A streamlined protocol for high-throughput amplification-based analysis of DNA samples via nanopore sequencing (based on the 96-well PCR barcoding kit)

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.

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.

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

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

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

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

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.

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.

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.

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.

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

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.

Measure the concentration of your samples using Qubit high-sensitivity/broad-range (depending on expected concentration) dsDNA assays (or similar).

Pool your barcoded samples together based on concentration to achieve approximate equimolarity.

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.

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

Incubate the sample as follows:

A	B
Temperature	Time
20 °C	5 mins
65 °C	5 mins

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 .

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

Incubate at room temperature (~20 °C) for 10 mins.

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

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:

A	B
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:

A	B
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.

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.