This protocol describes how to carry out sequencing of a DNA sample using the Ligation Sequencing Kit V14 (SQK-LSK114) to generate high duplex rates. The library preparation is the same as the standard ligation sequencing protocol where DNA is repaired and end-prepped before the sequencing adapters are ligated onto both ends of the DNA. However, a different third-party repair buffer and ligase are recommended, and a higher concentration of successfully prepared library is loaded on the R10 Version HD flow cell for optimum duplex read capture. We have also included detailed instructions for performing duplex basecalling and calculating duplex rates.

To generate high duplex output, it is important to follow the library preparation protocol to ensure successful ligation of sequencing adapters onto both ends of the DNA strands.

Steps in the sequencing workflow:

Prepare for your experiment

You will need to:

• Extract your DNA, and check its length, quantity and purity. The quality checks performed during the protocol are essential in ensuring experimental success.
• Ensure you have your sequencing kit, the correct equipment and third-party reagents
• Download the software for acquiring and analysing your data
• Check your flow cell to ensure it has enough pores for a good sequencing run

Library preparation

Library preparation	Process	Time	Stop option
DNA repair and end-prep	Repair the DNA and prepare the end for adapter attachment and perform a bead clean-up	35 minutes	4°C overnight
Adapter ligation and clean-up	Ligate the sequencing adapters to the DNA ends and perform a bead clean-up.	20 minutes	4°C short-term storage or for repeated use, such as re-loading your flow cell. -80°C for single-use, long-term storage.
Priming and loading the flow cell	Prime the flow cell and load the prepared library for sequencing	5 minutes

Sequencing and analysis

You will need to:

• Start a sequencing run using the MinKNOW software which will perform duplex basecalling from the device.

The NEBNext® FFPE DNA Repair Buffer v2 can now be purchased as a single item through the Nanopore store.

The FFPE DNA Repair Buffer v2 (EXP-NEBFFPEV2) contains enough reagents for 24 reactions.

FFPE DNA Repair Buffer v2 (EXP-NEBFFPEV2) contents:

Name	Acronym	Cap colour	No. of vials	Fill volume per vial (μl)
NEBNext® FFPE DNA Repair Buffer V2	FFPE DNA Repair Buffer V2	Lilac	1	168

The additional NEBNext reagents required for this protocol:

• Salt-T4® DNA Ligase (NEB, M0467)

We have validated and recommend the use of all the third-party reagents used in this protocol. Alternatives have not been tested by Oxford Nanopore Technologies.

For all third-party reagents, we recommend following the manufacturer's instructions to prepare the reagents for use.

We recommend performing fragmentation and size selection prior to library prep to make samples more uniform in size to improve ligation of sequencing adapters and in turn, duplex output. By depleting short fragments (<25 kb) by size selection and decreasing very long fragments (>100 kb) by fragmentation, high pore occupancy can be attained as there will be fewer open pores and very long reads that can cause terminal blocking.

For more information on fragmentation and size selection, please see the linked pages.

To ensure you have enough DNA to load on a flow cell to reach high pore occupancy and output, yield from the library prep can be improved by increasing the waiting times of the clean-up steps with AMPure XP Beads (AXP) from five to 15 minutes. Pore occupancy can be viewed in MinKNOW after sequencing has commenced.

How to QC your input DNA

It is important that the input DNA meets the quantity and quality requirements. Using too little or too much DNA, or DNA of poor quality (e.g. highly fragmented or containing RNA or chemical contaminants) can affect your library preparation.

For instructions on how to perform quality control of your DNA sample, please read the Input DNA/RNA QC protocol.

Chemical contaminants

Depending on how the DNA is extracted from the raw sample, certain chemical contaminants may remain in the purified DNA, which can affect library preparation efficiency and sequencing quality. Read more about contaminants on the Contaminants page of the Community.

Note: We are in the process of reformatting our kits with single-use tubes into a bottle format.

Single-use tubes format:

Bottle format:

Note: This Product Contains AMPure XP Reagent Manufactured by Beckman Coulter, Inc. and can be stored at -20°C with the kit without detriment to reagent stability.

Note: The DNA Control Sample (DCS) is a 3.6 kb standard amplicon mapping the 3' end of the Lambda genome.

Duplex basecalling is the sequencing of both DNA strands and the consensus basecall for both strands leads to a further increase in accuracy of up to Q30 with the super-accurate (SUP) basecaller model.

We recommend performing duplex basecalling on MinKNOW (5.8) when you are setting up your sequencing run, as described below.

To generate high duplex output, it is important to follow the library preparation protocol to ensure successful ligation of sequencing adapters to both ends of the DNA strands and to perform duplex basecalling with the super-accurate (SUP) basecaller.

Once you have loaded your flow cell, the sequencing run can be started on MinKNOW, our sequencing software that controls the device, data acquisition and real-time basecalling. For more detailed information on setting up and using MinKNOW, please see the MinKNOW protocol.

MinKNOW can be used and set up to sequence in multiple ways:

• On a computer either direcly or remotely connected to a sequencing device.
• Directly on a MinION Mk1C, GridION and PromethION 24/48 sequencing device.

For more information on using MinKNOW on a sequencing device, please see the device user manuals:

• MinION Mk1B user manual
• MinION Mk1C user manual
• MinION Mk1D user manual
• GridION user manual
• PromethION user manual
• PromethION 2 Solo user manual

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To start a sequencing run on MinKNOW:

1. Navigate to the start page and click Start sequencing.

2. Fill in your experiment details, such as name and flow cell position, flow cell type (FLO-MIN114HD or FLO-PRO114HD) and sample ID.

3. Select the Ligation Sequencing Kit V14 (SQK-LSK114).

4. Keep run options to their default settings for run limit and minimum read length.

5. Set up basecalling and barcoding using the following parameters:

• Toggle the basecalling and duplex switches to ON.
• Next to "Models", click Edit options and choose Super-accurate basecaller (SUP) from the drop-down menu.
• Keep all other options at their default settings.
• Click Continue to output and continue.

6. Keep the output format and filtering options to their default settings.

• Click Continue to final review to continue.

7. Click Start on the Review page to start the sequencing run.

Duplex output can be viewed during a sequencing run via the MinKNOW UI as Gb of data in a duplex pair:

Duplex Q score can also be viewed on the UI during sequencing:

To determine total percentage of duplex reads, we calculate as follows:

= ((template + complement)/total) * 100

We expect a duplex output of ~60-70% of all bases which are part of a duplex pair with our R10 Version HD Flow Cells. For example:

= ((40 + 40)/100) * 100 = 80% of all bases part of a pair

• Duplex output = 40%
• Simplex output = 20%

This graph illustrates the duplex output presented as template and complement strands to highlight the need to combine the duplex rates. In this example, duplex rate is 40%.

Calculating output

Separating files

Note: Ensure your samtools version has filter expressions (1.11+)

To count bases for calculating duplex rates, the simplex and duplex must be separated into separate BAM files by using the following samtools command:

samtools view -@4 -e '[dx] == 1' $BAM -o duplex.bam -U simplex.bam

Counting

Once the simplex and duplex bases have been separated, these can be counted:

For example: samtools stats simplex.bam | grep ^SN | grep "total length" SN total length: 1467425598 # ignores clipping

samtools stats duplex.bam | grep ^SN | grep "total length" SN total length: 394815325 # ignores clipping

Total simplex	All duplex	All duplex / total simplex X 100	All duplex X 2 / total simplex X 100
1467425598	394815325	27%	54%

Finding IDs

The separated simplex and duplex .BAM files can be used to find their IDs, as follows:

• duplex.bam (semi-colon separated): samtools view duplex.bam | cut -f 1 | uniq | sed 's/;/\n/' | sort | uniq > duplex_ids.txt

• simplex.bam (single ID): samtools view simplex.bam | cut -f 1 | uniq | sort | uniq > simplex_ids.txt

Finding simplex-only data

As of Dorado v0.3.0, all simplex data (paired and unpaired) will be present in the uBAM, together with the duplex data. For downstream applications, it can be useful to access the unpaired simplex data.

To separate out the unpaired simplex data:

1. Find the IDs unique to the simplex data
2. Filter the original BAM using the simplex-only IDs

Find the IDs unique to the simplex data, as follows.

• duplex.bam (semi-colon separated): samtools view duplex.bam | cut -f 1 | uniq | sed 's/;/\n/' | sort | uniq > duplex_ids.txt

• simplex.bam (single ID): samtools view simplex.bam | cut -f 1 | uniq | sort | uniq > simplex_ids.txt

Filter the original BAM using the simplex-only IDs, as follows.

samtools view -N unique_to_simplex_ids.txt -o unique_to_simplex.bam simplex.bam

In addition, the duplex yield can also be measured in a different way by comparing the amount of data unique to simplex and the amount of duplex data using samtools stats:

samtools stats mybam.bam | grep ^SN

For example:

samtools stats unique_to_simplex.bam | grep ^SN SN total length: 689315580 # ignores clipping

samtools stats duplex.bam | grep ^SN SN total length: 394733687 # ignores clipping

394 mb X 2 = 788 mb which are part of a pair = 788/(788+689) = 53% total duplex

After sequencing and basecalling, the data can be analysed. For further information about options for basecalling and post-basecalling analysis, please refer to the Data Analysis document.

In the Downstream analysis section, we outline further options for analysing your data.

There are several options for further analysing your basecalled data:

1. EPI2ME workflows

For in-depth data analysis, Oxford Nanopore Technologies offers a range of bioinformatics tutorials and workflows available in EPI2ME, which are available in the EPI2ME section of the Community. The platform provides a vehicle where workflows deposited in GitHub by our Research and Applications teams can be showcased with descriptive texts, functional bioinformatics code and example data.

2. Research analysis tools

Oxford Nanopore Technologies' Research division has created a number of analysis tools, that are available in the Oxford Nanopore GitHub repository. The tools are aimed at advanced users, and contain instructions for how to install and run the software. They are provided as-is, with minimal support.

3. Community-developed analysis tools

If a data analysis method for your research question is not provided in any of the resources above, please refer to the resource centre and search for bioinformatics tools for your application. Numerous members of the Nanopore Community have developed their own tools and pipelines for analysing nanopore sequencing data, most of which are available on GitHub. Please be aware that these tools are not supported by Oxford Nanopore Technologies, and are not guaranteed to be compatible with the latest chemistry/software configuration.

We also have an FAQ section available on the Nanopore Community Support section.

If you have tried our suggested solutions and the issue still persists, please contact Technical Support via email (support@nanoporetech.com) or via LiveChat in the Nanopore Community.

We also have an FAQ section available on the Nanopore Community Support section.

If you have tried our suggested solutions and the issue still persists, please contact Technical Support via email (support@nanoporetech.com) or via LiveChat in the Nanopore Community.