Jun 29, 2026
OceanOmics Centre: eDNA Amplicon Clean-Up and Pooling (Manual)
- Laura Missen1,
- Marcelle Ayad2,
- Ebony Thorpe1,
- Georgia Nester1,
- Anna Depiazzi1,
- OceanOmics UWA1
- 1Minderoo OceanOmics Centre at UWA;
- 2Minderoo Foundation
External link: https://www.uwa.edu.au/oceans-institute/partnerships/minderoo-oceanomics-centre-at-uwa
Protocol Citation: Laura Missen, Marcelle Ayad, Ebony Thorpe, Georgia Nester, Anna Depiazzi, OceanOmics UWA 2026. OceanOmics Centre: eDNA Amplicon Clean-Up and Pooling (Manual). protocols.io https://dx.doi.org/10.17504/protocols.io.bp2l6okqzlqe/v1
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: June 03, 2026
Last Modified: June 29, 2026
Protocol Integer ID: 318472
Keywords: eDNA, environmental DNA, OceanOmics, Marine vertebrate, metabarcoding, 12S, 16S, marine eDNA, normalisation, minipooling, COI, amplicon, pooling sample amplicon, oceanomics centre for marine vertebrate edna metabarcoding, follow oceanomics centre, sample amplicon, oceanomics centre, sample pool, marine vertebrate edna metabarcoding, oceanomic, control pool, amplicon clean, minipool, control amplicon, coi metabarcoding of marine vertebrate, minipools from the same project, resulting masterpool, amplicon, pooling, following qpcr, pooling workflow, masterpool, qpcr triplicate, only pool, marine vertebrate
Abstract
This protocol describes the post-qPCR amplicon clean-up and pooling workflow used by the OceanOmics Centre for marine vertebrate eDNA metabarcoding assays. It applies to five assays: 16S-FishD, MiFish-U, MiFish-E2, MarVer1, and COI-Leray.
Following qPCR, amplicons are first combined into minipools. A minipool is generated for each plate by pooling sample amplicons and control amplicons separately. Each plate therefore produces two minipools: one sample pool and one control pool. Amplicons are pooled proportionally according to their average End Point Fluorescence (EPF) calculated from qPCR triplicates. For each assay, additional control-only pools are generated for non-template controls (NTCs) and internal positive controls (ITCs).
Each minipool undergoes a 1.8× AMPure XP bead clean-up and quality control assessment. Following QC, minipools from the same project and assay are combined into masterpools. The resulting masterpools are used as input for library preparation and next-generation sequencing.
An automated version of this protocol is also available and requires a Biomek i7 Workstation. Follow OceanOmics Centre: eDNA Amplicon Clean-Up and Pooling (Automated) if you wish to use this method.
The following protocol is a pre-requisite: OceanOmics: 12S, 16S, and COI Metabarcoding of Marine Vertebrates.
Guidelines
Standard lab set-up:
- Ensure competency and training requirements are met.
- Ensure the laboratory is appropriately configured with separation of pre- and post- PCR rooms. Amplified PCR products should never come in contact with equipment used for non-amplified DNA.
- Wear gloves and a lab coat.
- Use tips with filters to avoid contamination.
- Wipe workspace with 10% bleach, leave to decontaminate for 10 minutes before wiping with ddH20.
- Sterilise all equipment (vortex, minispin, pen markers, outside of tip boxes, pipettes) before use.
- UV pipettes, racks, tubes (lids open) for 15 minutes before use.
- Ensure all reagents are aliquoted in appropriate amounts, and stored according to manufacturers' recommendations. Never pipette directly from reagent stocks. Bring all reagents to room temperature before use.
- All waste must be disposed of following institutional biosafety regulations.
Materials
Equipment
- Qubit 4.0 Fluorometer
- UV sterilisation cabinet
- Dyna-Mag-2 magnetic rack (ThermoFisher)
Equipment
4150
NAME
TapeStation System
TYPE
Agilent
BRAND
G2992AA
SKU
LINK
Consumables
- Eppendorf DNA LoBind 1.5mL tubes
- 50mL Falcon tubes
- Nuclease-free water
Reagents
AMPure XP Reagent, 450 mLBeckman CoulterCatalog #A63882
100% Molecular grade ethanol
Resuspension BufferIllumina, Inc.Catalog #FC-131-1096
Qubit dsDNA HS assay kitFisher ScientificCatalog #Q32854
D1000 ScreenTapeAgilent TechnologiesCatalog #5067-5582
D1000 ReagentsAgilent TechnologiesCatalog #5067-5583
D1000 LadderAgilent TechnologiesCatalog #5067-5586
Safety warnings
This protocol involves the use of potentially hazardous biological materials, chemicals (including ethanol and enzymatic reagents), and electrical laboratory equipment. Always wear appropriate personal protective equipment (PPE), including a lab coat, gloves, and eye protection. Be familiar with the relevant Safety Data Sheets (SDS) and laboratory risk assessments before commencing work. Follow institutional safety guidelines and dispose of waste in accordance with local regulations. Exercise caution when handling sharp instruments, thermal cyclers, and high-speed centrifuges to prevent injury or equipment damage.
Before start
1. Remove AMPure XP beads from the fridge and allow to equilibrate at room temperature for 30 minutes.
2. Remove Resuspension Buffer (RSB) (Illumina) from the -20°C freezer and allow to equilibrate at room temperature for 30 minutes.
3. Bring amplicon sample pools to room temperature, then vortex and spin-down.
4. Prepare 2 x 50mL falcon tubes. Label one as 'WASTE', to decant liquid waste, and the other '80% EtOH' to prepare an 80% ethanol dilution.
5. Dilute molecular grade ethanol to 80% using nuclease-free water, 3 mL is required per pool.
6. Prepare 2 x 1.5mL DNA LoBind tubes labelled CLEANMINIP with project, assay, type (sample/control), plate #, date, and initials. Repeat for all minipools.
7. Preheat the dry block heater to 37°C.
Bead Clean-Up Minipools
This protocol follows previous metabarcoding steps described in OceanOmics Centre: 12S, 16S and COI Metabarcoding of Marine Vertebrates.
Aliquot up to 200 µL of sample minipool into a new labelled 1.5mL DNA LoBind tube.
NOTE: volumes for each minipool can be found in the 'minipool_summary.csv' output file from the
Aliquot up to 100 µL of control minipool into a new labelled 1.5mL DNA LoBind tube.
Vortex AMPure XP beads for 30 seconds until fully resuspended.
Pipette 1.8X volume of AMPure XP beads into each minipool tube.
For example, if sample minipool volume is 200 µL, add 360 µL (1.8X) of AMPure XP beads. If control minipool volume is 100 µL, add 180 µL (1.8X) of AMPure XP beads.
Pipette to mix minipool containing beads 10x until homogenised, changing tips each time.
Incubate minipool tubes containing AMPure XP beads on the benchtop at room temperature for 5 minutes.
Place tubes into the magnetic rack and incubate on the benchtop at room temperature for 5 minutes. The supernatant should become clear as the beads pellet to the magnet.
Using a pipette, aspirate the supernatant without disturbing the bead pellet. Discard the supernatant into the designated waste tube. Use a new pipette tip between each sample.
With the tubes on the magnetic rack, pipette 1000 µL of 80% ethanol to each tube.
Incubate tubes on the magnetic rack at room temperature for 1 minute.
Using a 1000 µL pipette, aspirate the supernatant without disturbing the bead pellet. Discard the supernatant into the designated waste tube. Use a new pipette tip between each sample.
Repeat steps 10-12 for a total of 2 ethanol washes.
Spin-down tubes then place back on the magnetic rack.
Using a 20 µL pipette, remove any residual liquid. Use a new pipette tip for each sample.
With the tubes on the magnetic rack, open each tube lid and allow to air dry at room temperature for 5 minutes.
Remove the tubes from the magnetic rack.
Pipette 52 µL RSB to each tube directly onto the beads. Use a new pipette tip between each sample.
Vortex for 30 seconds until fully resuspended.
Incubate in a dry block heater at 37°C for 5 minutes.
After incubation, briefly spin-down then place tubes into the magnetic rack and incubate at room temperature for 5 minutes. The supernatant should become clear.
Using a 100 µL pipette, transfer 50 µL of the supernatant into the corresponding labelled 1.5 mL DNA LoBind tube labelled Clean MINP (minipool). Do no disturb the bead pellet.
Safe stopping point.
Clean MINIP can be stored at 4°C for short-term storage (if processing the same week), or -20°C for long-term storage. Avoid freeze/thaw cycles as this can degrade DNA.
QC Clean Minipool
Prepare and label a new 1.5mL DNA LoBind tube for each minipool.
Make a 1:10 dilution of all sample minipools by combining 9 µL nuclease-free water and 1 µL sample into the new labelled 1.5mL DNA LoBind tube.
NOTE: no dilution is required for control minipools.
Quantify 2 µL of all control and diluted pools using Qubit dsDNA High Sensitivity kit.
Run all minipools on a Tapestation (Agilent) using a D1000 Screen Tape and D1000 Reagents to check amplicon size in sample pools, and to check for contamination in control pools.
NOTE: the minipools may have to be diluted for this step. Refer to the D1000 assay guidelines: 
D1000_QuickGuide.pdf301.9KB
D1000_QuickGuide.pdf301.9KB The expected fragment sizes for each assay are shown below:
Expected result
This is an example of an expected result for 16S-Fish/D assay:
Prepare Masterpools
In the MasterPool_Calculation_Workbook, enter the Qubit concentrations of each clean minipool, and the number of samples per plate.
MasterPool_Calculation_Workbook.xlsx28.2KB NOTE: the number of samples can be found in the 'minipool_summary.csv' output file obtained from the R-Script: plates_to_pooling
The total required yield (ng) may be modified, but should be consistent for all pools. If the 'volume to add to masterpool' cell is highlighted red, reduce the required yield (ng) for all pools.
Prepare a new 1.5mL DNA LoBind tube labelled as MASTP (masterpool).
Following the volumes calculated in the workbook, pipette the required volume of each pool into the 1.5mL MASTP tube.
Quantity concentration of masterpools using HS dsDNA Qubit
- Make a 1:10 dilution of all 'Sample' pools by combining 9 µL nuclease-free water and 1 µL sample.
- No dilution is required for 'Control' pools.
- Quantify all pools using Qubit dsDNA HS kit.
Enter concentration values into the Quantification Table of the workbook.
Safe stopping point.
MASTP can be stored at 4°C for short-term storage (if processing the same week), or -20°C for long-term storage. Avoid freeze/thaw cycles as this can degrade DNA.
Blend Sample and Control MASTP for input into library preparation using the 'Blending Table' in the workbook. The total volume depends on the library preparation and the assay.
- For Illumina TruSeq DNA PCR-Free Library Preparation, a total volume of 50 µL is required.
- For Lucigen NxSeq AMPFree Low DNA Library Preparation, a total volume of 17 µL is required.
- For 16S-Fish/D, MiFish-U/E2 and MarVer1, a 250ng input is required for library preparation.
- For COI-Leray, a 100ng input is required for library preparation.
Safe stopping point.
Blended MASTP can be stored at 4°C overnight or at -20°C long-term.
Proceed to library preparation:
- OceanOmics: eDNA Amplicon Library Preparation with Illumina TruSeq DNA PCR-Free Kit
- OceanOmics: eDNA Amplicon Library Preparation with Lucigen NxSeq AMPFree Low DNA Kit
Protocol references
MiFish-U:
Miya, M., Sato, Y., Fukunaga, T., Sado, T., Poulsen, J., Sato, K., Minamoto, T., Yamamoto, S., Yamanaka, H., & Araki, H. (2015). MiFish, a set of universal PCR primers for metabarcoding environmental DNA from fishes: detection of more than 230 subtropical marine species. Royal Society open science, 2(7), 150088.
MiFish-E2:
Miya, M. a. S., T. (2019). Multiple species detection using MiFish primers. Pages 55–92 in
Environmental DNA sampling and experimental manual version 2.1. Ed. by eDNA Methods
Standardization Committee, The eDNA Society, Otsu, Japan.
MarVer1:
Valsecchi, E., Bylemans, J., Goodman, S. J., Lombardi, R., Carr, I., Castellano, L., Galimberti, A., & Galli, P. (2020). Novel universal primers for metabarcoding environmental DNA surveys of marine mammals and other marine vertebrates. Environmental DNA, 2(4), 460-476.
16S-Fish/D:
Berry, T. E., Osterrieder, S. K., Murray, D. C., Coghlan, M. L., Richardson, A. J., Grealy, A. K., Stat, M., Bejder, L., & Bunce, M. (2017). DNA metabarcoding for diet analysis and biodiversity: A case study using the endangered Australian sea lion (Neophoca cinerea). Ecology and evolution, 7(14), 5435-5453.
Deagle, B. E., Gales, N. J., Evans, K., Jarman, S. N., Robinson, S., Trebilco, R., & Hindell, M. A. (2007). Studying seabird diet through genetic analysis of faeces: a case study on macaroni penguins (Eudyptes chrysolophus). PloS one, 2(9), e831.
COI-Leray:
Leray, M., J. Y. Yang, C. P. Meyer, et al. 2013. A New Versatile Primer Set Targeting a Short Fragment of the Mitochondrial COI Region for Metabarcoding Metazoan Diversity: Application for Characterizing Coral Reef Fish Gut Contents. Frontiers in Zoology 10: 34.