Sep 11, 2025
Version 2
Illumina Dual Index Amplicon Sequencing Sample Preparation 16S (515F–806R primer pair) V.2
This protocol is a draft, published without a DOI.
- Alexander Eiler1,
- Eivind Stensrud2
- 1Alex private;
- 2eDNA solutions AB
- eDNAsolutionsTech. support phone: +46 (704) 139-481 email: eivind@ednasolutions.se
Protocol Citation: Alexander Eiler, Eivind Stensrud 2025. Illumina Dual Index Amplicon Sequencing Sample Preparation 16S (515F–806R primer pair). protocols.io https://protocols.io/view/illumina-dual-index-amplicon-sequencing-sample-pre-g7ukbznuxVersion created by Eivind Stensrud
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: August 18, 2025
Last Modified: September 11, 2025
Protocol Integer ID: 224876
Keywords: illumina dual index amplicon sequencing sample preparation, earth microbiome project primer, primer pair, preparation of pcr product, pcr product, amplicon, earth microbiome project, primer, primer set
Abstract
Preparation of PCR products for amplicon sequencing using The Earth Microbiome Project 515F–806R primer set .
Guidelines
| Index 2 (i5) | Index 2 (i5) Sequence | Index 1 (i7) | Index 1 (i7) Sequence | |
| Illu_N501F | TAGATCGC | Illu_N701R | TCGCCTTA | |
| Illu_N502F | CTCTCTAT | Illu_N702R | CTAGTACG | |
| Illu_N503F | TATCCTCT | Illu_N703R | TTCTGCCT | |
| Illu_N504F | AGAGTAGA | Illu_N704R | GCTCAGGA | |
| Illu_N505F | GTAAGGAG | Illu_N705R | AGGAGTCC | |
| Illu_N506F | ACTGCATA | Illu_N706R | CATGCCTA | |
| Illu_N507F | AAGGAGTA | Illu_N707R | GTAGAGAG | |
| Illu_N508F | CTAAGCCT | Illu_N708R | CCTCTCTG | |
| Illu_N521F | CTTGCTTT | Illu_N709R | AGCGTAGC | |
| Illu_N522F | GGCTTCAA | Illu_N710R | CAGCCTCG | |
| Illu_N523F | AATCGGCA | Illu_N711R | TGCCTCTT | |
| Illu_N524F | GGTTCAAA | Illu_N712R | TCCTCTAC | |
| Illu_N525F | ACTTCGAC | Illu_N733R | GGTATAAG | |
| Illu_N526F | TGACTTGC | Illu_N735R | CAGCTAGA | |
| Illu_N527F | TAGGACCT | Illu_N736R | CCATAGCA | |
| Illu_N528F | GGAGACTT | Illu_N738R | GGTATAGC | |
| Illu_N529F | AGGTTACG | Illu_N739R | GGTTATGC | |
| Illu_N530F | AATTCGCT | Illu_N740R | TAGGCAAG | |
| Illu_N531F | TCAGCTAA | Illu_N741R | TTGTCCAT | |
| Illu_N532F | GCGATATG | Illu_N743R | TCTAGGCA |
Materials
Thermo Scientific, dNTP Mix (2 mM each): R0242
New England Biolabs, Q5 High-Fidelity DNA Polymerase: M0491LVIAL
New England Biolabs, Q5 Reaction Buffer: B9027SVIAL
Eurofins, NGSgrade Oligos (HPLC purified) 515F-806 primer pair
Thermo Fisher Scientific, Invitrogen, TE, pH 8.0, RNase-free: AM9858
Thermo Fisher Scientific, Invitrogen, UltraPure DNase/RNase-Free Distilled Water: 10977035
Troubleshooting
Abstract
Preparation of PCR products for amplicon sequencing using The Earth Microbiome Project primer set, 515F–806R.
Introduction
Standard procedure for preparing libraries for metabarcoding using the 515F–806R primer pair (Caporaso et al., 2011) from the Earth Microbiome Project
The primer set is regarded as near universal for prokaryotes (bacteria and archaea) and the original designed primers were modified by Parada et al. 2016 (https://doi.org/10.1111/1462-2920.13023) and Appril et al. 2015 (doi: 10.3354/ame01753).
To improve the quality of the analysis, eDNA solutions adapted the workflow to work with New England Biolabs Q5 High-Fidelity DNA Polymerase and lower cycling numbers than the published workflow at Earth Microbiome Project.
Safety warnings
The lab work should be conducted in designated rooms for DNA-extractions, pre-PCR and post-PCR.
Perform plate set-up in the pre-PCR lab, and all PCR products should only be worked with in the post-PCR.
These rooms should have a strict one-way flow, meaning no equipment or samples should be moved from post-PCR to pre-PCR.
Always include (a) positive control(s) with a known mock-community, and add (a) negative plate control(s) in addition to the negative control(s) from the DNA extraction.
Before starting
Ensure that metadata files are finalised before starting library preparations in the lab.
Necessary equipment:
Pipettes P10, P100/P200 + P1000
Pipette tips (w + w/o filters)
Eppendorf tubes
96-well plate / PCR strips w/ lids
96-well plate seal
Seal applicator
Microplate rack / PCR strip rack
Clean bench and equipment (ex. pipettes and racks) with 10% chlorine, wait 10 minutes, then proceed to clean with 70% Ethanol and wait additional 10 minutes.
If working in an UV-hood, to sterilize the hood, turn on the UV for 30 minutes.
Avoid using chlorine on metal, and ethanol on hood plastics.
DNA extraction
Conduct the DNA extraction, we recommend using the DNeasy PowerWater Sterivex Kit (remember to add negative DNA extraction(s)) for water samples. Follow the manufacturerer recommendation with the exeption of a dual elution of 100 µl volume. (https://www.qiagen.com/us/products/discovery-and-translational-research/dna-rna-purification/dna-purification/microbial-dna/dneasy-powerwater-sterivex-kit)
Quantify the extracted DNA using PicoGreen.
Primers
Earth Microbiome primer set (515F–806R primers)
NGSgrade Oligos (HPLC purified) (Eurofins), the primer set has attached Illumina adapters from Sinclair et al. 2015. https://doi.org/10.1371/journal.pone.0116955 and Juottonen et al. https://doi.org/10.1111/1462-2920.15058 incorporate random nucleotides to reduce amplification bias in the forward primer.
Resuspend the primers with TE Buffer or Nuclease-Free water to obtain a 100 μM
stock solution (store in -20 ℃).
To reduce risk of cross-contamination and primer degradation, we recommend to make several aliquots of 10 μM primers by diluting with either TE-buffer or Nuclease-Free Water.
Forward primer (56mer):
515F (Parada) (5'-3'): ACA CTC TTT CCC TAC ACG ACG CTC TTC CGA TCT NNNN GTG YCA GCM GCC GCG GTA A
Reverse primer (41mer):
806R (Apprill) (5'-3'): AGA CGT GTG CTC TTC CGA TCT GGA CTA CNV GGG TWT CTA AT
Illumina adapters and spacers are marked in bold, while the taxa specific primers are in roman.
First PCR reactions
Work in pre-PCR room.
Calculate the volumes of each reagent needed to create the Mastermix for the number of samples you are running (+10% extra to account for pipetting lost and potential errors).
Remember to run the samples in duplicates/triplicates to reduce PCR-stochasticity.
| A | B | C | D | E | |
| Reagent | Stock conc. | Mastermix conc. | Volume 1 reaction (µl) | Volume X reactions (µl) | |
| 5X Q5 Reaction Buffer | 5X | 1X | 5 | ||
| Forward primer | 10 µM | 0,2 µM | 0,5 | ||
| Reverse primer | 10 µM | 0,2 µM | 0,5 | ||
| dNTP Mix | 2 mM each | 40 µM each | 0,5 | ||
| Q5 High-Fidelity DNA Polymerase | 0,25 | ||||
| Nuclease-Free Water | 16,25 | ||||
| Template DNA | 2* | ||||
| Total | 25 |
PCR cycling conditions has been optimized with an Applied Biosystems QuantStudio 3.
Add up to 2 µl DNA template from environmental samples. For positive control, add 1 µl mock community template and 1 µl Nuclease-Free water. For negative control, add 2 µl Nuclease-Free water.
Remember to wear gloves and lab coat. Change gloves whenever necessary.
Clean the working area and equipment with 10% chlorine, wait 10 minutes before continuing.
Clean the working area and equipment with 70% ethanol, wait 10 minutes before continuing.
Briefly vortex and spin down all reagents in a microcentrifuge, except the polymerase. Mix the polymerase by pipette mixing 10 times and spin briefly down in a microcentrifuge.
Prepare a Mastermix according to Table 1. Remember to make 10% extra to account for pipetting loss and potential errors.
Vortex and spin down Mastermix in a microcentrifuge.
Add 23 µl Mastermix to each well according to plate layout.
Add 2 µl template DNA according to plate layout.
Add 2 µl Negative control according to plate layout.
Add 1 µl mock community + 1 µl PCR water according to plate layout
Seal the plate, vortex and spin the plate down with a centrifuge.
Run PCR using the cycling conditions described in the Table below.
PCR cycling conditions (first round of PCR)
| A | B | C | D | |
| STEP | TEMP | TIME | ||
| Initial Denaturation | 98°C | 2 minutes | ||
| 20-35 cycles* | Denaturation | 98°C | 10 seconds | |
| Annealing | 50°C | 30 seconds | ||
| Extension | 72°C | 20 seconds | ||
| Final extension | 72°C | 2 minutes | ||
| Hold | 4°C | ∞ |
PCR cycling conditions have been optimized with an Applied Biosystems QuantStudio 3.
Aim to use as few cycles as possible, we recommend to run a few test samples with 25 cycles.
PCR visualization
Conduct in post-PCR lab.
After the first PCR reaction, pool the replicated samples and inspect the PCR products using Agarose gel electrophoresis (1%).
Successful amplification should give bands at ~390 bp (See picture below).
Sample overview in the wells:
1: 100 bp ladder, 2: Empty, 3-8: Environmental samples, 9: PCR negative, 10: Mock community.
Purification
Conduct in post-PCR.
Perform purification with magnetic beads (Cytiva Sera-Mag Select) using 0.8x beads concentration following the manufacturer recommendations.
Second PCR reactions
A second PCR is conducted for attaching standard illumina handles (bold) and index primers
Multiplex_fwd:
AATGATACGGCGACCACCGAGA{TCTACAC}-[i5 index] ACACTCTTTCCCTACACGACG
Multiplex_rev:
CAAGCAGAAGACGGCATACGAGAT-[i7 index]-GTGACTGGAGTTCAGACGTGTGCTCTTCCGATCT
We use 20 different forward index/barcode primers and 20 different reverse index/barcode primers.
Combining both primers (20X20) makes it possible to generate 400 unique tags to run 400 samples of each unique primer set in one final pool for sequencing.
(See Guidelines for index sequences)
Second PCR cycling conditions
Prepare Mastermix and mix with forward and reverse indices according to plate layout in the pre-PCR lab.
Bring the plate(s) with Mastermix and primers to the post-PCR lab, where samples from the 1st PCR reaction is added according to plate layout.
Calculate the volumes of each reagent needed to create the Mastermix for the number of samples you are running (+10% extra to account for pipetting lost and potential errors).
| A | B | C | D | E | |
| Reagent | Stock conc. | Mastermix conc. | Volume 1 reaction (µl) | Volume X reactions (µl) | |
| 5xQ5 Reactio Buffer | 5X | 1X | 4,00 | ||
| Forward index (i5, illu-N5xx) | 5 μM | 0,25 μM | 1,00 | ||
| Reverse index (i7, illu-N7xx) | 5 μM | 0,25 μM | 1,00 | ||
| dNTPs | 2 mM each | 200 μM each | 2,00 | ||
| Q5 HF DNA polymerase | 2 U/μl | 0,02 U/μl | 0,20 | ||
| Template from 1st PCR | 2,00 | ||||
| Nuclease-Free water | 9,80 | ||||
| ∑ | 20,00 |
PCR cycling conditions has been optimized with an Applied Biosystems QuantStudio 3.
Add 2 µl of purified PCR product of both environmental samples, positive and negative controls.
Remember to wear gloves and lab coat. Change gloves whenever necessary.
Clean the working area and equipment with 10% chlorine, wait 10 minutes before continuing.
Clean the working area and equipment with 70% ethanol, wait 10 minutes before continuing.
Briefly vortex and spin down all reagents in a microcentrifuge, except the polymerase. Mix the polymerase by pipette mixing 10 times and spin briefly down in a microcentrifuge.
Prepare a Mastermix according to Table 1. Remember to make 10% extra to account for pipetting loss and potential errors.
Vortex and spin down Mastermix in a microcentrifuge.
Add 16 µl Mastermix to each well according to plate layout.
Add 1 µl Forward and Reverse primer (unique combination per sample) to each well on the 96-well plate.
Move to Post-PCR
Seal the plate while transfering to Post-PCR room.
Add 2 µl template DNA according to plate layout.
Seal the plate, vortex and spin the plate down with a centrifuge.
Run the second PCR using the cycling conditions described in the Table below.
PCR cycling conditions (second round of PCR)
| A | B | C | D | |
| STEP | TEMP | TIME | ||
| Initial Denaturation | 98 C | 30 sec | ||
| 15 cycles | Denaturation | 98 C | 10 sec | |
| Annealing | 66 C | 30 sec | ||
| Extension | 72 C | 30 sec | ||
| Final Extension | 72 C | 2 min | ||
| Hold | 6 C | ∞ |
PCR cycling conditions have been optimized with an Applied Biosystems QuantStudio 3.
Number of cycles can be reduced to 10.
PCR visualization
Conduct in Post-PCR.
After the second PCR reaction, pool the replicated samples and inspect the PCR products using Agarose gel electrophoresis (1%).
Successful amplification should give bands at 500 bp (See picture below).
Note that some primer dimers ~150 bp are expected, and is removed in the purification step.
Purification
Conduct in post-PCR.
Perform purification with magnetic beads (Cytiva Sera-Mag Select) using 0.75x beads concentration following the manufacturer recommendations.
Quantify and pool in equimolar concentration
Conduct in post-PCR:
Quantify and pool the samples to create a final library using PicoGreen, if pooling samples from different marker lengths, see Sheet 3.
Make sure to reach the requirements for illumina sequencing, with at least 20 μL of 10 nM library pool (Sheet 3).
Read more about the requirements here:
Conduct in post-PCR.
Calculation conducted in the Google Sheets.
Pool the PCR samples in equal DNA amount (ng) or for unequal length amplicons, in equal molecule amount (mol).
You will get one tube with a mix of all the samples in it.
To calculate the volume of each sample to be pooled (DNA amount mixing):
Use the lowest concentration sample to define the minimum amount of DNA (ng) that you have available from a single sample: the DNA concentration (ng/μL) of the lowest concentration sample multiplied with its volume (μL). This will be your target DNA amount for each sample.
Calculate how many μLs of each sample you need to achieve the target DNA amount: divide the target DNA amount with the concentration of each sample.
Pipette into one tube the calculated volume of each sample. Aim to use the same pipette for all samples (dilute or pipette multiple times) to avoid pipette calibration errors.
Quality control
Ensure that primer dimers have been removed from the final pool by using Agarose gel electrophoresis (1%). If primer dimers are still present, conduct a new bead purification as described above, or conduct gel extraction.
If the DNA concentration is too low, <10nM, you can either conduct another bead purification and elute with a lower elution volume (recommended).
Or use a SpeedVac to further up-concentrate the DNA (Remember that SpeedVac also up-concentrates salts, which may be unfavorable for sequencing.