Apr 14, 2025

Amplicon PCR & Library Preparation for Illumina Sequencing

Environmental DNA
DOI
https://dx.doi.org/10.17504/protocols.io.261gerqxol47/v1
  • 1School of Biosciences, Western Bank, University of Sheffield, Sheffield, S10 2TN, UK
NEOF - NERC Environmental Omics Facility VF
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DOI: https://dx.doi.org/10.17504/protocols.io.261gerqxol47/v1
External link: https://doi.org/10.1002/edn3.70203
Protocol Citation: NEOF - NERC Environmental Omics Facility VF, Helen Hipperson, Kathryn H. Maher, Gavin J. Horsburgh, Lucy S. Knowles 2025. Amplicon PCR & Library Preparation for Illumina Sequencing. protocols.io https://dx.doi.org/10.17504/protocols.io.261gerqxol47/v1
Manuscript citation:
Dina‐Leigh Simons, Helen Hipperson, Tom J. Webb, Matthew Spencer, Gavin John Horsburgh, Nova Mieszkowska (2025) Characterizing Rocky Intertidal Biodiversity Using Environmental DNA Metabarcoding From Local to National Scales.Environmental DNA doi: 10.1002/edn3.70203
License: This is an open access protocol distributed under the terms of the Creative Commons Attribution License,  which permits unrestricted use, distribution, and reproduction in any medium, provided the original author and source are credited
Protocol status: Working
We use this protocol and it's working
Created: March 10, 2025
Last Modified: April 14, 2025
Protocol Integer ID: 124106
Keywords: Metabarcoding, Amplicon, Illumina, 2-step PCR, Library, sequencing, prep, protocol, cost-effective, preparation for illumina next generation sequencing, illumina next generation sequencing, amplicon sequences from multiple sample, amplicon pcr, generating amplicon sequence, sequencing, metabarcoding protocol, amplicon, library preparation for illumina, identifying tag sequence, pcr2, pcr, pcr1, tag sequence, using qpcr, multiple sample, sample, unique sample, library preparation, generation sequencing, qpcr
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Abstract
This metabarcoding protocol is for generating amplicon sequences from multiple samples in preparation for Illumina Next Generation Sequencing.

Amplicons are generated for each sample (PCR1) then cleaned using magnetic beads. Unique sample identifying tag sequences are attached to each sample (PCR2), pooled, cleaned again and finally quantified using qPCR.
Materials
  • Amplicon-specific primers with Illumina sequencing primer sites (see Step 1)
  • 0.2ml 96-well PCR plates
  • 1.5ml microcentrifuge tubes
  • Meridian Bioscience MyTaq HS Mix (BIO25046)*
  • Thermocycler
  • Electrophoresis gel equipment and reagents
  • Promega ProNex‱ Bead Kit (NG2003)*
  • 96-well and microcentrifuge tube magnetic racks
  • Fi5 & Fi7 IDT Primers (see Step 3)
  • Agilent D1000 ScreenTapes (5067-5582) and buffer (5067-5602)*
  • Promega QuantiFluor dsDNA System (E2670)*
  • Real-time PCR thermal cycler
  • KAPA Library Quantification (KK4873)*
  • Low TE (Tris10mM,EDTA0.1mM) recipe for 500ml:
5ml 1M Tris-HCl (pH 8.0)
100µl 0.5M EDTA (pH8.0)
495ml ddH2O
Autoclave.

*Can be subsituted.
Safety warnings
Use Good Lab Practice and wear protective equipment. Avoid contact with skin. All chemicals can be disposed of down the sink with copious amounts of water to dilute the ethanol down to >20% of the waste.
PCR1
Using amplicon-specific primers (XX) tailed with Illumina sequencing primer sites (F – small RNA primer; R – read 2 primer), e.g.:

F 5’-3’; ACACTCTTTCCCTACACGACGCTCTTCCGATCTNNNNNXXXXXXXXXXXXXXXXXXXX
R 5’-3’; GTGACTGGAGTTCAGACGTGTGCTCTTCCGATCTXXXXXXXXXXXXXXXXXXX

PCR1 adds 72bp to your product size due to adapters.

Controls
When running any PCR amplification reaction, positive and negative controls should be added to determine if the PCR is working as expected (+ve control) and to make sure no contamination is being introduced (-ve control). Negative controls from the DNA extraction may also be included. You may also choose to add replicates of either/both extraction and PCR.

PCR the Amplicon/s from your DNA
AB
µL/reactionReagent
10Meridian Bioscience MyTaq HS Mix
2F primer (at 1 to 5 μM)*
2R primer (at 1 to 5 μM)*
5Sterile ddH2O
1⧫DNA (at ~10 ng/μl)
Table 1: PCR1 Recipe
* You may need to optimise the amount of primer used to avoid excess/unincorporated/dimer being carried through to the next step, while still getting good/strong amplification.
⧫ If your DNA concentration is particularly low (which can be the case in studies on diet & eDNA) increase the amount of DNA you use and decrease the amount of ddH2O accordingly.

PCR program:
95°C for 5 min
Followed by 25-35 cycles of:
94°C for 30 sec
Tm°C* for 30 secs to 1min 30sec *this depends on the Tm of your specific primers
72°C for 30 secs to 2 min (depending on length)
Then:
72°C for 10 min
Electrophoresis Agarose Gel
Run 4μl of PCR product on a 1% agarose gel along with a 100bp ladder to make sure samples have amplified, the amplicon is the correct size and that there is a minimal amount of primer dimer.

Bead Cleaning
Prior to PCR2 we need to bead clean the samples to remove short extraneous DNA fragments and reagents left over from PCR1.
Purification with Promega Pronex‱ Beads
  1. Allow an aliquot of beads to reach RT (30 min), and vortex to resuspend them.
  2. Add a ratio of resuspended Pronex beads appropriate for your amplicon size (see Table 2). Mix well by pipetting up and down 10 times and incubate at RT for 5-10 min.
  3. Place on a magnetic rack to separate beads from the solution. When the liquid is completely clear, aspirate the supernatant and keep in a new plate (just in case your target amplicon is accidentally removed). Do not disturb the pellet of separated magnetic beads and do not remove the samples from the magnetic plate.
  4. With the samples still on the magnetic place, add 100 μl of ProNex Wash Buffer (or 80% ethanol). Incubate at RT for 30-60 seconds, then carefully aspirate out and discard.
  5. Repeat step 4.
  6. Allow the beads to dry at RT for 5-10 min, while on the magnetic stand. Drying will allow traces of ethanol to evaporate, but over-drying the beads (if the pellet cracks) can significantly decrease elution efficiency.
  7. Remove samples from the magnetic plate and elute with 15 μl of Low TE (get a fresh sterile aliquot to ensure no contamination). Mix well by pipetting up and down 10 times (ensure that the beads are fully immersed & mixed with the Low TE). Incubate at RT for 5-10 min.
  8. Place on a magnetic rack to separate beads from the solution (~ 1 min).
  9. Remove the solution containing your DNA to a fresh plate without carrying over any beads.

ABC
Approximate Size Cutoff (bp)ProNex® Chemistry Ratio (v/v)Estimated Equivalent AMPure® XP Ratio
1003X1.8X
1502X1.5X
2501.5X0.95X
3501.3X0.75X
4751.2X0.65X
5501.15X0.625X
6501.1X0.6X
8001.05X0.55X
10001X0.5X
Table 2: Ratios for Size-Selective Purification of DNA Fragments
The ratios of ProNex Chemistry to sample are higher than those used in other methods with similar size-selective chemistries. If you have experience with another size-selective DNA purification product, such as AMPure‱ XP or SPRIselect‱, do not use the same ratios of binding reagent to sample, as this will result in the loss of desired fragments.

PCR2
Useing tailed PCR to add unique identifier sequences (dual-plexed: Fi5 and Ri7 primers in unique combination for each sample) and Illumina sequencing sites to the amplicon products.

The general sequences of the forward and reverse primers with included 10bp barcode are illustrated below.

Fi5_01
5’CAAGCAGAAGACGGCATACGAGATTATCTTCTCGGTGACTGGAGTTCAGACGTGTGCTCTTCCGATC*T3’
Ri7_01
5’AATGATACGGCGACCACCGAGATCTACACCGTCGCCTATACACTCTTTCCCTACACGACGCTCTTCCGATC*T3’

PCR2 adds 73 bp to your overall product size.

PCR to Add Unique Identifying Sequences

AB
µL/reactionReagent
10Meridian Bioscience MyTaq HS Mix
0.5Fi5 Primer (at 10 μM)*
0.5Fi7 Primer (at 10 μM)*
1Sterile ddH2O
8Template from PCR1
Table 3: PCR2 Recipe
*IDT Illumina Unique Dual (UD) Indexes, 1536 unique indexes available.
PCR program:
95°C for 5 min
Followed by ⧫8-12 cycles of:
98°C for 10 sec
65°C* for 30 secs
72°C for 30 secs
Then:
72°C for 10 min

⧫ Determine how many cycles you need from the concentration.

TapeStation

Run a selection (~4 per 96-well plate) of pre and post PCR2 samples on the Agilent TapeStation or BioAnalyser to check that the samples have increased in size from the addition of PCR2 primers.

Figure 2: Example TapeStation image of two samples' PCR1 and PCR2 products.

Quantification and Pooling
Quantification
Quantify all PCR2 products using a fluorometer and the Promega QuantifFluor dsDNA kit. Qubit and other fluorometers may be used, Nanodrop is not recommended.

  1. Take 1ml of 20X TBE from the Promega QuantiFluor dsDNA kit and add 19ml ddH2O.
  2. Add 50µl of the Quantifluor dsDNA dye and vortex. This is your Quantifluor mix.
  3. Add 2µl of a set of pre-made standards to a dry BMG black plate (0, 3.125, 6.25, 12.5, 25, 50, 100ng/µl)*.
  4. Add 2µl of PCR2 product to the black plate.
  5. Add 200µl of the Quantifluor mix to each well of your DNA and standards in the black plate.
  6. Read the plate in the BioTek Fluorometer.
  7. Leftover Quantifluor can be kept at 4°C, wrapped in foil, for future use.

* Mix well before use. Standards can be created by making a serial dilution of the DNA standard from the Quantifluor kit and Low TE.

Pooling
Using the formula below calculate the volumes of each sample so that each sample is evenly represented when pooling together. Add 8 - 24 samples into pools.


Pool Bead Cleaning
Prior to sequencing we need to bead clean the samples to remove short extraneous DNA fragments and reagents left over from PCR2.

Purification with Promega Pronex‱ Beads
  1. Allow an aliquot of beads to reach RT (30 min), and vortex to resuspend them.
  2. Add a ratio of resuspended Pronex beads appropriate for your amplicon size (see Table 2). Mix well by pipetting up and down 10 times and incubate at RT for 5-10 minutes.
  3. Place on a magnetic rack to separate beads from the solution. When the liquid is completely clear, aspirate the supernatant and save. Do not disturb the pellet of separated magnetic beads and do not remove the samples from the magnetic plate.
  4. With the samples still on the magnetic rack, add 200μl of ProNex Wash Buffer (or 80% ethanol). Incubate at RT for 30-60 seconds, then carefully aspirate out and discard.
  5. Repeat step 4.
  6. Allow the beads to dry at RT for a 5-10 min, while on the magnetic rack. Drying will allow traces of ethanol to evaporate, but over-drying the beads (if the pellet cracks) can significantly decrease elution efficiency.
  7. Remove samples from the magnetic rack and elute with 25μl of Low TE. Mix well by pipetting up and down 10 times (ensure that the beads are fully immersed & mixed with the low TE). Incubate at RT for 5-10 min.
  8. Place on a magnetic rack to separate beads from the solution (~ 1 min).
  9. Remove the solution containing your DNA to a fresh tube without carrying over any beads.

TapeStation
If there is no primer dimer present at Step 4, you may skip this step.

Run all pools on the Agilent Tapestation or BioAnalyser to check that no primer dimer is present, this will be a small peak shorter than your amplicon size (usually 50-150bp). If primer dimer is shown, repeat the bead clean and check again.

If the primer dimer contamination is large or cannot be removed efficiently with beads, the libraries may be run a Sage Science BluePippin.

Library Quantification
Due to the presence of unlabelled DNA from PCR2, quantification of the library by qPCR is most accurate. Using a commercial kit we can quantify products for Illumina sequencing on the a qPCR that only target DNA with i5 and i7 indexes attached.

KAPA qPCR
  1. Make a serial dilution of each library: 100, 1000 and 10000 -fold. 
  2. For 100-fold dilution, Combine 99µl of the dilution buffer with 1µl of the library. Vortex. 
  3. For 1000-fold dilution, take out 1µl from 1st dilution (100-fold) and add 9µl of dilution buffer. Vortex. 
  4. For 10000–fold, take out 1µl from 2nd dilution (100-fold) and add 9µl of dilution buffer. Vortex.
  5. Repeat steps 2-4 two times to produce three independent dilutions of the library.
  6. Prepare KAPA Library Quantification Kit master mix with 6µl of SYBR FAST primer mix and 2µl of sterile ddH20 per well.
  7. Add 2µl of the kit standards, diluted sample libraries, or dilution buffer (no template control) to appropriate wells in the optical 96-well plate.
  8. Dispense 8µl of the master mix to the appropriate wells in the 96-well plate.
  9. Spin plate and ensure that there are no bubbles at the bottom of wells or debris on the bottom of the plate. Cover plate with an optical adhesive film.
  10. Run the plate on a qPCR machine with the following profile:
95°C for 5 min
Followed by 35 cycles of:
95°C for 30 sec
60°C for 45 sec

Pool into Final Library
Pool samples in equimolar amounts into one tube (per sequencing run).

The final library is now ready to be sequenced, you may wish to dilute the library and re-quantify using qPCR. Most Illumina sequencing protocols dilute from a concentration of 4nM though commercial companies usually ask for ≥15nM in 20-30µl.
Protocol references
Campbell, N.R., Harmon, S.A. and Narum, S.R. (2014) ‘Genotyping‐in‐thousands by sequencing (GT‐SEQ): A cost effective SNP genotyping method based on custom amplicon sequencing’, Molecular Ecology Resources, 15(4), pp. 855–867. doi:10.1111/1755-0998.12357.