes at room temperature on a microfuge tube rack.
4. Spin down the tube for 15 seconds and place the tube on a magnetic rack. Wait for about 1 minute for the beads to completely pellet near the magnet. Note that the liquid
5. Keeping the tube on the magnet, pipette off the supernatant and discard without disturbing the pellet.
6. Keeping the tube on the magnet, add 200 μL of freshly prepared 70% ethanol without disturbing the pellet. Remove the ethanol using a pipette and discard.
7. Repeat Step 5.
8. Remove the tube from the magnetic rack and spin down the tube for 15 seconds. Replace the tube on the magnetic rack and allow the pellet to reform.
9. Keeping the tube on the magnet, remove any residual liquid (~10-30 μL). With the tube open, allow the pellet to dry for 30 seconds. Caution: Do not allow the pellet to dry to the point of cracking. Be prepared to immediately move on to Step 10.
10. Remove the tube from the magnetic rack and resuspend the pellet in 10 μL of EB. Use the pipette to continuously wash the EB over the pellet until it completely dissociates.
11. Incubate the tube for 5 minutes away from the magnetic rack.
12. Spin down the tube for 15 seconds and replace the tube on the magnetic rack. Wait for about 1 minute for the beads to completely pellet near the magnet. Note that the liquid supernatant shoμLd be completely clear.
13. Remove and retain 10 μL of supernatant. Pipette the supernatant into a thin-walled PCR tube. Discard the tube containing the pellet. The PCR product is now ready for the ONT Rapid Barcode Kit.

Rapid Barcode Index Attachment

Summary

After the optional SPRI bead clean-up step, DNA amplicons are ready for rapid barcode index attachment. Note that up to 24 DNA amplicons can be uniquely tagged during a single library prep and sequencing run using the Rapid Barcode Kit 24 V14. ONT manufactures another kit with up to 96 unique barcode indexes.

Steps

1. Obtain one unique Rapid Barcode (RB) mix for each PCR amplicon. Record each sample number and its associated RB.
2. Spin down RB tubes. Pipette 1 μL of each unique RB directly into a PCR tube containing 10 μL of its associated DNA amplicon. Vortex briefly or pipette to mix.
3. Ensure that PCR tubes are sealed shut, and then incubate tubes in a thermal cycler using the following conditions:

Temperature	Time
30 °C	2:00 minutes
80 °C	2:00 minutes
4 °C	hold

Explanation

The 20 °C incubation step will first ensure optimal transposase activity, promoting the addition of the RB indexes. The 80 °C step will denature and inactivate the transposase to prevent erroneous addition of unique indexes to the wrong DNA amplicons during pooling.

1. Spin down PCR tubes. DNA amplicons are now uniquely tagged using the ONT RBs.
2. Pipette each tagged DNA amplicon (~11.5 μL) into a single class-wide 1.5 mL microfuge tube. This represents the pooled amplicon library.

Library Clean-Up with Solid Phase Reversible Immobilization (SPRI) Beads

Summary

Library clean-up will remove remnants of PCR (e.g., primers, polymerase, etc.), residual transposome and unused rapid barcodes, as well as any dyes or indicators used during previous steps (e.g., Cresol red). Note that this step must be carried out even if the optional PCR clean-up step was performed with a class.

Steps:

1. Re-suspend AMPure XP beads by using the pipette to mix or vortexing.
2. Add an appropriate volume of AMPure XP beads to the pooled amplicon library using the following formula: VolumeAMPure XP beads= (11.5 μL)(No. of DNA amplicon samples)
3. Use a pipette to gently mix the beads and amplicon library such that the beads are evenly distributed throughout the tube.
4. Incubate the mix at room temperature for 10 minutes, flicking occasionally to ensure that beads remain suspended in solution. Note that DNA amplicons are coordinating with SPRI beads during this time.
5. Spin down the mix for 15 minutes to form an initial bead pellet. Then, place the tube on a magnetic rack to pellet the beads. Allow up to one minute for the beads to pellet. Keeping the tube on the magnetic rack, use an appropriate pipette to remove all of the liquid from the tube and discard. Avoid disturbing the pellet. Remember that the DNA is located in the pellet at this point.
6. Keeping the tube on the magnetic rack, add 1000 μL of 80% ethanol to the pellet. Then, remove the supernatant and discard without disturbing the pellet.
7. Repeat Step 6.
8. Remove the tube from the magnetic rack and spin down the tube for 15 seconds. Replace the tube on the magnetic rack and allow the pellet to reform.
9. Keeping the tube on the magnet, remove any residual liquid (~10-30 μL). With the tube open, allow the pellet to dry for 30 seconds. Caution: Do not allow the pellet to dry to the point of cracking. Be prepared to immediately move on to Step 10.
10. Remove the tube from the magnetic rack and resuspend the pellet in 15 μL of EB. Use the pipette to continuously wash the EB over the pellet until it completely dissociates. Note that this may take several repeated washes with the same 15 μL EB.
11. Incubate the tube for 10 minutes away from the magnetic rack at room temperature.
12. Spin down the tube for 15 seconds and replace the tube on the magnetic rack. Wait for about 1 minute for the beads to completely pellet near the magnet. Note that the liquid supernatant shoμLd be completely clear.
13. Remove and retain 11 μL of supernatant. Pipette the supernatant into a 1.5 mL microfuge tube. Discard the tube containing the pellet. The pooled amplicon library is now ready for the attachment of rapid adapters and motor proteins.

Rapid Adapter Attachment

Summary

The final step of the library preparation is attaching the motor protein to the ends of the DNA. The Rapid Adapter (RA) reagent contains the motor protein and can attach it to the rapid adaptor chemistry on the ends of the DNA which were added by the transposome complex in a previous step.

Steps:

1. In a 1.5ml tube dilute Rapid Adapter (RA) as follows:

Reagent	Volume
Rapid Adapter (RA)	1.5ul
Adapter Buffer (ADB)	3.5ul
Total	5ul

1. Add 1ul of diluted Rapid Adapter to indexed DNA.
2. Incubate for 10 min at room temperature.

Flow Cell Check

Summary

The flow cell contains the nanopores which facilitate the sequencing. The number of active pores needs to be determined for every flow cell before it is to be used to ensure that there are an adequate number for the sequencing experiment. Because we are sequencing amplicons and don’t need very high-throughput, we suggest using the Flongle flow cell which has a maximum of 126 pores rather than the Minion, which has maximum of 2048.

Steps: 1. Open MinKNOW software. 2. Click the “Start” tab on the upper left of the screen. 3. Click “Flow cell check”

!!! warning "Important' The sequencer will take a few minutes to get to the proper temperature and run the flow cell check. When complete, the number of usable pores on the flow cell will be displayed. The flow cell is considered under warranty if it is checked within 2 weeks of the delivery date, was properly stored at 4 degrees, but has less than 50 pores. If under warranty, contact Oxford Nanopore within 2 days of performing the flow cell check.

Flow Cell Priming

Summary

Before loading the flow cell with the DNA library, the flow cell needs to be “primed” for sequencing. The reagents required for priming and loading the Flongle flow cell are contained in the Flongle Sequencing Expansion (EXP-FSE002), a separate box from the library prep reagents, which contains glass vials of Sequencing Buffer (SB), Flow Cell Flush (FCF), and Library Beads (LIB).

1. Mix 117ul FCF (from the Flongle expansion kit) and 3ul of FCT (from the Rapid Barcoding kit).
2. Peel back the seal from the Flongle flow cell.
3. Slowly expel liquid from the pipette by turning the dial clockwise until a bead of liquid forms at the tip.
4. Place the pipette tip in the sample port, keeping the pipette vertical.
5. Turn the dial clockwise to load the flush buffer and primer into the flow cell, stopping right before all liquid is entirely expelled to avoid pipetting air into the flow cell.

Flow Cell Loading

1. Make a sequencing mix:

Reagent	Volume
Sequencing Buffer (SB)	15ul
Loading Beads (LIB)	10ul
DNA Library	5ul
Total	30ul

1. Pipette up sequencing mix, slowly expel liquid from the pipette by turning the dial clockwise until a bead of liquid forms at the tip.
2. Place the pipette tip in the sample port, keeping the pipette vertical.
3. Turn the dial clockwise to load to the flush buffer and primer the flow cell, stopping right before all liquid is entirely expelled to avoid pipetting air into the flow cell.
4. Place the sticker back on.
5. Proceed to "Start Sequencing" on MinKNOW software.

Bioinformatics

Full DNA Barcoding Analysis of DNA barcodes can be done using DNA Subway2.0 (coming soon).

Epi2melabs wf-amplicon may also be used to generate consensus sequences for each sample. This conensus sequence can

Data analysis

Goal: Determine the taxa sampled.

Example data

Plant DNA Barcoding :

This dataset was generated at a workshop at James Madison University in June 2024.

• Nanopore sequencing report example: Download
• FastQ files: Download
• Example EPI2ME report: Download

Bioinformatics

This tutorial will use the EPI2ME amplicon workflow developed by Oxford Nanopore.

Workflow version: 1.1.3

Instruction tip

Students can work individually or in groups to analyze their data. Assuming each student sample is assigned their own barcoded samples, they can generate an individual report specific to their example.

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Analyze data

1. Open the EPI2ME software and choose either Sign in or Continue as guest.

   Bioinformatics

   In this example, we use the Continue as guest option.

2. Click on the Launch icon and choose the Amplicon workflow.

   Bioinformatics

   In this example, choose Run Locally.

   Failure

   If you see the "Setup required" error message, follow the instructions to install or configure any needed software (for example, installing and starting Docker).

   Option

   Depending on your access and your computer, the launch page is where you will decide on running the workflow locally (on your computer) or on the cloud.

3. Click on the Launch button to proceed with the workflow.

4. Under Setup local analysis, complete the required options to load your sequencing dataset and choose analysis parameters.

   Bioinformatics

   In this example, we use the following parameters:

   Input Options

   • FASTQ: "fastq_pass" folder provided in this tutorial example data. This folder will be generated by your experiment.
   • Analyse unclassified reads: Leave as default.
   • Reference: Leave as default.

   Sample Options

   • All options left as default.

   Pre-processing Options

   • All other options left as default.

   Variant calling options

   • All options left as default.

   Output Options

   • All options left as default.

   Advanced Options

   • All options left as default.

   Miscellaneous Options

   • All options left as default.

   Nextflow Configuration

   • All options left as default.

   Nextflow Options

   • All options left as default.

5. Click Launch workflow and then Launch again.

   Time

   The time required for this analysis depends on your dataset. On a computer with the recommended configuration for running a MinION, the test dataset was completed in approximately 5 minutes.

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Interpret results

The wf-amplicon pipeline generates a detailed report (wf-amplicon-report.html) that provides comprehensive insights into sequencing data. However, the file we will need for our analysis in in the folder "output" generated by epi2melabs afer the run. The output folder contains folders for each sample (i.e. "barcode01"), which contain a folder named "consensus". A .fasta file in the consensus folder is a consensus DNA sequence of the DNA Barcoding amplicon from all the sequencing reads for the sample.

The DNA sequence from the .fasta file can be copied and pasted into the blue line of DNA Subway under "Enter sequences in FASTA format".

Instruction tip

DNA Subway 2.0 is coming late Spring/Summer 2025 and is equiped with a new "Blue Line" for Nanopore sequencing data which contains tools for generating the consensus Nanopore sequence reads.

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