Mar 25, 2025
High-throughput individual insect metabarcoding for identification and interaction data
- 1Newcastle University
- Foraging Ecology Research Group
- Network Ecology Group
Protocol Citation: Jordan P Cuff, Thomas Howells, James JN Kitson, Ben SJ Hawthorne, darren.evans 2025. High-throughput individual insect metabarcoding for identification and interaction data. protocols.io https://dx.doi.org/10.17504/protocols.io.n2bvj3qrnlk5/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: November 06, 2023
Last Modified: March 25, 2025
Protocol Integer ID: 90463
Keywords: metabarcoding, biomonitoring, entomology, high-throughput sequencing, community ecology, field techniques
Funders Acknowledgements:
European Union Horizon 2020
Grant ID: 773554 (EcoStack)
Abstract
This protocol is designed for extracting DNA from individual invertebrates for metabarcoding to detect and identify ecosystem service and disservice providers and their interactions. The reagents and methods proposed offer a cost effective and high-throughput method for molecular analyses of individual invertebrates using standard lab equipment. Where specialist equipment us used, attempts are made to suggest low-cost alternatives.
Image Attribution
Created in BioRender. Cuff, J. (2025) https://BioRender.com/559f6hh
Materials
For field collection and initial storage:
- Small collection pots or tubes for storage of samples
- 100 % ethanol for preservation
- Chemgene/diluted bleach for sterilisation of equipment
For DNA extraction:
- Hardened carbon steel ball bearings
- 2.2 mL deep well plates for initial lysis
- Deep-well and standard Kingfisher 96-well plates
- Plate seals for long-term storage
For DNA amplification and subsequent steps:
- Tagged PCR primers with bridge primer sequence for Nextera library preparation by PCR
- 2X hot-start Taq polymerase mastermix
- Molecular grade water
- 96-well PCR plates
- Mineral oil
- 0.1X and 1X SPRI beads
- Nextera Illumina adapter index primers
Buffers and reagents:
- Sodium chloride
- Tris-HCl
- EDTA
- GITC
- Nuclease-free water
- SDS
- PEG
- Tris-HCl
- 100 % ethanol
- Papain
Equipment:
- -20 °C freezer
- Geno/Grinder 2010 or similar bead beater for homogenisation
- Thermocycler
- Magnetic stand (for plates and tubes)
- Centrifuge
- Microcentrifuge
- Vortex
- Pipettes (preferably including multichannel, ideally including 96-well)
- Ideally, Kingfisher Apex or similar
- Illumina sequencer
Safety warnings
Check safety guidelines for individual reagents before commencing work. Some reagents will be toxic, corrosive or otherwise present health and safety risks. Appropriate personal protective equipment should be used at all times, not only for personal safety but also reduction of contamination risk.
Ethics statement
Check national and institutional policies for insect research. Follow best practice guidelines and stay up to date with the latest developments in insect welfare (e.g., through the Insect Welfare Research Society). Only kill as many invertebrates as is necessary, and always do so as humanely as possible.
Sampling of invertebrates
Sampling of invertebrates
3d 0h 30m
3d 0h 30m
Select suitable sites and locations for insect sampling. This protocol is based on hand-collected foliar and flower visitor invertebrates, but other collection methods are viable; be wary of cross-contamination and surface DNA though, and consider surface sterilisation accordingly. Consider how systematic the study needs to be and the various constraints imposed on the data by the study design.
Following collection, store insects in sterile tubes. For flower visitors, store insects dry and consider killing them with ethyl acetate on a cotton ball; for other insects, storage in 100 % ethanol may be ideal. Kill invertebrates as humanely as possible. Freezing is widely considered to be among the optimal methods.
3d
Transfer individual invertebrates into 96-well plates (ideally deep-well, e.g., 2.2 mL, with fixable lids for grinding/washing). Consider the distribution of experimental controls ahead of subsequent steps to streamline downstream liquid handling.
Our recommended PCR plate layout, which could be adopted here for streamlining downstream. Created in BioRender. Cuff, J. (2025) https://BioRender.com/559f6hh
Store samples at -20 °C until ready to process.
15m
Preparation and homogenisation of samples
Preparation and homogenisation of samples
16h 8m
16h 8m
The DNA extraction protocol is largely adapted from the BOMB-Bio tissue nucleic acid extraction protocol. See their documentation for additional detail.
CITATION
Add 100 µL TNES lysis buffer to each well of a 96-well plate.
Note
For the TNES buffer, follow the recipe provided by BOMB-Bio (100 millimolar (mM) Tris-HCl, 52 millimolar (mM) NaCl, 10 millimolar (mM) EDTA, 10 Mass / % volume SDS).
5m
Two protocols are presented below, which simply differ in whether samples are ground or surface-washed. For herbivorous insects, we recommend grinding to facilitate analysis of internal DNA (either consumed plants or parasitoids). For flower visitors, we recommend surface washing (exactly the same protocol, but without ball bearings during the grinding step). This still facilitates analysis of DNA from the insect itself, but also pollen on the surface of the insect. These protocols could both be used for a wider range of potential applications.
Herbivorous insects1 step
Add one 3 mm hardened carbon steel bead to each well.
Note
Beads are usually shipped coated in manufacturing oil (especially the carbon steel beads). To remove this, place beads in a borosilicate glass beaker or Duran bottle with plastic pouring lip and lid removed then bake for at least 12 hours at 250 °C.
5m
Grind the samples in a tissue grinder/homogeniser/lyser at 1750 RPM for 1 minute.
1m
Add 10 µL 20 mg/mL papain to each well.
Note
For streamlining, you could add the papain prior to grinding, but be aware that this may reduce its efficacy.
2m
Incubate overnight (~16:00:00 ) at 37 °C .
16h
DNA extraction
DNA extraction
1h 5m
1h 5m
Centrifuge the plate at 2000 x g, Room temperature, 00:02:00 .
2m
Prepare Kingfisher Apex reagent plates as below.
Note
The Kingfisher Apex is ideal for automated high-throughput extraction of DNA, but will not always be available. Alternative equipment can achieve similar results, including just using magnetic racks. In that case, rather than transferring beads between plates, remove the supernatant and replace it with the next reagent, as described for normalisation below.
For the sample plate, to each well of a 96-well deep-well plate, add 60 µL of the lysate from STEP 11, 120 µL of 1.5X GITC buffer, 120 µL of 1 mg/mL SeraMag Speed Beads in TE (10 millimolar (mM) Tris and 1 millimolar (mM) EDTA) and 240 µL isopropanol.
Note
For the 1.5X GITC buffer, follow the recipe provided by BOMB-Bio (6 Molarity (M) GITC, 75 millimolar (mM) Tris-HCl, 3 % volume sarkosyl, 30 millimolar (mM) EDTA, 0.15 % volume antifoam).
5m
For the two ethanol plates, to each well of two 96-well deep-well plates, add 400 µL 80 % ethanol.
2m
For the isopropanol plate, to each well of a 96-well deep-well plate, add 400 µL isopropanol.
1m
For the elution plate, to each well of a 96-well standard plate, add 100 µL molecular biology grade water.
1m
Insert plates into the Kingfisher Apex and run a preset programme with the below steps.
30m
Pick up the 96 deep-well tip comb from a 96-well standard plate.
30s
Bind DNA to the beads by mixing at medium speed for 00:05:00 with a slow post-mix for 00:05:00 .
10m
Collect the beads in three 00:00:01 collections.
10s
Release the beads into the isopropanol plate and mix for 00:01:00 at medium speed, and collect the beads in three 00:00:01 collections.
1m 30s
Release the beads into one of the 80 % ethanol plates and mix for 00:01:00 at medium speed, and collect the beads in three 00:00:01 collections.
1m 30s
Release the beads into one of the 80 % ethanol plates and mix for 00:01:00 at medium speed, and collect the beads in three 00:00:01 collections.
1m 30s
Dry the beads above the well for 00:02:00 .
2m
Release the beads into the elution plate and mix for 00:02:00 at fast speed with a slow post-mix for 00:03:00 , and collect the beads in four 00:00:20 collections.
6m
Leave the tip comb in an empty 96-well standard plate.
30s
Store the eluted DNA at -20 °C until ready for subsequent steps.
PCR
PCR
3h 22m
3h 22m
Decide how samples will be distributed across plates (but don't distribute the DNA yet). Consider including a negative control in each row and column to detect any contaminants in each tagged forward and reverse primer. Among these wells, include any DNA extraction negative controls. Include positive controls (ideally mixed samples of species not found in the same study system), perhaps one adjacent to negative controls and the other adjacent only to samples (but both on separate rows and columns). Include blank controls (ideally wells into which no reagents or at least no primers are added), perhaps one adjacent to negative controls and the other adjacent only to samples (but both on separate rows and columns).
If using multiple PCR primer pairs, familiarise yourself with the annealing temperatures for each and prepare separate PCR plates for each. For optimal accuracy, consider running replicates of each reaction (e.g., triplicates).
Our recommended PCR plate layout, which could be adopted here for streamlining downstream. Created in BioRender. Cuff, J. (2025) https://BioRender.com/559f6hh
10m
Prepare enough PCR mastermix for each sample.
For a full plate, the below values will usually suffice (with some overage to account for pipetting error), but check your specific Taq polymerase mix for any differences:
| A | B | |
| Reagent | Volume (μL) | |
| Molecular grade water (DNase free) | 422.4 | |
| 2X hot-start PCR mastermix | 528 |
Note
These values are for 10 µL reaction volumes, which have been demonstrated to be effective for community metabarcoding. Consider running them in triplicate for more accurate results.
2m
For ease, if able to use a 96-well pipette, consider creating a "primer plate" containing both PCR primers for each well at 5 µM concentration; this is especially effective when using multiple plates. For 10 µL reaction volumes, this will subsequently involve adding 7.75 µL hot-start Taq polymerase and water mastermix (described in the step above) to each well, followed by 1.25 µL of each primer mix to its corresponding well. It is possible to use this strategy with a multichannel pipette, but the next suggestion may be easier.
If not using a 96-well pipette, consider making eight mastermixes each with sufficient reagents for 12 samples (7.75 µL mastermix per well, so 97.65 µL for 12 with some overage for pipetting error), each containing a different forward primer (with 0.625 µL of 5 micromolar (µM) forward primer per well, so 7.875 µL per mastermix for 12 with some overage for pipetting error). Distribute 8.375 µL of forward primer + mastermix to each well across the rows corresponding to each forward primer. Then distribute 0.625 µL of reverse primer to each well in the corresponding columns making sure to change pipette tip for each sample to avoid cross-contaminating the forward primers.
Distribution of tagged PCR primers across the 96-well plate. Created in BioRender. Cuff, J. (2025) https://BioRender.com/559f6hh
15m
Add 1 µL DNA to each corresponding sample or positive control well, and 1 µL molecular grade water to each negative control other than extraction negative control(s).
10m
Distribute one drop of mineral oil into each well of the PCR plate(s) (~20 µL ).
Note
This can be achieved by taking a large volume of mineral oil into the pipette tip and then gently depressing the plunger so that a drop forms and falls from the tip into each well.
Mineral oil improves sealing of reactions by preventing evaporation and condensation. By reducing evaporation and thus loss of product, this also reduces potential cross-contamination.
2m
Briefly centrifuge the plate to ensure that the oil is above the PCR mix and everything is at the bottom of each well without air bubbles.
1m
Load the PCR plate into a thermocycler. Ensure that the temperature regime matches the enzyme used (including any heat activation for hot-start Taq) and that the annealing temperature matches the PCR primers used.
Note
Given differences between labs and samples, and inaccuracies in temperature calibration, considering running a temperature gradient PCR with known samples to check the specificity of your PCR primers.
2m
Run your PCR programme.
2h
The samples should now be checked for successful amplification, contamination in negatives and any secondary banding. Gel electrophoresis will achieve this, but digital systems like the Qiagen Qiaxcel will do this and facilitate equalisation by generating amplicon-specific DNA concentations.
40m
Normalisation
Normalisation
1h 55m
1h 55m
Note
Equalisation is more effective than normalisation, but requires amplicon concentration data. Both can be time and labour intensive though, so can be skipped if time restricted for large-scale projects, although at the expense of data recovery and evenness.
If the PCRs were replicated (i.e., each sample run multiple times for each PCR primer pair used), these can be merged together into one plate at this point, or carried forward separately. Keeping the replicates separate increases the number of libraries to prepare and sequence later, but better facilitates inconsistencies between samples. To merge triplicates, assuming use of 10 µL reaction volumes, pipette 8 µL from each well of two of the three plates into the corresponding well of the third. Briefly centrifuge the merged plate to move the oil to the top of the product again.
Note
To avoid pipetting oil from the oil-sealed PCR products, plunge the pipette to the first stop and fully insert the pipette tip into the bottom of the well, then release sharply. The PCR product will be taken up quickly, whereas the relatively viscous oil will be taken up slowly, thus being outcompeted by the PCR product.
5m
Prepare 0.1X SPRI bead solution and bring to room temperature. The below steps detail how to make this solution, but it is also commercially available.
20m
If using beads such as Sera-Mag Magnetic SpeedBeads (carboxylated, 1 µm, 3 EDAC/PA5), take 1 mL of well-mixed bead solution and wash the beads twice with TE+Tween buffer (10 millimolar (mM) Tris base, 1 millimolar (mM) EDTA, 0.05 % volume Tween 20, pH 8.0) by magnetising the beads, removing the supernatant, adding the TE+Tween, remagnetising the beads and removing the supernatant, and repeating the addition and removal of TE+Tween once more.
5m
To the beads, add the following mix:
| A | B | |
| Reagent | Volume | |
| 5 M NaCl | 25 mL | |
| Molecular grade water | 3.582 mL | |
| 1 N HCl | 0.168 mL | |
| 1 M Tris base | 0.5 mL | |
| 0.1 M disodium EDTA | 0.5 mL |
5m
Add 20 mL of 50 % volume PEG to the tube to reach a 1X bead solution (alongside making the 0.1X solution, this will be useful later).
2m
Add 5 mL of 1X bead solution to 45 mL of the following mix to make a 0.1X solution:
| A | B | |
| Reagent | Volume | |
| 5 M NaCl | 25 mL | |
| Molecular grade water | 3.582 mL | |
| 1 N HCl | 0.168 mL | |
| 1 M Tris base | 0.5 mL | |
| 0.1 M disodium EDTA | 0.5 mL | |
| 50 % PEG | 20 mL |
5m
If ≥20 µL of PCR product is available in each well, pipette 20 µL of 0.1X SPRI bead solution into each well of a new 96-well plate. If less DNA is available, add the available volume of 0.1X SPRI beads and add the same volume of PCR product to these beads in the next step.
Note
When working with magnetic beads, ensure they are fully mixed with no residue at the bottom of the container.
2m
Add 20 µL of PCR product (or whatever volume of beads was used in the last step) to each corresponding well of 0.1X beads, avoiding oil, and mix by vortexing (1500 rpm, Room temperature , 00:01:00 ).
Note
To avoid pipetting oil from the oil-sealed PCR products, plunge the pipette to the first stop and fully insert the pipette tip into the bottom of the well, then release sharply. The PCR product will be taken up quickly, whereas the relatively viscous oil will be taken up slowly, thus being outcompeted by the PCR product.
2m
Incubate at Room temperature for 00:05:00 .
5m
Place on a magnetic stand for 00:05:00 .
5m
Remove all but 5 µL of the mixture from each well via pipette without disturbing the beads, which should be settled on the magnet (although can be hard to see with 0.1X solutions).
5m
Add 200 µL 80 % volume ethanol to each well.
2m
Remove the ethanol and add a further 200 µL 80 % volume ethanol to each well.
4m
Remove the ethanol as completely as possible by pipetting.
Note
A small amount of remaining ethanol isn't debilitating in this case, unlike usual magnetic bead cleaning protocols, because we will complete a subsequent purification immediately after.
4m
Add 10 µL molecular grade water to each well, shake at 1500 rpm, Room temperature , 00:01:00 and incubate at Room temperature for 00:05:00 .
7m
Place on a magnetic stand for 00:05:00 .
5m
Pool 8 µL from each well into a single tube, leaving magnetic beads behind in the 96-well plate.
Note
To pool samples using a multi-channel pipette, pipette each column of samples into a new strip of 200 µL tubes, then merge all of these tubes into a single 1.5 mL tube at the end.
4m
Mix the pooled samples by flicking and inverting, and transfer 700 µL into a new 2.5 mL microcentrifuge tube.
1m
Add 700 µL 1X magnetic beads for DNA purification (see note below for size selection) and mix by vortexing. Incubate for 00:05:00 .
Note
This is a good opportunity for size selection as well, especially if the PCR product contains any secondary bands. If using SPRI beads, adjust the volume added to select different fragment sizes.
6m
Place on a magnetic stand for 00:05:00 .
5m
Add 400 µL 80 % volume ethanol.
1m
Remove the ethanol and add a further 400 µL 80 % volume ethanol.
1m
Remove the ethanol completely. Briefly centrifuge the sample to collect and remove any residual ethanol. Allow to air-dry until the aggregation of magnetic beads transitions from 'glossy' (shiny reflection of light) to 'matte' (dull dark brown mass), but not so long that it dries completely (i.e., begins to turn a rusty red and shows cracks).
2m
Add 100 µL molecular grade water and mix by vortexing. Incubate for 00:05:00 at Room temperature .
6m
Place the sample on a magnetic stand for 00:05:00 .
5m
Remove 95 µL of the water and place it in a new tube. This will be the library used in the subsequent section.
1m
Library preparation and sequencing
Library preparation and sequencing
3h 9m
3h 9m
Quantify the concentration of the library from the previous step (e.g., using Qubit dsDNA assay).
5m
Dilute library to ≤ 5 ng/µL in molecular grade water.
Note
Without diluting the library, there is a risk that the adapters will not be added to all DNA (i.e., there is too much DNA) which will compromise the sequencing yield.
2m
Add a drop of mineral oil to one PCR tube for each library, alongside an additional tube for a negative control.
1m
Per library, assemble the following reaction:
| A | B | |
| Reagent | Volume | |
| 2X hot-start PCR mastermix | 7.5 | |
| Molecular grade water | 1 | |
| Nextera sequencing adapter primer mix | 1.5 | |
| DNA library | 5 |
5m
Briefly centrifuge the tubes to ensure that the oil is above the PCR mix.
1m
Load the PCR reactions into a thermocycler. Ensure that the temperature regime matches the enzyme used (including any heat activation for hot-start Taq) and that the annealing temperature matches the PCR primers used.
5m
Run your PCR programme for 12-20 cycles, with 12 for ~5 ng/µL libraries and more for lower concentrations.
2h
Quantify the concentration of the library from the previous step (e.g., using Qubit dsDNA assay).
5m
Check that the adapters have been added to the DNA by determining and comparing the amplicon sizes for libraries before and after this second PCR. Amplicons should be longer after the second PCR. This is ideally assessed using a digital system like TapeStation, but even gel electrophoresis will work.
40m
Once successful adapter addition is confirmed, libraries can be pooled so that each is equimolar in the final mixture based on the concentrations determined above (e.g., by Qubit).
Note
This can be achieved by dividing the maximum concentration across the libraries by each library concentration and pooling that many µL from each. These values may need to be multiplied up to ensure adequate yield for sequencing.
Be careful if using multiple PCR primer pairs! Different amplicons have different molecular weights, so pool based on fmol rather than ng/µL.
5m
Check with your sequencing provider how many fmol they will need in how many µL. This will be based on the sequencer, sequencing cartridge and any QC processes they follow. The libraries should be ready for sequencing.
Protocol references
CITATION
Citations
Oberacker P, Stepper P, Bond DM, Höhn S, Focken J, Meyer V, Schelle L, Sugrue VJ, Jeunen GJ, Moser T, Hore SR, von Meyenn F, Hipp K, Hore TA, Jurkowski TP. Bio-On-Magnetic-Beads (BOMB): Open platform for high-throughput nucleic acid extraction and manipulation.
https://doi.org/10.1371/journal.pbio.3000107Step 4
Oberacker P, Stepper P, Bond DM, Höhn S, Focken J, Meyer V, Schelle L, Sugrue VJ, Jeunen GJ, Moser T, Hore SR, von Meyenn F, Hipp K, Hore TA, Jurkowski TP. Bio-On-Magnetic-Beads (BOMB): Open platform for high-throughput nucleic acid extraction and manipulation.
https://doi.org/10.1371/journal.pbio.3000107Acknowledgements
This protocol was developed in and for Newcastle University's School of Natural and Environmental Sciences Molecular Diagnostics Facility. JPC and DME were funded by European Union Horizon 2020 Grant Award Number 773554 (EcoStack).