Nov 26, 2024
Chagos eDNA metabarcoding protocol
- Rosie Dowell1
- 1ZSL
Protocol Citation: Rosie Dowell 2024. Chagos eDNA metabarcoding protocol. protocols.io https://dx.doi.org/10.17504/protocols.io.e6nvwb5nzvmk/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 25, 2024
Last Modified: November 26, 2024
Protocol Integer ID: 112738
Abstract
Full eDNA metabarcoding protocol for Chagos eDNA metabarcoding project, funded by the Bertarelli Marine Science Program.
Extraction from frozen encapsulated filters frozen, simplified from Spens et al, 2017.
Extraction from frozen encapsulated filters frozen, simplified from Spens et al, 2017.
Before extraction: Carefully wipe all surfaces with 5% bleach using clean tissue paper. Dry and wipe with 70% Ethanol using tissue paper. Syringes, tubes and pipettes should be sterilised in UV box for 30 mins prior to each set of extractions.
Label all filters using lab marker. Turn on incubators. Turn on heating block to 70°C to heat ddH2O for elution.
Make a premix of Lysis working solution by adding 720 μL ATL buffer and 80 μL proteinase K per sample provided by the kit.
Using a sterile filter tip, load 800 μL of lysis working solution into a sterile 3ml syringe with luer adapter.
Keeping the outlet end loosely closed with the outlet cap. Carefully add the 800 μL Lysis working solution to the filter using a 3ml sterile syringe with luer adapter. If bubbles/lysis solution can be seen from the outlet, tighten the cap. Close with an new inlet cap and handshake vigorously for a few seconds.
Incubate samples, while rotating, at 56°C for 24 hours.
Transfer: Remove ALL the liquid from outlet of capsule by using a Luer Lock syringe to pull the lysis buffer out of the filter capsule and add to 5 mL LoBind tube.
From this point on, this follows Qiagen B&T protocol. Add Buffer AL and ice-cold molecular grade 99% ethanol to the sample in equal volumes. Sample:Buffer:Ethanol = 1:1:1. Note: AL and ethanol can be premixed.
Vortex vigorously.
Pipet the mixture (max 650 μL at a time) into a DNeasy Mini Spin column in a 2 mL collection tube provided in the kit.
Spin in micro-centrifuge at 6000 ∗ g (8000 rpm) for 1 min.
Discard flow through.
Repeat steps 10-12 until all sample is filtered through DNeasy Mini spin column (most likely 4 times)
Place the DNeasy Mini spin column in a new 2 ml collection tube, add 500 μl Buffer AW1, and centrifuge for 1 min at 6000 ∗ g (8,000 rpm). Discard flow-through and collection tube.
Place the DNeasy Mini spin column in a new 2 ml collection tube, add 500 μl Buffer AW2, and centrifuge for 3 min at 20,000 ∗ g (14,000 rpm) to dry the DNeasy membrane. Discard flow-through and collection tube.
Place spin column in a new collection tube, centrifuge 1 min at 17,000 ∗ g (13,000 rpm).
Transfer spin column to a new 1.5 or 2 mL DNA LoBind tube with caps removed.
Place tubes with spin columns, four at a time, on a 70°C heating plate, add 100 μl 70°C ddH2O to the membrane, immediately transfer spin column with filter to room temperature.
Incubate at room temperature for 10 min.
Centrifuge for 1 min at 6,000 ∗ g (8,000 rpm)
Re-elute DNA from DNA LoBind tube. (Apply eluate back on spin column on heating plate).
Incubate at room temperature for 10 min.
Centrifuge for 1 min at 6,000 ∗ g (8,000 rpm)
Discard the spin column.
Transfer DNA to pre-marked DNA LoBind tube with lid intact.
Qubit 2 μL for DNA measurement.
Store at -20°C or at -80°C.
Extraction from Durapore filters, rolled and stored either frozen or in RNAlater in 5ml Cryovials. Also simplified from Spens et al, 2017.
Extraction from Durapore filters, rolled and stored either frozen or in RNAlater in 5ml Cryovials. Also simplified from Spens et al, 2017.
Before extraction: Carefully wipe all surfaces with 5% bleach using clean tissue paper. Dry and wipe with 70% Ethanol using tissue paper. Syringes, tubes and pipettes should be sterilised in UV box for 30 mins prior to each set of extractions.
Label all filters using lab marker. Turn on incubators. Turn on heating block to 70°C to heat ddH2O for elution.
Make a premix of Lysis working solution by adding 720 μL ATL buffer and 80 μL proteinase K per sample provided by the kit.
If stored in RNAlater, remove the buffer using a sterile pipette tip and add to a labelled tube.
Add 800 μL Lysis working solution to the filter in the cyrovial. Close the cap and handshake vigorously for a few seconds.
Incubate all samples, while rotating, at 56°C for 24 hours.
Using a pipette, transfer ALL the liquid from the cryovial to a 5 mL LoBind tube. Add the lysis buffer from the pellet incubation.
From this point on, this follows Qiagen B&T protocol. Add Buffer AL and ice-cold molecular grade 99% ethanol to the sample in equal volumes. Sample:Buffer:Ethanol = 1:1:1. Note: AL and ethanol can be premixed.
Vortex vigorously.
Pipet the mixture (max 650 μL at a time) into a DNeasy Mini Spin column in a 2 mL collection tube provided in the kit.
Spin in micro-centrifuge at 6000 ∗ g (8000 rpm) for 1 min.
Discard flow through.
Repeat steps 10-12 until all sample is filtered through DNeasy Mini spin column (most likely 4 times)
Place the DNeasy Mini spin column in a new 2 ml collection tube, add 500 μl Buffer AW1, and centrifuge for 1 min at 6000 ∗ g (8,000 rpm). Discard flow-through and collection tube.
Place the DNeasy Mini spin column in a new 2 ml collection tube, add 500 μl Buffer AW2, and centrifuge for 3 min at 20,000 ∗ g (14,000 rpm) to dry the DNeasy membrane. Discard flow-through and collection tube.
Place spin column in a new collection tube, centrifuge 1 min at 17,000 ∗ g (13,000 rpm).
Transfer spin column to a new 1.5 or 2 mL DNA LoBind tube with caps removed.
Place tubes with spin columns, four at a time, on a 70°C heating plate, add 100 μl 70°C ddH2O to the membrane, immediately transfer spin column with filter to room temperature.
Incubate at room temperature for 10 min.
Centrifuge for 1 min at 6,000 ∗ g (8,000 rpm)
Re-elute DNA from DNA LoBind tube. (Apply eluate back on spin column on heating plate).
Incubate at room temperature for 10 min.
Centrifuge for 1 min at 6,000 ∗ g (8,000 rpm)
Discard the spin column.
Transfer DNA to pre-marked DNA LoBind tube with lid intact.
Qubit 2 μL for DNA measurement.
Store at -20°C or at -80°C.
Qubit DNA Quantification
Qubit DNA Quantification
The Qubit kit consists of the Qubit sample buffer, Qubit reagent and two standards (or if 1X qubit kits are bought, the buffer and reagent are pre-mixed). The Qubit sample buffer is simply the solution used to dilute your samples, the Qubit reagent is the fluorescent dye that, in this case, binds to dsDNA. It is advisable to prepare standards each time you use the Qubit. The Qubit requires specific 0.5ml thin wall tubes. The final volume in the assay tubes need to be 200ul, but you can vary the volume of DNA you add, 1-20ul, to allow concentration to be maintained within detection range. Standards are always added at a volume of 10ul.
If there is lots of samples, there is a qubit flex which allows for running strips of tubes rather than one at a time. The chemistry is exactly the same but the tubes are smaller, and must be thin walled as above.
Remove Qubit reagents from the fridge 30 minutes before use and vortex just before use.
If not using 1X buffer, make enough master mix for duplicates of each sample + 2 for standards
Per sample: 199ul Qubit reaction buffer and 1ul of Qubit reagent
Label Qubit tubes for the standards and the duplicate sample tubes.
To each standard tube add 190ul of master mix and 10ul of the relevant standard.
To each sample tube add 198ul of master mix and 2ul of library.
Turn on the Qubit and select DNA - dsDNA HS assay.
Select to read new standards.
Follow on screen instructions to read standards and insert the first assay tube.
After reading the first assay tube select “calculate stock concentration”.
Change the sample volume to 2μl and ensure that the units are ng/μl - record the concentrations.
1st round PCR with metabarcoding assays
1st round PCR with metabarcoding assays
18S (AmarZett)
Triplicate 10 μL PCRs should be performed for each sample to be sequenced.
The PCR reaction should be made up of 2 μL of 5x HOT FIREPol® Blend Master Mix Ready to Load, 0.4 μL of forward and reverse primers (10nM), 5.7 μL of ddH2O and 1.5 μL of DNA template.
The PCR protocol comprised 95°C denaturation for 15 minutes, followed by 30 cycles of 95°C for 10 seconds, 57°C for 30 seconds and 72°C for 30 seconds, with a final elongation step of 72°C for 7 minutes.
Pool the three replicates for each sample and run 5 μL on 1% agarose gel with SYBR safe stain to check for successful amplification (TLUM light and UV32 filter). COI (Leray)
COI (Leray)
Triplicate 25 μL PCRs should be performed for each sample to be sequenced.
The PCR reaction should be made up of 5 μL HOT FIREPol 5X MasterMix, 0.5 μL of each primer (10μM), 2 μL extracted DNA and 17 μL H2O for a final reaction volume of 25 μL.
The PCR protocol comprised 95°C denaturation for 95°C for 15 minutes followed by 35 cycles of 95°C for 1 60s, 55°C for 60s and 72°C for 60s, with a final extension of 72°C for 5 minutes.
Pool the three replicates for each sample and run 5 μL on 1% agarose gel with SYBR safe stain to check for successful amplification (TLUM light and UV32 filter). 12S (MiFish-U)
12S (MiFish-U/E)
Triplicate 10 μL PCRs should be performed for each sample to be sequenced.
The PCR reaction should be made up of 3 μL HOT FIREPol 5X MasterMix, 0.7 μL of each primer (MiFish-U-F & -R (10μM) and MiFish-E-F & -R (5μM)), 2.5 μL extracted DNA and 6.7 μL H2O for a total volume of 15 μL. Thermal conditions consisted of an initial denaturation step of
The PCR protocol comprised 95°C denaturation for 95°C for 2 minutes followed by 40 cycles of 95°C for 15s, 65°C for 30s and 72°C for 30s, with a final extension of 72°C for 5 minutes.
Pool the three replicates for each sample and run 5 μL on 1% agarose gel with SYBR safe stain to check for successful amplification (TLUM light and UV32 filter). If not using the green mastermix, each sample will need to be mixed with loading dye and gel red for visualisation (no gel stain). 16S (EMP)
16S (EMP)
Triplicate 25 μL PCRs should be performed for each sample to be sequenced.
The PCR reaction should be made up of 5 μL HOT FIREPol 5X MasterMix, 0.5 μL of each primer (10μM), 1 μL extracted DNA and 18 μL H2O to make the mixture up to 25 μL.
The PCR protocol comprised 94°C for 12 minutes followed by 35 cycles of 94°C for 45s, 50°C for 60s and 72°C for 90s, with a final extension of 72°C for 10 minutes.
Pool the three replicates for each sample and run 5 μL on 1% agarose gel with SYBR safe stain to check for successful amplification (TLUM light and UV32 filter). If not using the green mastermix, each sample will need to be mixed with loading dye and gel red for visualisation (no gel stain).
Primer sequences (inlcuding illumina adapters)
Primer sequences (inlcuding illumina adapters)
| A | B | |
| Primer | Primer sequence in bold (5’ – 3’) | |
| 18S AmarZett (Amaral-Zettler et al., 2009) | ||
| 1380F | TCGTCGGCAGCGTCAGATGTGTATAAGAGACAGCCCTGCCHTTTGTACACAC | |
| 1510R | GTCTCGTGGGCTCGGAGATGTGTATAAGAGACAGCCTTCYGCAGGTTCACCTAC | |
| COI (Leray et al., 2013) | ||
| mlCOIintF | TCGTCGGCAGCGTCAGATGTGTATAAGAGACAGGGWACWGGWTGAACWGTWTAYCCYCC | |
| jgHCO2198 | GTCTCGTGGGCTCGGAGATGTGTATAAGAGACAGTAIACYTCIGGRTGICCRAARAAYCA | |
| 12S MiFish (Miya et al, 201) | ||
| MiFish-U-F | TCGTCGGCAGCGTCAGATGTGTATAAGAGACAGGTCGGTAAAACTCGTGCCAGC | |
| MiFish-U-R | GTCTCGTGGGCTCGGAGATGTGTATAAGAGACAGCATAGTGGGGTATCTAATCCCAGTTTG | |
| MiFish-E-F | TCGTCGGCAGCGTCAGATGTGTATAAGAGACAGGTTGGTAAATCTCGTGCCAGC | |
| MiFish-E-R | GTCTCGTGGGCTCGGAGATGTGTATAAGAGACAGCATAGTGGGGTATCTAATCCTAGTTTG | |
| V4 16S (Earth Microbiome Project) | ||
| 515-F | TCGTCGGCAGCGTCAGATGTGTATAAGAGACAGGTGYCAGCMGCCGCGGTAA | |
| 806-R | GTCTCGTGGGCTCGGAGATGTGTATAAGAGACAGGGACTACNVGGGTWTCTAAT | |
First round PCR bead clean up (protocol the same for each marker from this point)
First round PCR bead clean up (protocol the same for each marker from this point)
Before you start: The volume of each sample should be about 70μl (70 μL total PCR product with 5 μL run on gel). Bring the Ampure XP beads to room temperature (~30 minutes) and thoroughly mix. Make up 400μl of 80% ethanol for each sample to be cleaned. Prepare an empty tip box filled with tissue to discard liquid waste into. Add the beads to a reagent reservoir or strip of tubes if cleaning a plate of samples.
Vortex clean-up beads for 30s.
Add 60 μL of beads to each sample and pipette mix 10 times - incubate the plate at room temperature for 5 minutes.
Put the plate on the magnetic stand for 2 minutes. There are two magnetic plate stands, one in Kev’s drawer and one in the drawer under the MiSeq, you can use either. They pull the beads in slightly different ways so choose whichever you find easiest to not disturb the bead pellets.
Remove 120 μl of supernatant and discard but avoid disturbing the beads.
If beads are also being aspirated, it might be necessary to remove less initially and remove the rest of the volume with a 10μl tip afterwards.
With the plate still on the magnet add 200μl of 80% ethanol without disturbing the beads - Incubate for 30 seconds.
Remove and discard the supernatant.
Repeat steps 5-6 for a total of two washes.
Incubate the plate for 5 minutes at room temperature to dry the beads.
Using a 10μl pipette remove and discard any remaining ethanol from each sample.
Remove the plate from the magnet, add 52.5μl of RSB or PCR grade water to each well and pipette up and down 10 times to re-suspend the beads - Incubate at room temperature for 2 minutes.
Put the plate back onto the magnet and incubate for 2 minutes or until the solution has cleared.
Transfer 50μl of the clean product to a new PCR tube / plate.
Index PCR
Index PCR
Individual samples are identified post sequencing by the indexes that are added during this PCR. The libraries are dual-indexed, the combination of indexes is unique for each sample. Before starting in the lab decide which indexes are to be used – we have previously used Nextera XT index kits B or C, which come with enough indexes to dual index 96 samples, four times (each box can has 96 indexes and can index two plates (192 samples). The Illumina guidance on index choice and low-plexity pooling is included in Appendix-1.
Make the PCR master mix, using 10 μL of 5X hot fire pol and 25 μL of water per sample.
Aliquot 35 μL of master mix into the bottom of each well, try to avoid the edges of the wells so you can keep tips clean when pipetting the indexes.
Arrange index 1 and 2 primers in the “Index Fixture Plate” (available in the post PCR genomic lab). Take a picture of them in order so you can be sure of the index combinations.
Set a fresh PCR plate on the “index Fixture Plate” and add 5ul of the corresponding index primer 1 to each well. To reduce the need for a new pipette tip for each well, pipette the indexes onto opposite sides of well walls to avoid contamination.
Add 5ul of the corresponding index primer 2 to each well.
Add 5ul of cleaned first round PCR product to the corresponding well and pipette mix.
Seal the plate and briefly centrifuge.
Run a PCR protocol comprising 95°C denaturation for 15 minutes, followed by 8 cycles of 95°C for 30 seconds, 55°C for 1 minute, 72°C for 1 minute, and a final elongation step of 72°C for 10 minutes.
Index PCR clean up
Index PCR clean up
Before you start: Bring the Ampure XP beads to room temperature (~30 minutes) and thoroughly mix. Make up 400μl of 80% ethanol for each sample to be cleaned. Prepare an empty tip box filled with tissue to discard liquid waste into. Add the beads to a reagent reservoir or strip of tubes if cleaning a plate of samples.
Vortex clean-up beads for 30s.
Add 56 μL of beads to each sample and pipette mix 10 times- Incubate the plate at room temperature for 5 minutes.
Put the plate on the magnetic stand for 2 minutes.
Remove 42 μL of supernatant and discard, avoid disturbing the beads.
If beads are also being aspirated, it might be necessary to remove less initially and remove the rest of the volume with a 10μl tip afterwards.
With the plate still on the magnet add 200 μL of 80% ethanol without disturbing the beads - Incubate for 30 seconds.
Remove and discard the supernatant.
Repeat steps 5-6 for a total of two washes.
Incubate the plate for 5 minutes at room temperature to dry the beads.
Using a 10μl pipette remove and discard any remaining ethanol from each sample.
Remove the plate from the magnet, add 27.5 μL of PCR grade water to each well and pipette up and down 10 times to re-suspend the beads - Incubate at room temperature for 2 minutes.
Put the plate back onto the magnet and incubate for 2 minutes or until the solution has cleared.
Transfer 25 μL of the clean product to a new PCR plate.
PCR Check on TapeStation (D1000 HS)
PCR Check on TapeStation (D1000 HS)
Label 0.2ml tubes, the first for the ladder, then one for each sample.
Add 3μl of sample buffer to each tube.
To the first tube add 1μl of ladder.
Add 1ul of PCR product to the remaining tubes.
Seal the tubes and put onto the plate vortex for 60 seconds.
Briefly spin down.
Remove the lids (cut hinged lids so they come off totally), check liquid is still in the very bottom of the tube and put into the tube holder in the TapeStation.
Insert a tape, and make sure the loading tip rack is full.
In the TapeStation controller software select the relevant tube loading positions and name the samples in the sample table.
Start the run (save the run stating it is the 1st round PCR, your name and the date).
Once complete: In TapeStation analysis click on “Electropherogram” and check each sample for amplicon sizing (target gene +100BP) and signs of primer dimer.
Sample QC: Qubit
Sample QC: Qubit
Vortex and centrifuge Qubit reagents
In a 1.5ml tube prepare 5 reactions of master mix by adding 5μl of Qubit reagent to 995μl of sample buffer, vortex and briefly centrifuge
Add the following to 5 Qubit tubes -
Tube-1 190μl of master mix, 10μl of standard 1
Tube-2 190μl of master mix, 10μl of standard 2
Tube-3 198μl of master mix, 2μl of pooled sample
Tube-4 198μl of master mix, 2μl of pooled sample
Tube-5 198μl of master mix, 2μl of pooled sample
Turn on the Qubit and select DNA - dsDNA HS assay
Select to read new standards
Follow on screen instructions to read standards and sample tubes
After reading the first sample tube select “calculate stock concentration”, change the sample volume to 2μl and change the units to ng/μl
Record the concentrations for the three sample replicates
Calculate nM values based on amplicon size from previous TapeStation values
MiSeq Loading
MiSeq Loading
Denature and Dilute Sample and PhiX
Prepare a fresh dilution of 0.2N NaOH - 400μl H2O + 100μl 1N NaOH (in fridge)
To a 1.5ml tube add 10μl of sample pool (or 1:10 sample pool if necessary) and 10μl of 0.2N NaOH
To a different 1.5ml tube add 2μl of 10nM PhiX, 3μl of H2O and 5μl of 0.2N NaOH
Vortex the library and PhiX tubes for 5 seconds and spin for 1 minute at 400rcf
Incubate the tubes for 5 minutes at room temperature
To the denatured library tube add 980μl of HT1
To the denatured PhiX tube add 990μl of HT1, now 20pM (20pM dilution can be stored at -20 for three weeks)
Vortex library and PhiX and centrifuge for 1 minute
Dilute the sample to 3.5pM using the volumes from the Library Dilution Template sheet
Dilute the PhiX to 3.5pM by adding 175μl of 20pM PhiX to 825μl of HT1
Invert library and PhiX tube to mix and pulse centrifuge
Put 950μl of 3.5pM library in a 1.5ml tube and add 50μl of 3.5pM PhiX, put this sample on ice
Prepare Reagent Cartridge
Remove the cartridge from the water bath and dry, tap on paper towel to remove as much water as possible from the base
Invert cartridge 10 times to mix reagents and inspect the reservoirs to make sure they are fully defrosted
Tap on the bench to make sure there are no bubbles at the bottom of the tubes
Set aside the MiSeq cartridge until the library is ready to Load
Load Flow Cell
On the MiSeq press sequence
Remove the flow cell and PR2 reagent box from fridge
Carefully remove flow cell from the buffer and rinse thoroughly with dd H2O, dry using Kimwipes paying particular attention to the edges of the glass
Wet another Kimwipe with absolute ethanol and clean the glass slide on both sides, avoid contact with the rubber gasket
Check the flow cell has no smudges or fibres from the wipes
Follow the instructions on the MiSeq to load the flow cell and PR2 buffer bottle, empty and replace the waste bottle
Heat Denature Sample
Using the heat block incubate the combined Library and PhiX tube for 2 minutes at 96⁰C
After the incubation invert the tube 2 times to mix and place in ice water bath for 5 minutes
Load MiSeq Cartridge
Using a 1000μl pipette tip pierce well 17 (it might be 16 – its labelled sample) on the reagent cartridge
Load 600μl of the heat denatured Library / PhiX pool into the cartridge
Load the cartridge into the MiSeq and wait for it to read the RFID
Follow on screen instructions, check the cycle numbers for the sequencing reads and the indexes
Wait for the MiSeq to complete its pre-run checks and then press start