TITLE: Quantitative multi-target amplicon sequencing workflow
AUTHORS: Mateusz Buczek, Monika Prus-Frankowska, Piotr Łukasik

[DESCRIPTION]

High-throughput sequencing of marker gene amplicons is a powerful tool for the discovery of biodiversity, at different levels. However, the method has multiple caveats.
Here, we describe in detail a versatile laboratory protocol for high-throughput and cost-effective characterization of microbiota across large numbers of wild insects. By simultaneously targeting different marker regions, in a quantitative manner,  the protocol enables the determination of insect host identity and reconstruction of bacterial diversity, alongside bacterial quantification combined with rigorous contamination control. It can be easily extended to encompass other marker regions and customized depending on specific needs, research questions, or specimen properties. 
The protocols encompasses DNA extraction from individual specimens (specimen homogenization, homogenate purification) and Illumina amplicon library preparation for two target regions: insect mitochondrial cytochrome oxidase I gene and the V4 region of bacterial 16S rRNA gene. It also explores the sequencing options and briefly outlines bioinformatics solutions that are published elsewhere.
The protocol is labor- and cost-effective and relies on readily available reagents and equipment. It is highly customizable: we suggest alternative solutions, including omission of certain optional elements, or addition of alternative reagents and approaches.
To date, we have successfully applied the protocol to nearly 20,000 individual insects, obtaining insights into microbial distribution, diversity, and transmission across multiple populations or large multi-species community subsamples of wild insects.

[STEPS]
SECTION: A. DNA extraction
3. Sample homogenization

Material: 
 • Lysis buffer „Vesterinen” [0.4M NaCl, 10mM Tris-HCl, 2mM EDTA, 2% SDS] – before use, place
in a container with warm water to help dissolve any precipitate
 • Proteinase K (stored frozen at -20oC) 
 • Ceramic beads: 2,8mm and 0,5mm 
 • 2 ml screw-cap tubes
 • 50 ml Falcon tubes, tweezers, Petri dish, paper towels, metal spatula, torch

Equipment:
• Homogenizer. We use Bead Ruptor Elite (Omni International), with a block suitable for 24 2ml screw cap tubes. A different homogenizer may require different tubes. 
Make sure that the tubes are made for plastic resistant for bead beating!

Homogenization:

1.      Prepare 91 tubes and sign them with sample ID on the side (the inscriptions on the lid may wear off). For each plate you will have 88 insects and 3 negative controls - "extraction blanks" (3C, 6C, 9C). 

2.      For standard size insects1) (size 1-2) in 50 ml falcon tube prepare a mix of a "Vesterinen" lysis buffer and proteinase K for 100 samples. For one sample in that case we use 195 ul of lysis buffer and 5 ul of proteinase K, so for a 100 – 19 500 ul of buffer and 500 ul proteinase K. However, if you see that the insects are bigger prepare more buffer, because you will use more than 200 ul of buffer mix per sample. 

3.      Open the tubes and place them on a 96 well rack. Make sure that the caps are placed on aluminium foil to avoid contamination from the bench. To each tube add 200 ul of prepared buffer mix. Then into each tube add one small spoon of 0.5mmm beads (make sure to sterilize the spoon under the flame first!) and one big bead 2.8mm. You can add big beads by hand, but before change gloves to minimalize the risk of contamination. 
Note that bigger insects will need more buffer mix and more big beads (2-3 instead of 1).

4.      Close the tubes with blanks – 3C, 6C, 9C. 

5.      Transfer your specimens to the tubes. Sterilize the tweezers at least between the different eppendorfs with insects to homogenization. If each individual is in seperate eppendorf, then sterilize the tweezers between every individual. Make sure to put an insect on a paper towel before you put in a tube with buffer, to get rid off external ethanol. Close the tube after adding the insect2).
(Remember to note on the homogenization sheet3) all of the important informations about the sample – trap ID, size of an insect, volume of a buffer, info about the Order/Group/species, sex.  Sterilize the tweezers with a flame after each sample.)

6.      Place the tubes in the homogenizer and grind two times for 30 s with a 5m/s speed.

7.      Place the samples in a thermal block/water bath at 55oC for 2 hours. If possible, vortex samples 1-2 times during incubation. 

8.     Remove the samples from the thermal block and, after cooling to room temperature, start DNA extraction or store in fridge or freezer.

1) Insect size categorys: [1] up to 2mm, [2] up to 5mm, [3] up to 1cm, [4] up to 2cm, [5] over 2 cm. In case of size category 1, 2 and most of the 3 standard amount (200 ul) of homogenization mix is enough. If you need to homogenize really big insects, e.g. bees, you need to cut it into smaller pieces. Divide bee into two pieces and put each into a 2 ml screw cap tube. Don’t forget to sterilize the tweezers/scalpel after each sample.  After incubation pour the content of one tube into another, vortex and store in a fridge or freezer.

2)
 
3)
 
SECTION: A. DNA extraction
4. Quantification 16S rRNA spike-in plasmid preparation

A. Sequence synthesizing, plasmid production, transformation into E. coli and bacteria culture. 
Plasmid sequences with artificial 16S rRNA targets are derrived from Tourlousse et al., 2016. 
In our workflow we are using Ec5502 at the stage of DNA extraction, and Ec5001 during PCR I. 

Use commercial services (i.e. Eurofins) to synthesise spike-in sequences and insert them onto pEX-A128 plasmid harboring ampicillin resistance.

After obtaining the plasmid you need to tranfer them into E.coli and culture the bacteria. For the description of those steps please look into the protocol by E. Iwaszkiewicz-Eggebrecht et al. (2023) dx.doi.org/10.17504/protocols.io.14egn33ryl5d/v2

B. Plasmid isolation from E. coli
The plasmids were isolated from E. coli using a GeneMATRIX PLASMID MINIPREP (EURx) kit following the manufacturer's instructions.
 
C.Plasmid digestion with restriction enzymes. 

To avoid complications during PCR that might have arisen because of circular shape pf plasmids, we decided to linearise them by cutting with a restriction enzyme. 

For plasmid Ec5502 we choose EcoRI-HF enzyme (NEB) and for Ec5001 AatII enzyme (NEB). Both are  cutting the plasmid effectively and leaving the barcode region intact. 

Materials:        
Restriction enzyme EcoRI-HF (for Ec5502) or AatII (For Ec5001) - we are using New England Biolabs enzymes, for the enzymes from a different manufacture the digestion protocol might be different 
CutSmart Buffer (provided with enzyme)        
Molecular water (nuclease free) 

PREPARE ON ICE!

C.1. Measure the concentration of isolated plasmid DNA on Qubit.
C.2. Remove restriction enzyme and CutSmart buffer from the frezer and put it on ice. 
C.3. Calculate how many ul of plasmid you need to add to the reaction in order to have ca. 100 ng of plasmid DNA in each reaction. At first add molecular water to the tube, then plasmid and CutSmart buffer. As a last component add restriction enzyme. Do not vortex the reaction! Just mix it by pipetting. Centrifuge shortly.
C.4. Put the strip tube/plate in a thermal cycler and set up the program. The incubation and inactivation temepratures and time can be found in table below.   

 

 C.5. After digestion freeze the samples or prepare 220 MLN copies single-use aliquots according to the instruction below. 

D. Preparing 220 MLN copies single-use aliquots.
 
Materials and equipment:
PolyA 100 (TE with PolyA tails in conc. 100 ng/ul)
Digested plasmid from previous step
Qubit fluorometer and Qubit HS assay kit
     
D.1. Measure digested plasmid DNA concentration using Qubit flurometer. 

D.2. Based on the received information calculate how many copies of a plasmid you have in one μl. 
You can do this based on the assumption that 1 ng of 16S plasmid DNA contains 220 MLN copies of the plasmid.
(This assumption is based on a calculation taking into account the size of the DNA molecule)

D.3. Based on performed calculations dilute spike-in’s samples with PolyA 100 into the concentration of 220 mln of copies/μl and prepare single-use aliquots of 10 μl each. 
To avoid molecula binding to the surface of the tube, use DNA low-binding eppendorf tube. 
Store in a frezer in -20˚C. 

E. Preparing 10 000 copies dilutions. 
In our laboratory we are using, as a standard dose, 20 000 copies per 40 ul of homogenate (20% of the sample/insect).

Materials: 
 • Ec5502 16S spike-in – 10 ul aliquote from previous step
 • sterile water
 • DNA low bind eppendorf tubes

Preparation: 
(1) add 210 ul of sterile water into aliquot tube (concentration after dilution 10 mln/ul)
(2) to the fresh tube add 180 ul of sterile water and 20 ul of dilution from step 1 (concentration after dilution 1 mln/ul)
(3) to the fresh tube add 1980 ul of sterile water and 20 ul of dilution from step 2. 
The final concentration after 3 dilutions is 10 000 copies/ul. 

!!! Start preparing plasmids only after you have completed point 1 of DNA Extraction protocol. 
It is important to minimize plasmid concentration loss due to the adhesion of plasmid molecules to the tube walls.

SECTION: A. DNA extraction
5. Homogenate purification

Materials: 
• SPRI magnetic beads (stored in the fridge) – take out from the fridge and keep in room temperature 
1 hour before use 
• deepwell storage plate
• 80% ethanol (freshly made) 
• TE x 1 
• magnetic stand
• clean PCR plates with caps

DNA extraction:

1.       From the tubes after digestion transfer 40 ul of homogenate into proper well on deep well plate.
 If the tubes after digestion were sored in a fridge or freezer make sure to warm them up on thermoblock before (the homogenate in the fridge becomes thick and difficult to pipet).

2.      If you are using spike-in add 2ul of freshly prepared 10 000 copies/ul dilution into each well. 

3.      Add 80 μL (x 2.0) of SPRI magnetic beads, and mix by pipetting (or vortex). 

4.      Incubate samples at room temperature for 5 minutes.

5.      Place the plate on the magnetic stand for 2 min. 

6.     Remove sthe supernatant. Make sure to not disturb the magnetic beads. 

7.      Pour alcohol into the plastic trough (reservoir) and add 200 μL of 80 % EtOH to each well using multichannel pipett. 

8. Make sure that the deep well plate still stays on the magnetic stand. Incubate for 30 seconds. 

9.      Remove the supernatant. 

10.      Repeat steps 7 – 9 for two ethanol washings.  

11.      Wait 5 min until dry (on magnetic stand). 

12.      Remove the plate from the magnetic stand, add 20 μL of TE x 1 into each well and mix by pipetting (or vortex). 

13.      Incubate the samples at room temperature for 5 minutes to elute the DNA. 

14.      Place on magnet stand for 2 min.

15.      Transfer supernatant (20 μL), containing DNA, to new 96-well PCR plate. Sign the plate and store in fridge (for few days) or freezer.


 
SECTION: A. DNA extraction
6. Measuring DNA concentration

Materials: 
• Quant-iT™ PicoGreen™ dsDNA Assay Kit (stored in the fridge) -  allow the Quant-iT PicoGreen reagent to warm to room temperature before opening the vial. It might take a while, so before using check if the vial content is liquid(!!!).
• Black, flat bottom 96-well plate

Equipment:
• Synergy HTX reader


Protocol for whole 96-well plate:
1.       Prepare 10 ml of 1xTE working solution (used for diluting the Quant-iT PicoGreen reagent, DNA
samples and PicoGreen standard) by mixing 9.5 ml of sterile, distilled water with 0.5 ml of 20xTE concentrate buffer.

2.      Mix 4975 ul of 1xTE, that you prepared in point one, with 25 ul of concentrated DMSO to obtain 5 ml of Quant-iT PicoGreen reagent working solution. Cover tube with reagent with aluminum foil, as PicoGreen reagent is susceptible to photodegradation.

3.      Prepare PicoGreen five-point standard curve. Add 2ul of PicoGreen standard (100 ug/mL) to 98ul of 1xTE. Take 10ul of this solution and add 90ul of 1xTE to obtain 10-fold dilution. Repeat this step twice to obtain 100 and 1000-fold dilutions. Arrange diluted standard on the plate adding 50ul of each dilution, starting from the highest concentration and ending with blank (50ul of 1xTE).

 
4.      Prepare the 96-well plate as in the picture:


 
5.      Add 50ul of Quant-iT PicoGreen reagent working solution to each well (standard curve and DNA samples). Use plastic reagent tray and multi-channel pipette to speed up the process. Cover plate with aluminum foil, mix well and incubate 2-5 minutes.

6.     Samples are ready to measure DNA concentration on Synergy HTX reader [15 min].



SECTION: B. Amplicon library preparation
7. Overview

For amplicon library preparation we use a two-step PCR approach. During the first step (PCR1) we amplify three target genes: COI, 16S V1-2 region and 16S V4 region, in multiplex reaction. Next we clean PCR1 products with magnetic beads. Finally, during the second step (PCR2) we add the indexes and illumina adapters to the amplified sequences.
 
SECTION: B. Amplicon library preparation
8. PCR I primers preparation

For each target gene we have 8 forward and 12 reverse primers with unique tags. What gives us in total 96 uniqe pairs combinations. Informative variable-length inserts (tags) are used to mitigate index switching that occures during sequencing on Illumina platform.

Primers scheme example:
 
Using stock primers [100uM] we are preparing primer mixes (separately for forward and reverse). Each mix contains three primers in concentration: COI [2.5uM], 16S V4 [10uM] and 16S V1-2 [5uM].

 
 
Finally primer mixes aliquots should be distributed into strip tubes: MA_P5 - MH_P5, M1_P7 - M8_P7 and M9_P7 - M12_P7. This will allow you to use multi-channel pipette during adding primer mixes to the PCR reactions.

SECTION: B. Amplicon library preparation
9. PCR I

Set up PCR 1 by creating a master mix as described below. We carry out PCR 1 in a total volume of 10 μL. To prepare the master mix for a 96-well plate, multiply the volumes for one reaction by 100. Plate will contain: 91 samples, two negative controls (with and without PCR spike-in), two positive controls and one empty well for PCR 2 negative control.

 
1.    In sterile tube prepare PCR reaction mix by mixing 2x QIAGEN Multiplex PCR Master Mix and sterile water, acording to the table above.

2.    If you are using PCR spike-in add 6 ul of PCR mix prepared in point one to well D12 of your PCR pate, it will be first negative control (without PCR spike-in). Otherwise skip poin 2 and go directly to point 4.

3.    Prepare plasmid spike-in in similar manner as for DNA extraction. You need to use different plasmid than before (e.g. Ec5001) and diluted it to lower concentration, 1000 copies per sample. Add 4 ul of diluted spike-in, containing 99 000 copies of plasmid to PCR mix and vortex it.

4.    Ditribute 6 ul of PCR mix to each well on the plate, except D12 (if you use plasmids) and H12.

5.    Using multi-channel pipette add correct primers mixes to the correct wells, according to the scheme belowe. Each well should contain unique pairs of primers.

 
6.    Add 2ul of DNA to the wells A1 - C12. Add 2ul of sterile water to wells D12 and E12. Add to the wells F12 and G12 2ul of DNA samples, that contains all target genes, so it will work as positive controls.

7.    Put the lids on the plate, centrifuge the plate and place in the thermocycler.

8.    Run PCR 1 using the following conditions [~2h]:


 
SECTION: B. Amplicon library preparation
10. Electrophoresis I

Run PCR 1 products on a 2,5% agarose gel (2 μL PCR product + 2 μL loading dye). It is enough to run just a subset of samples on the gel to see if the PCR reaction went properly. However remember to add all negative and posive controls on the gel.




SECTION: B. Amplicon library preparation
11. PCR  product cleaning

Clean up PCR 1 products by following the steps below. Make sure you prepare SPRI Beads as explained in the "Materials" section before starting this step. 

    1. Prepare fresh aliquots of 80% EtOH.
    2. Transfer all of PCR1 product to a 96 deep-well plate (8 - 10 μL per sample) and add 20 μL of SPRI Beads, and mix by pipetting (or vortex the entire plate using a plate shaker).
    3. Incubate mixture for 5 min at room temperature.
    4. Place the plate on a magnetic stand for 2 min.
    5. Remove supernatant (use multichannel pipette).
    6. Pour alcohol into a 100 mL reagent reservoir and, using a multichannel pipette, transfer 100 μL of 80% EtOH to each well.
    7. Incubate on a magnetic stand for 1 min.
    8. Remove supernatant.
    9. Repeat steps 6 – 8 above.
    10. Wait up to 4 min until the magnetic bead pellet gets dry (on magnetic stand).
    11. Remove samples from the magnetic stand, add 20 μL of TE buffer and mix by pipetting up and down (or vortex the entire plate using a plate shaker).
    12. Incubate mixture for 5 min at room temperature.
    13. Place back the plate on the magnetic stand for 2 min.
    14. Transfer supernatant (20 μL) to a new 96-well plate. 
    15. Mark the plate with a unique identifier and store at 4ºC (for a few days) or at -20ºC (for a longer period).

SECTION: B. Amplicon library preparation
12. PCR 2 (indexing)

Set up PCR 2 by creating a master mix as described below. We carry out PCR 2 in a total volume of 10 μL. To prepare the master mix for a 96-well plate, multiply the volumes for one reaction by 100. Plate will contain all samples from PCR 1 plus PCR 2 negative control in well H12.

 
1.    In sterile tube prepare PCR reaction mix by mixing 2x QIAGEN Multiplex PCR Master Mix and sterile water, acording to the table above.

2.    Ditribute 7 ul of PCR mix to each well on the plate.

3.    Using multi-channel pipette add correct indexing primers to the correct wells, that each well will contain unique pair of primers.

4.    Add 1ul of clean PCR 1 product to each well. Add 2ul of sterile water to well H12 (negative control of indexing).

7.    Put the lids on the plate, centrifuge the plate and place in the thermocycler.

8.    Run PCR 1 using the following conditions [~1h]:

 

SECTION: B. Amplicon library preparation
13. Electrophoresis II

Run PCR 2 products on a 2,5% agarose gel (2 μL PCR product + 2 μL loading dye). 

SECTION: C. Amplicon library pooling, QC, and sequencing
14. Pooling

Create a pool of samples (libraries) from each 96-well plate by mixing PCR 2 products in an 1.5 mL microtube, based on gel band intensity: 
    very strong band - 0.5 μL
    strong band - 1 μL
    medium band - 2 μL
    weak band - 4 μL
    no visible band - 8 μL

 
 

SECTION: C. Amplicon library pooling, QC, and sequencing
15. Pools cleaning

Clean the "plate pools" created above with Promega ProNex® Size-Selective Purification System. Follow the manufacturer's instructions, using 1/1.5 (v/v) pool to magnetic beads ratio. As standard we use 80 μL of pool for cleaning.
SECTION: C. Amplicon library pooling, QC, and sequencing
16. Measure DNA concentrations of the "plate pools" on Qubit Fluorometer using dedicated kit. Please follow the manufacturer's instructions. 

You can use Qubit‱ 1X dsDNA High Sensitivity (HS) or Broad Range (BR) Assay Kits.

SECTION: C. Amplicon library pooling, QC, and sequencing
17. Master Pool

Create Master Pool by mixing together the “plate pools” obtained in step 12, based on sample number (not all plates had to be full) and the DNA concentration. If pools that you are mixing togather differs in target genes composition (e.g one plate has only COI barcode and other plates all three target genes) then you have to take it into account.
SECTION: C. Amplicon library pooling, QC, and sequencing
18. Master Pool cleaning

Clean Master Pool by repeating point 15, but use 1/1.3 (v/v) pool to magnetic beads ratio. Suspend the final product in TE x 0.1 (or ddH2O). Volume in which to suspend depends on desired DNA concentration.
SECTION: C. Amplicon library pooling, QC, and sequencing
19. Measure DNA concentrations of the Master Pool on Qubit Fluorometer using dedicated kit. Please follow the manufacturer's instructions. 

You can use Qubit‱ 1X dsDNA High Sensitivity (HS) or Broad Range (BR) Assay Kit.
SECTION: C. Amplicon library pooling, QC, and sequencing
20. Quality check

Check quality of Master Pool (e.g. lack of primer-dimmers) by running them on Bioanalyzer or Tapestation before submitting for sequencing. In an unlikely event of detecting primer-dimers, repeat cleaning step as in point 16.



SECTION: C. Amplicon library pooling, QC, and sequencing
21. Sequencing options and considerations

The library pool is compatible with all Illumina platforms that produce at least 2x250 bp reads. The currently available platforms - Illumina MiSeq, NextSeq 2000, and NovaSeq 6000 - differ dramatically in output, cost of a sequencing flow cell, and cost per million read pairs, as listed in Table X.1 below:

 

Amplicon libraries prepared according to the current protocol can generally be combined with other library types, for example, metagenomic libraries prepared using Illumina TruSeq or NEBNext Ultra DNA kits, as long as there is no direct overlap in index sequences: we have never encounteres issues.
On the other hand, because relatively few researchers use longer-read flow cells on NextSeq and NovaSeq platforms, sequencing facilities may be unwilling to combine your samples with those submitted by other researchers for sequencing in a single lane or flow cell (as is done more often for 2x150bp sequencing submission), and you will generally need to pay for the whole sequencing flow cell.

When selecting the sequencing platform, we would recommend aiming for not less than 50,000 read pairs per library on average. To maximize the value per read and per library, you would ideally submit pools comprising many libraries, and use highest-throughput platforms. For example, the cost of sequencing a library to an average coverage of 100,000 read pairs on NovaSeq 6000 platform may be less than 1 EUR when submitting 8000 libraries at once - but the price would increase more than ten-fold when using MiSeq for the sequencing. At the same time, few projects so far aimed to sequence thousands of amplicon libraries --- and even if they did, preparing and pooling that many libraries will likely take a long time, translating to a long wait for the data. Also, the availability of sequencers and the turnaround time may be a consideration - and may be greater for lower-throughput platforms. Then, despite NovaSeq's superior per-read value, for logistics reasons, NextSeq or MiSeq may be preferred options for your project. Alternatively, you may consider pooling amplicon libraries from across many projects, and combining amplicons with genomic/metagenomic libraries.

When selecting sequencing platforms, it is also good to be aware of differences in data characteristics. For example, in sequencing data produced by the MiSeq platform, read quality tends to decrease substantially towards the end of the read, and is much lower and drops more quickly for the second read in the pair. The decrease in quality along the read length tends to be much less in data generated using platforms that rely on the newer patterned flow cell - NextSeq and NovaSeq (Fig. X1). On the other hand, platforms using patterned flow cells may be affected more by cross-contamination among samples that happens during sequencing: https://www.illumina.com/techniques/sequencing/ngs-library-prep/multiplexing/index-hopping.html


When sequencing libraries using flow cells with more than one lane (specifically, NovaSeq SPrime flow cell has two lanes), you may want to submit a single library Master Pool for sequencing on both lanes --- or alternatively, submitting different Master Pools, even with overlapping index combinations, for sequencing on different lanes. The latter approach will allow you to pack your libraries less densely in index space, likely reducing the extent of cross-contamination among samples.

SECTION: Protocol overview
1. Protocol overview, general instructions, and considerations

Quantitative Multi-Target Amplicon Sequencing is an approach originally developed for the characterization of microbiota in large collections of diverse insects. It enables the molecular identification of the host insects and the characterization of the abundance and genotype- and clade-level diversity and distribution of their associated bacteria. This is achieved through simultaneous amplification of insect (COI) and bacterial (16S rRNA) marker genes, the latter in a quantitative manner, and the preparation of Illumina-compatible sequencing libraries in a highly multiplexed manner.

This protocol has been initially developed and optimized for the cost-effective preparation of large numbers of amplicon sequencing libraries for these two target regions. However, it can be easily adapted to incorporate other marker regions for different organisms potentially present in samples, or adapted for other types of biological samples. Parts of the protocol can be executed on their own and combined with other methods. Many of the solutions implemented in the protocol, including quantification of targets or use of "phased" primers, are optional.

The protocol presented here starts with a set of 88 diverse ethanol-preserved insects selected from a Malaise trap. This number, alongside different positive and negative controls, is sufficient to fill a 96-well plate. 
It ends with a library pool suitable for sequencing using one of Illumina platforms compatible with at least 500-cycle kits.

Specifically, the protocol comprises the following steps:

A. DNA extraction
    > Sample homogenization
    > Quantification spike-in preparation and addition
    > Homogenate purification
    > DNA quantification
B. Amplicon library preparation
    > The overview of the library preparation process
    > First PCR
    > PCR product validation
    > PCR product purification
    > Second (indexing) PCR
C. Amplicon library pooling, QC, and sequencing
    > Estimating library concentrations
    > Library pooling
    > Library pool QC and preparation for sequencing
    > Sequencing options and considerations
D. An overview of the data analysis workflow
    > 
SECTION: A. DNA extraction
2. Specimen selection and preparation

Typically, we start with a batch of ethanol-preserved insects from a sweep net or a Malaise trap catch
Generally, we pre-select insects based on morphology, to maximize diversity and avoid sequencing large numbers of specimens representing the same species.
We have also successfully used insect specimens pre-barcoded following Meyer protocols that include HotSHOT DNA extraction step (https://link.springer.com/article/10.1186/s12915-021-01141-x). See Andriienko et al. (2024) for details (https://doi.org/10.1101%2F2024.04.30.591865).
We typically roughly sort insects by size. Very large insects may require a modified treatment, such as using a greater volume of lysis buffer (see section 3).
We use 88 individual insects per plate (one per well). The remaining wells are used for negative and positive controls.
We carefully organize and preserve sample metadata
We recommend rinsing specimens in clean ethanol before processing, especially if they originate from dense multi-species samples, and may have dust or body fragments of other insects attached.
It is recommended to air-dry at least the larger insects before processing them, to avoid the inhibitory effects of excess ethanol.... but wait until tubes for the DNA extraction (next step) are ready to go