Nov 19, 2021
A streamlined protocol for high-throughput amplification-based analysis of DNA samples via nanopore sequencing (based on the 96-well PCR barcoding kit)
- Jordan Cuff1,
- HooperAS 2,
- Shrinivas Nivrutti Dighe2,
- Angela Marchbank2,
- Peter Kille2
- 1Newcastle University;
- 2Cardiff University
- Foraging Ecology Research Group
Protocol Citation: Jordan Cuff, HooperAS , Shrinivas Nivrutti Dighe, Angela Marchbank, Peter Kille 2021. A streamlined protocol for high-throughput amplification-based analysis of DNA samples via nanopore sequencing (based on the 96-well PCR barcoding kit). protocols.io https://dx.doi.org/10.17504/protocols.io.bw2cpgaw
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: July 30, 2021
Last Modified: August 26, 2023
Protocol Integer ID: 52004
Keywords: nanopore, sequencing, multiplex, amplification, 96-well, pcr barcoding, environmental, using nanopore sequencing, nanopore sequencing, available kits from nanopore facilitate, well ligation kit from oxford nanopore, nanopore facilitate, nanopore, effective sequencing, effective sequencing of long fragment dna, holistic dna amplification, yield dna sample, oxford nanopore, samples of genomic dna, dna sample, low dna yield, genomic dna, analysis of dna sample, sites onto all extant dna, sequencing run, well pcr barcoding kit, extant dna, long fragment dna, well ligation kit, pcr barcoding kit, dna, pcr barcoding, large numbers of sample, streamlined protocol, smaller numbers of sample
Abstract
Nanopore sequencing facilitates the rapid and cost-effective sequencing of long fragment DNA for a massive range of applications. When looking to holistically analyse low-yield DNA samples using nanopore sequencing, the optimal method is likely to involve the PCR Barcoding Kit. This effectively involves blunt end ligation of priming sites onto all extant DNA for holistic amplification to achieve yields suitable for nanopore sequencing. The currently available kits from nanopore facilitate the multiplexing of 96 samples in one sequencing run using this method, but the reagent costs are inherently multiplicative. This protocol is designed to streamline (in terms of cost, reagents and time) the process of sequencing up to 96 samples of genomic DNA through nanopore sequencing.
This protocol is best applied to large numbers of samples (up to 96). For smaller numbers of samples, consider the smaller "PCR Barcoding" kits provided by nanopore which similarly achieve holistic DNA amplification and sequencing, but without the need for additional adapter ligation. The protocol is best suited to samples with low DNA yields (100 ng input is recommended). If you can input 1000 ng of DNA from each of your samples, consider using the 96-well Ligation Kit from Oxford Nanopore which can similarly be streamlined in terms of cost and time, but avoids the amplification step.
Guidelines
This protocol is designed specifically around the Oxford Nanopore Technologies PCR Barcoding Expansion 1-96 (EXP-PBC096) kit. The kit protocol deviates from the standard PCR barcoding kit protocol, existing as a hybrid between that and the ligation kit (SQK-LSK109). As such, both EXP-PBC096 and SQK-LSK109 will be required (alternative protocols are possible using SQK-LSK110).
This protocol is designed to streamline (in terms of both cost/reagents and time) the process of sequencing many (up to 96) samples of genomic DNA using nanopore sequencing. The process involves ligating priming sites onto the genomic DNA, priming and amplifying the full DNA strands and then ligating adapters onto the amplified products.
Materials
Equipment:
Qubit spectrophotometer (or alternative DNA concentration measurement system)
96-well plate magnetic rack
Thermocycler
MinION nanopore sequencer
Pipettes (P2, P20, P200, P1000, or otherwise overlapping range of volumes)
Consumables:
Plasticware
Ethanol
Ice
Oxford Nanopore Technologies PCR Barcoding Expansion 1-96 (EXP-PBC096)
Oxford Nanopore Technologies Ligation Sequencing Kit (SQK-LSK109)
Oxford Nanopore Technologies Flow Cell Priming Kit (EXP-FLP002)
Oxford Nanopore Technologies flongle and/or R9.4.1/R10.3 flow cell
SPRI beads
AMPure beads
Qubit BR dsDNA assay kit (HS dsDNA may be more applicable in some cases; alternative DNA concentration measurement methods are equally viable)
Nuclease-free water
NEB Ultra II End Prep reaction buffer and enzyme mix
NEB Blunt TA Ligase Master Mix
NEB 2X LongAmp Taq Master Mix
NEBNext FFPE DNA Repair buffer and enzyme mix
NEBNext Quick T4 DNA Ligase buffer and enzyme
10mM Tris-HCl pH 8.0 with 50 mM NaCl
Troubleshooting
Safety warnings
Please review the COSSH documentation for all reagents to ensure safe use and what to do in the event of accidents.
Before start
This protocol is best applied to large numbers of samples (up to 96). For smaller numbers of samples, consider the smaller "PCR barcoding" kits provided by nanopore which similarly achieve holistic DNA amplification and sequencing, but without the need for adapter ligation at the end. The protocol is best suited to samples with low DNA yields (100 ng input is recommended). If you can input 1000 ng of DNA from each of your samples, consider using the 96-well ligation kit from Oxford Nanopore which can similarly be streamlined in terms of cost and time, but avoids the amplification step.
Prepare DNA
Extract and purify your DNA in a manner appropriate for the sample type, the hypothesis being tested and this protocol. Consider the length of DNA that you are ideally interested in and how the tissue/sample type might best be lysed/extracted from. This protocol will be most successful when using samples with low concentrations of contaminants/inhibitors.
Consider whether a size selection/clean/concentration would be appropriate for the samples. If targeting long fragments, clean the samples with a 0.4X SPRI Select bead clean, washing twice with 200 μL 70% ethanol and eluting into 12.5 μL nuclease-free water (the step-by-step protocol for bead cleans is given below in Step 7). If the samples are low concentration or contain a high concentration of inhibitors/contaminants, consider carrying out a 1X bead clean to concentrate them and remove inhibitors.
Consider running the samples in duplicate/triplicate for best results. Ideally, these would use different barcodes downstream for post-sequencing comparison of results between samples to ascertain consistency.
Ligation of barcode adapters
3h
Before beginning the ligation process, it would be ideal to quantify the concentration of your samples using Qubit high-sensitivity/broad-range (depending on application) dsDNA assays or similar, but this can be omitted if necessary.
10m
In a new 0.2 ml tube (or 96-well plate), set up the following reaction for each sample:
| A | B | |
| Component | Volume | |
| DNA in nuclease-free water | 12.5 µL | |
| Ultra II End Prep Reaction Buffer | 1.75 µL | |
| Ultra II End Prep Enzyme Mix | 0.75 µL | |
| Total | 15 µL |
15m
Consider including negative and positive controls alongside your samples which can be sequenced for later quality control.
Incubate the samples as follows:
| A | B | |
| Temperature | Time | |
| 25 °C | 10 mins | |
| 65 °C | 5 mins | |
| 4 °C | 1 min |
16m
Get your barcode adapter ligation reaction ready in the order given:
| A | B | |
| Component | Volume | |
| End-prepped DNA (from last step) | 15 μL | |
| BCA (Barcode Adapter) | 8 μL | |
| Blunt TA Ligase Master Mix | 10 μL | |
| Total | 33 μL |
15m
Incubate the samples as follows:
| A | B | |
| Temperature | Time | |
| Room temperature (~20 °C) | 20 mins | |
| 70 °C | 10 mins | |
| On ice/4 °C | 1 min |
31m
Clean the adapter-ligated DNA with 0.4X AMPure beads (13.2 μL), washing twice with 200 μL 70% ethanol. Resuspend in 26 μL nuclease-free water, leaving 1 μL behind to prevent bead uptake.
1h 30m
If unfamiliar with bead cleaning, it will be described once here which can be referred to for subsequent cleans. Stay vigilant regarding the volume/ratio of beads to add, the solution used to wash the beads (i.e., ethanol vs. SFB/LFB) and the resuspension volume.
First, resuspend the beads in solution, ensuring they are at room temperature.
Add the required volume of beads (e.g., 0.4X clean of 33 μL sample requires an addition of 13.2 μL bead solution) and mix by gentle flicking and inversion (slower) or pipetting (risks fragmentation).
Incubate the solution for 5 mins (unless not physically mixed, in which case incubate for at least 10 mins). The solution can be incubated on a mixer to ensure thorough mixing.
Prepare the cleaning solution (e.g., make up fresh 70 % ethanol or thaw LFB/SFB).
Place the sample on a magnet for 5 mins to separate the beads from the rest of the solution.
Keeping the sample on the magnet, pipette off the supernatant and wash the beads with the cleaning solution (e.g., 200 μL 70 % ethanol) without disturbing the pelleted beads.
Incubate for 30 seconds and remove the ethanol supernatant by pipetting without disturbing the pelleted beads. Wash the beads with a further round of the cleaning solution (e.g., 200 μL 70 % ethanol) without disturbing the pelleted beads.
Pipette off any residual cleaning solution (briefly spinning down first can help, but ensure the sample is back on the magnet before attempting to remove any solution). Briefly air dry the beads (waiting for the beads to lose their glossy shimmer and become matte - this is usually ~30-120 seconds for SPRI and AMPure depending on the efficiency with which cleaning solution is removed).
Resuspend the pellet with the required resuspension volume (e.g., 25 μL nuclease-free water) and incubate for 5 mins.
Pellet the beads on a magnet until the solution is clear and colourless, and the beads are fully separated.
Remove and retain the required volume of resuspended DNA in a new tube.
Amplification of libraries
5h 15m
LongAmp Hot Start Taq 2X Master Mix, Barcode Primer 1 (BP 01).
Set up the following reaction:
| A | B | |
| Volume | Reagent | |
| 10 µl | Adapter-ligated DNA | |
| 14 µl | Nuclease-free water | |
| 1 µl | Barcode Primer (BC01-96, one per reaction) | |
| 25 µl | 2X LongAmp Taq Master Mix | |
| Total: 50 µl |
25m
Mix the reaction by gentle flicking and inversion, and incubate as follows:
| A | B | C | |
| Temperature | Time | Cycles | |
| 95 °C | 3 mins | 1 | |
| 95 °C | 0:15 mins | 18 | |
| 62 °C | 0:15 mins | ||
| 65 °C | V mins | ||
| 65 °C | 6 mins | 1 | |
| 4 °C | - | - |
The extension length (V) depends on the expected DNA length. Approximately 50 seconds per kbp is recommended (e.g., 16:40 for expected fragment sizes of ~20 kbp). Sizes can be pre-determined using gel electrophoresis or digital analysis tools such as TapeStation or Qiaxcel.
3h 15m
Measure the concentration of your samples using Qubit high-sensitivity/broad-range (depending on expected concentration) dsDNA assays (or similar).
1h
Pool your barcoded samples together based on concentration to achieve approximate equimolarity.
15m
Divide each concentration by the largest concentration across all samples to determine the volume of each to add to achieve approximate equimolarity.
Clean the adapter-ligated DNA with 1X AMPure beads, washing twice with 200 μL 70% ethanol. Resuspend in 50 μL nuclease-free water, taking 48 μL out (leaving 2 μL behind to prevent bead uptake), and using 1 μL to measure concentration.
20m
Ligation of adapters
1h 20m
Get your adapter ligation reaction ready in the order given:
| A | B | |
| Component | Volume | |
| DNA | 47 µL | |
| DNA CS | 1 µL | |
| NEBNext FFPE DNA Repair Buffer | 3.5 µL | |
| NEBNext FFPE DNA Repair Mix | 2 µL | |
| Ultra II End-prep reaction buffer | 3.5 | |
| Ultra II End-prep enzyme mix | 3 | |
| Total | 60 µL |
10m
Incubate the sample as follows:
| A | B | |
| Temperature | Time | |
| 20 °C | 5 mins | |
| 65 °C | 5 mins |
10m
Clean the sample with 1X AMPure beads, washing twice with 200 μL 70% ethanol. Resuspend in 61 μL nuclease-free water, taking 60 μL out (leaving 1 μL behind to prevent bead uptake).
You can theoretically skip this clean, but if you do, do not incubate for longer than 10 mins after the next step.
20m
Get your ligation reaction ready in the order given:
| A | B | |
| Component | Volume | |
| DNA | 60 µL | |
| Adapter Mix AMX | 5 µL | |
| Ligation Buffer LNB (or T4 buffer) | 25 µL | |
| NEBNext Quick T4 DNA Ligase | 10 µL | |
| Total | 100 µL |
10m
Incubate at room temperature (~20 °C) for 10 mins.
10m
Clean the sample with 1X AMPure beads, washing twice with either LFB or SFB (DO NOT CLEAN WITH ETHANOL; SFB for target fragments less than 3 kbp in length, otherwise use LFB). Resuspend in 17 μL elution buffer (EB, or 10mM Tris-HCl pH 8.0 + 50mM NaCl), leaving 1 μL of it behind and measuring the concentration of a further 1 μL using Qubit or similar.
Target input should be:
5-50 fmol for R9.4.1 flow cell
25-75 fmol for R10.3 flow cells
3-20 fmol for flongle flow cells
20m
Loading of library (flongle)
30m
If looking to check the quality of the DNA for sequencing, or only looking to generate relatively small amounts of sequencing data, consider sequencing on a flongle.
Allow the flongle to reach room temperature for 20-30 mins ahead of loading of the library. Insert the flongle into the MinION sequencer and check the number of active pores available. Visually check the flongle for air bubbles.
Thaw the sequencing buffer (SQB), loading beads (LB), flush tether (FLT) and flush buffer (FB) at room temperature and vortex all but the FLT briefly and spin them down.
Put together the following flush solution:
| Volume | Reagent | |
| 3 µl | Flush Tether (FLT) | |
| 117 µl | Flush Buffer (FB) | |
| Total: 120 µl |
Of this, add 100 μL to the flongle via the priming port, with caution to avoid the introduction of air bubbles.
Ensure the beads of the LB are suspended by vortexing again immediately before pipetting, then assemble the following solution:
| Volume | Reagent | |
| 10 µl | Loading Beads (LB) | |
| 15 µl | Sequencing Buffer (SQB) | |
| 5 µl | DNA library | |
| Total: 30 µl |
Gently mix the library by gentle flicking and inversion immediately before loading 30 μL via the priming port. Begin sequencing using the default settings for 24 hours.
30m
Loading of library (full flow cell)
30m
Allow the flow cell to reach room temperature for 20-30 mins ahead of loading of the library. Insert the flow cell into the MinION sequencer and check the number of active pores available. Visually check the flow cell for air bubbles. Removing a small (<20 ul) amount of yellow buffer to purge any air.
Thaw the sequencing buffer (SQB), loading beads (LB), flush tether (FLT) and a full tube of flush buffer (FB) at room temperature, briefly vortex all but the FLT and spin them down. To the full FB tube, add 30 µl FLT, and mix by gentle flicking and inversion. Of this, add 800 µl to the flow cell via the priming port, with caution to avoid the introduction of air bubbles.
Ensure the beads of the LB are suspended by vortexing again immediately before pipetting, then assemble the following solution:
| A | B | |
| Volume | Reagent | |
| 25.5 µl | Loading Beads (LB) | |
| 34 µl | Sequencing Buffer (SQB) | |
| 4.5 µl | Nuclease-free water | |
| 10 µl | DNA library | |
| Total: 75 µl |
This was gently mix the library by gentle flicking and inversion, load 75 µl via the SpotON sample port in a dropwise manner, ensuring each drop flows into the port before adding the next. The flow cell can then begin sequencing using the default settings.
30m