PCR tiling of SARS-CoV-2 virus - automated Hamilton NGS STAR 96 (SQK-LSK109 with EXP-NBD196)

Version for device: MinION

1. Important This is a Legacy product

   This kit is available on the Legacy page of the store. We are in the process of gathering data to support the upgrade of this protocol to our latest chemistry. Further information regarding protocol upgrades will be provided on the Community as soon as they are available over the next few months. For further information on please see the product update page.

2. Introduction to the protocol

   To enable the support for the rapidly expanding user requests, the team at Oxford Nanopore Technologies have put together a partially automated end-to-end workflow based on the ARTIC Network protocols and analysis methods.

   We have developed this automated protocol on the Hamilton NGS Star 96 liquid handling robot. The protocol begins with manual preparation of the RNA samples, which includes reverse transcription and tiled PCR. The remainder of the library preparation is automated with minimal hands-on time which is required for sample quantification and deck re-loading.

   While this protocol is available in the Nanopore Community, we kindly ask users to ensure they are citing the members of the ARTIC network who have been behind the development of these methods.

   This protocol is based on the ARTIC amplicon sequencing protocol for MinION for nCoV-2019 by Josh Quick. The protocol generates 400 bp amplicons in a tiled fashion across the whole SARS-CoV-2 genome. Some example data is shown in the Downstream analysis and expected results section, this is generated using human coronavirus 229E to show what would be expected when running this protocol with SARS-CoV-2 samples.

   Primers were designed by Josh Quick using Primal Scheme; the primer sequences can be found here.

   Steps in the sequencing workflow:

   Prepare for your experiment
   you will need to:

   • Extract your RNA
   • 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

   Prepare your library
   You will need to:

   Off-deck:

   • Reverse transcribe your RNA samples with random hexamers
   • Amplify the samples by tiled PCR using separate primer pools
   • Combine the primer pools

   Note: On the Hamilton software, each step of the protocol is set up as a process. Currently, the off-deck section comprises Processes 1-2.

   On-deck:

   • Process 3: Purify and quantify the PCR products
   • Processes 4-8: Prepare the DNA ends for adapter attachment and ligate the native barcodes supplied in the kit to the DNA ends and pool the samples
   • Processes 10-11: Ligate the sequencing adapters supplied in the kit to the DNA ends

   Off-deck:

   • Prime the flow cell and load your DNA library into the flow cell

   
   Note: Timings are dependent on number of samples and include hands on time, such as deck loading and sample quantification

   Sequencing and analysis
   You will need to:

   • Start a sequencing run using the MinKNOW software, which will collect raw data from the device and convert it into basecalled reads
   • Start the EPI2ME software and select the barcoding workflow
3. Timings

   Off-deck library preparation:

   Process	X96 samples
   Reverse transcription	~20 minutes
   PCR tiling	~260 minutes

   Note: Manual preparation timings are for X96 samples. However, PCR tiling can take between ~200-260 minutes depending on the number of samples prepared.

   On-deck library preparation:

   Process	X24 samples	X48 samples	X96 samples	Hands-on time
   Deck set-up				~30 minutes
   Process 3: PCR tiling clean-up	~53 minutes	~60 minutes	~60 minutes
   Quantification				~20 minutes
   Deck loading				~30 minutes
   Processes 4-8: End-prep and native barcode ligation	~200 minutes	~240 minutes	~280 minutes
   Quantification				~10 minutes
   Deck loading				~30 minutes
   Processes 10-11: Adapter ligation and clean-up	~90 minutes	~90 minutes	~90 minutes
   Quantification				~10 minutes
   Total	5 hours 43 minutes	6 hours 30 minutes	7 hours	2 hours 10 minutes

4. Before starting

   This protocol requires total RNA extracted from samples that have been screened by a suitable qPCR assay. Here we demonstrate the level of sensitivity and specificity by titrating total RNA extracted from cell culture infected with Human coronavirus 229E spiked into 100 ng human RNA extracted from GM12878 to give approximate figures.

   Although not tested here, work performed by Josh Quick et al. on the Zika virus gives approximate dilution factors that may help reduction of inhibiting compounds that can be co-extracted from samples.

   Note: this is a guideline and not currently tested for SARS-CoV-2.

   qPCR ct	Dilution factor
   18–35	none
   15–18	1:10
   12–15	1:100

   When processing multiple samples at once, we recommend making master mixes with an additional 10% of the volume. We also recommend using pre- and post-PCR hoods when handling master mixes and samples. It is important to clean and/or UV irradiate these hoods between sample batches. Furthermore, to track and monitor cross-contamination events, it is important to run a negative control reaction at the reverse transcription stage using nuclease-free water instead of sample, and carrying this control through the rest of the prep.

   To minimise the chance of pipetting errors when preparing primer mixes, we recommend ordering the tiling primers from IDT in a lab-ready format at 100 µM.

5. Important Compatibility of this protocol

   This protocol should only be used in combination with:

   • Ligation Sequencing Kit (SQK-LSK109)
   • Native Barcoding Expansion 96 (EXP-NBD196)
   • FLO-MIN106 (R9.4.1) flow cells
   • Flow Cell Wash Kit (EXP-WSH004)

1. Input RNA guidelines

   Where sample RNA is added to the below reaction, it is likely advantageous to follow the dilution guidelines proposed by Josh Quick:

   qPCR Ct	Dilution factor
   18–35	none
   15–18	1:10
   12–15	1:100

   If the sample has a low copy number (ct 18–35) use up to 16 µl of sample. Use nuclease-free water to make up any remaining volume. Take note to be aware that co-extracted compounds may inhibit reverse transcription and PCR.

2. Input File Worklist

   Two worklist input excel files are required prior to running the protocol on the Hamilton NGS Star: one for processes03 to 08 and another for processes10 to 11. These will contain information regarding the appropriate number of samples, well identifiers and source concentration. In the first worklist, the target well is the well for pooled samples at the end of Process07. A maximum of 24 samples should be combined to make each pool, so for 96 samples, there will be 4 pools in target wells A1 to D1. Samples should be split evenly, so that the pools contain equivalent volumes. This worklist must be updated after the automated library preparation sections with the relevant plate information. Note that the pools are combined at the end of process08 into well A2, so the Source_Well for the second worklist will be A2.

   Example:
   

3. Hamilton NGS Star 96 and deck layout

   This method has been tested and validated using the Hamilton NGS Star 96 (with 8 channels and MPH96), including an on-deck thermal cycler (ODTC), a Hamilton Heater Shaker (HHS) and the Inheco Cold Plate Air Cooled (CPAC) Modules. All modules are used at different stages of the protocol. This protocol may require some fine tuning for the specific NGS Star set-up and the temperature/humidity of the customer laboratory.

   Please contact your Hamilton representative for further details.

   Deck layout

   

   • MIDI plates: Abgene™ 96 Well 0.8mL Polypropylene Deepwell Storage Plate (ThermoFisher, Cat # AB0859)
   • HSP plates: Bio-Rad Hard-Shell® 96-Well PCR Plates (Cat# HSP9601)
   • ODTC: On-Deck Thermal Cycler Module
   • HHS: Hamilton Heater Shaker Module
   • CPAC: Inheco Cold Plate Air Cooled Module
   • 20 ml reagent troughs: X150 Reagent tub MagNA Pure LC medium 20 plastic (zigzag troughs, Roche #03004058001
   • 60 ml self-standing large troughs (Hamilton, 56694-01)
4. Consumables and reagent quantities required:

   Consumables	X24 samples	X48 samples	X96 samples
   Hamilton 50 µl CO-RE tips with filter	293	561	1097
   Hamilton 300 µl CO-RE tips with filter	108	206	402
   Hamilton 1000 µl CO-RE tips with filter	91	109	145
   Hamilton 60 ml Reagent Reservoir, Self-Standing with Lid	5	5	5
   Hamilton PCR ComfortLid	2	2	2
   Bio-Rad Hard-Shell® 96-Well PCR Plate	10	10	10
   Roche Diagnostics MagNA Pure LC Medium Reagent Tubs 20	5	5	5
   Sarstedt Inc Screw Cap Micro Tube 2ml	4	6	8
   Abgene™ 96 Well 0.8mL Polypropylene Deepwell Storage Plate	4	4	4
   1.5 ml Eppendorf DNA LoBind tubes	4	4	4

   Reagents/kits	X24 samples	X48 samples	X96 samples
   AMPure XP Beads	7.639 ml	9.256 ml	12.422 ml
   80% ethanol	24.6 ml	35.3 ml	56.4 ml
   Nuclease-free water	16 ml	16.7 ml	19.9 ml
   NEBNext Ultra II End Repair/dA-Tailing Module (Cat# E7546)	1 small kit	2 small kits	2 small kits
   NEB Blunt/TA Ligase Master Mix (M0367)	2 small kits	3 small kits	1 large kit
   NEBNext Quick Ligation Module (Cat# E6056)	1 small kit	1 small kit	1 small kit
   Native Barcoding Expansion 96 (EXP-NBD196)	1 kit	1 kit	1 kit
   Ligation Sequencing Kit (SQK-LSK109)	1 kit	1 kit	1 kit
   SFB Expansion (EXP-SFB001)	1 kit (3 tubes)	1 kit (3 tubes)	1 kit (4 tubes)
   LunaScript™ RT SuperMix Kit	1 small kit	2 small kits	1 large kit
   Q5® Hot Start High-Fidelity 2X Master Mix (NEB, M0494)	1 small kit	1 small kit	1 small kit

   Note: These quantities are for one run of the protocol for the selected number of samples.

5. Native Barcoding Expansion 96 (EXP-NBD196) contents

   Kits in batches NBD196.10.0007 onwards have barcodes ordered in columns on the plate:

   

   Kits in batches prior to NBD196.10.0007 have barcodes ordered in rows:

   

   Name	Acronym	Cap colour	No. of vials	Fill volume per vial (μl)
   Native Barcode 01-96	NB01-96	-	1 plate	40 μl per well
   Adapter Mix II	AMII	Green	1	70

6. Ligation Sequencing Kit contents (SQK-LSK109)

   

   Name	Acronym	Cap colour	No. of vials	Fill volume per vial (µl)
   DNA CS	DCS	Yellow	1	50
   Adapter Mix	AMX	Green	1	40
   Ligation Buffer	LNB	Clear	1	200
   L Fragment Buffer	LFB	White cap, orange stripe on label	2	1,800
   S Fragment Buffer	SFB	Grey	2	1,800
   Sequencing Buffer	SQB	Red	2	300
   Elution Buffer	EB	Black	1	200
   Loading Beads	LB	Pink	1	360

7. Flow Cell Priming Kit contents (EXP-FLP002)

   

   Name	Acronym	Cap colour	No. of vials	Fill volume per vial (μl)
   Flush Buffer	FB	Blue	6	1,170
   Flush Tether	FLT	Purple	1	200

8. SFB Expansion contents (EXP-SFB001)

   

   Name	Acronym	Cap colour	No. of vials	Fill volume per vial (μl)
   Short Fragment Buffer	SFB	Grey	4	1,800

9. Native barcode sequences

   Component	Forward sequence	Reverse sequence
   NB01	CACAAAGACACCGACAACTTTCTT	AAGAAAGTTGTCGGTGTCTTTGTG
   NB02	ACAGACGACTACAAACGGAATCGA	TCGATTCCGTTTGTAGTCGTCTGT
   NB03	CCTGGTAACTGGGACACAAGACTC	GAGTCTTGTGTCCCAGTTACCAGG
   NB04	TAGGGAAACACGATAGAATCCGAA	TTCGGATTCTATCGTGTTTCCCTA
   NB05	AAGGTTACACAAACCCTGGACAAG	CTTGTCCAGGGTTTGTGTAACCTT
   NB06	GACTACTTTCTGCCTTTGCGAGAA	TTCTCGCAAAGGCAGAAAGTAGTC
   NB07	AAGGATTCATTCCCACGGTAACAC	GTGTTACCGTGGGAATGAATCCTT
   NB08	ACGTAACTTGGTTTGTTCCCTGAA	TTCAGGGAACAAACCAAGTTACGT
   NB09	AACCAAGACTCGCTGTGCCTAGTT	AACTAGGCACAGCGAGTCTTGGTT
   NB10	GAGAGGACAAAGGTTTCAACGCTT	AAGCGTTGAAACCTTTGTCCTCTC
   NB11	TCCATTCCCTCCGATAGATGAAAC	GTTTCATCTATCGGAGGGAATGGA
   NB12	TCCGATTCTGCTTCTTTCTACCTG	CAGGTAGAAAGAAGCAGAATCGGA
   NB13	AGAACGACTTCCATACTCGTGTGA	TCACACGAGTATGGAAGTCGTTCT
   NB14	AACGAGTCTCTTGGGACCCATAGA	TCTATGGGTCCCAAGAGACTCGTT
   NB15	AGGTCTACCTCGCTAACACCACTG	CAGTGGTGTTAGCGAGGTAGACCT
   NB16	CGTCAACTGACAGTGGTTCGTACT	AGTACGAACCACTGTCAGTTGACG
   NB17	ACCCTCCAGGAAAGTACCTCTGAT	ATCAGAGGTACTTTCCTGGAGGGT
   NB18	CCAAACCCAACAACCTAGATAGGC	GCCTATCTAGGTTGTTGGGTTTGG
   NB19	GTTCCTCGTGCAGTGTCAAGAGAT	ATCTCTTGACACTGCACGAGGAAC
   NB20	TTGCGTCCTGTTACGAGAACTCAT	ATGAGTTCTCGTAACAGGACGCAA
   NB21	GAGCCTCTCATTGTCCGTTCTCTA	TAGAGAACGGACAATGAGAGGCTC
   NB22	ACCACTGCCATGTATCAAAGTACG	CGTACTTTGATACATGGCAGTGGT
   NB23	CTTACTACCCAGTGAACCTCCTCG	CGAGGAGGTTCACTGGGTAGTAAG
   NB24	GCATAGTTCTGCATGATGGGTTAG	CTAACCCATCATGCAGAACTATGC
   NB25	GTAAGTTGGGTATGCAACGCAATG	CATTGCGTTGCATACCCAACTTAC
   NB26	CATACAGCGACTACGCATTCTCAT	ATGAGAATGCGTAGTCGCTGTATG
   NB27	CGACGGTTAGATTCACCTCTTACA	TGTAAGAGGTGAATCTAACCGTCG
   NB28	TGAAACCTAAGAAGGCACCGTATC	GATACGGTGCCTTCTTAGGTTTCA
   NB29	CTAGACACCTTGGGTTGACAGACC	GGTCTGTCAACCCAAGGTGTCTAG
   NB30	TCAGTGAGGATCTACTTCGACCCA	TGGGTCGAAGTAGATCCTCACTGA
   NB31	TGCGTACAGCAATCAGTTACATTG	CAATGTAACTGATTGCTGTACGCA
   NB32	CCAGTAGAAGTCCGACAACGTCAT	ATGACGTTGTCGGACTTCTACTGG
   NB33	CAGACTTGGTACGGTTGGGTAACT	AGTTACCCAACCGTACCAAGTCTG
   NB34	GGACGAAGAACTCAAGTCAAAGGC	GCCTTTGACTTGAGTTCTTCGTCC
   NB35	CTACTTACGAAGCTGAGGGACTGC	GCAGTCCCTCAGCTTCGTAAGTAG
   NB36	ATGTCCCAGTTAGAGGAGGAAACA	TGTTTCCTCCTCTAACTGGGACAT
   NB37	GCTTGCGATTGATGCTTAGTATCA	TGATACTAAGCATCAATCGCAAGC
   NB38	ACCACAGGAGGACGATACAGAGAA	TTCTCTGTATCGTCCTCCTGTGGT
   NB39	CCACAGTGTCAACTAGAGCCTCTC	GAGAGGCTCTAGTTGACACTGTGG
   NB40	TAGTTTGGATGACCAAGGATAGCC	GGCTATCCTTGGTCATCCAAACTA
   NB41	GGAGTTCGTCCAGAGAAGTACACG	CGTGTACTTCTCTGGACGAACTCC
   NB42	CTACGTGTAAGGCATACCTGCCAG	CTGGCAGGTATGCCTTACACGTAG
   NB43	CTTTCGTTGTTGACTCGACGGTAG	CTACCGTCGAGTCAACAACGAAAG
   NB44	AGTAGAAAGGGTTCCTTCCCACTC	GAGTGGGAAGGAACCCTTTCTACT
   NB45	GATCCAACAGAGATGCCTTCAGTG	CACTGAAGGCATCTCTGTTGGATC
   NB46	GCTGTGTTCCACTTCATTCTCCTG	CAGGAGAATGAAGTGGAACACAGC
   NB47	GTGCAACTTTCCCACAGGTAGTTC	GAACTACCTGTGGGAAAGTTGCAC
   NB48	CATCTGGAACGTGGTACACCTGTA	TACAGGTGTACCACGTTCCAGATG
   NB49	ACTGGTGCAGCTTTGAACATCTAG	CTAGATGTTCAAAGCTGCACCAGT
   NB50	ATGGACTTTGGTAACTTCCTGCGT	ACGCAGGAAGTTACCAAAGTCCAT
   NB51	GTTGAATGAGCCTACTGGGTCCTC	GAGGACCCAGTAGGCTCATTCAAC
   NB52	TGAGAGACAAGATTGTTCGTGGAC	GTCCACGAACAATCTTGTCTCTCA
   NB53	AGATTCAGACCGTCTCATGCAAAG	CTTTGCATGAGACGGTCTGAATCT
   NB54	CAAGAGCTTTGACTAAGGAGCATG	CATGCTCCTTAGTCAAAGCTCTTG
   NB55	TGGAAGATGAGACCCTGATCTACG	CGTAGATCAGGGTCTCATCTTCCA
   NB56	TCACTACTCAACAGGTGGCATGAA	TTCATGCCACCTGTTGAGTAGTGA
   NB57	GCTAGGTCAATCTCCTTCGGAAGT	ACTTCCGAAGGAGATTGACCTAGC
   NB58	CAGGTTACTCCTCCGTGAGTCTGA	TCAGACTCACGGAGGAGTAACCTG
   NB59	TCAATCAAGAAGGGAAAGCAAGGT	ACCTTGCTTTCCCTTCTTGATTGA
   NB60	CATGTTCAACCAAGGCTTCTATGG	CCATAGAAGCCTTGGTTGAACATG
   NB61	AGAGGGTACTATGTGCCTCAGCAC	GTGCTGAGGCACATAGTACCCTCT
   NB62	CACCCACACTTACTTCAGGACGTA	TACGTCCTGAAGTAAGTGTGGGTG
   NB63	TTCTGAAGTTCCTGGGTCTTGAAC	GTTCAAGACCCAGGAACTTCAGAA
   NB64	GACAGACACCGTTCATCGACTTTC	GAAAGTCGATGAACGGTGTCTGTC
   NB65	TTCTCAGTCTTCCTCCAGACAAGG	CCTTGTCTGGAGGAAGACTGAGAA
   NB66	CCGATCCTTGTGGCTTCTAACTTC	GAAGTTAGAAGCCACAAGGATCGG
   NB67	GTTTGTCATACTCGTGTGCTCACC	GGTGAGCACACGAGTATGACAAAC
   NB68	GAATCTAAGCAAACACGAAGGTGG	CCACCTTCGTGTTTGCTTAGATTC
   NB69	TACAGTCCGAGCCTCATGTGATCT	AGATCACATGAGGCTCGGACTGTA
   NB70	ACCGAGATCCTACGAATGGAGTGT	ACACTCCATTCGTAGGATCTCGGT
   NB71	CCTGGGAGCATCAGGTAGTAACAG	CTGTTACTACCTGATGCTCCCAGG
   NB72	TAGCTGACTGTCTTCCATACCGAC	GTCGGTATGGAAGACAGTCAGCTA
   NB73	AAGAAACAGGATGACAGAACCCTC	GAGGGTTCTGTCATCCTGTTTCTT
   NB74	TACAAGCATCCCAACACTTCCACT	AGTGGAAGTGTTGGGATGCTTGTA
   NB75	GACCATTGTGATGAACCCTGTTGT	ACAACAGGGTTCATCACAATGGTC
   NB76	ATGCTTGTTACATCAACCCTGGAC	GTCCAGGGTTGATGTAACAAGCAT
   NB77	CGACCTGTTTCTCAGGGATACAAC	GTTGTATCCCTGAGAAACAGGTCG
   NB78	AACAACCGAACCTTTGAATCAGAA	TTCTGATTCAAAGGTTCGGTTGTT
   NB79	TCTCGGAGATAGTTCTCACTGCTG	CAGCAGTGAGAACTATCTCCGAGA
   NB80	CGGATGAACATAGGATAGCGATTC	GAATCGCTATCCTATGTTCATCCG
   NB81	CCTCATCTTGTGAAGTTGTTTCGG	CCGAAACAACTTCACAAGATGAGG
   NB82	ACGGTATGTCGAGTTCCAGGACTA	TAGTCCTGGAACTCGACATACCGT
   NB83	TGGCTTGATCTAGGTAAGGTCGAA	TTCGACCTTACCTAGATCAAGCCA
   NB84	GTAGTGGACCTAGAACCTGTGCCA	TGGCACAGGTTCTAGGTCCACTAC
   NB85	AACGGAGGAGTTAGTTGGATGATC	GATCATCCAACTAACTCCTCCGTT
   NB86	AGGTGATCCCAACAAGCGTAAGTA	TACTTACGCTTGTTGGGATCACCT
   NB87	TACATGCTCCTGTTGTTAGGGAGG	CCTCCCTAACAACAGGAGCATGTA
   NB88	TCTTCTACTACCGATCCGAAGCAG	CTGCTTCGGATCGGTAGTAGAAGA
   NB89	ACAGCATCAATGTTTGGCTAGTTG	CAACTAGCCAAACATTGATGCTGT
   NB90	GATGTAGAGGGTACGGTTTGAGGC	GCCTCAAACCGTACCCTCTACATC
   NB91	GGCTCCATAGGAACTCACGCTACT	AGTAGCGTGAGTTCCTATGGAGCC
   NB92	TTGTGAGTGGAAAGATACAGGACC	GGTCCTGTATCTTTCCACTCACAA
   NB93	AGTTTCCATCACTTCAGACTTGGG	CCCAAGTCTGAAGTGATGGAAACT
   NB94	GATTGTCCTCAAACTGCCACCTAC	GTAGGTGGCAGTTTGAGGACAATC
   NB95	CCTGTCTGGAAGAAGAATGGACTT	AAGTCCATTCTTCTTCCAGACAGG
   NB96	CTGAACGGTCATAGAGTCCACCAT	ATGGTGGACTCTATGACCGTTCAG

1. MinION Mk1B IT requirements

   Sequencing on a MinION Mk1B requires a high-spec computer or laptop to keep up with the rate of data acquisition. For more information, refer to the MinION Mk1B IT requirements document.

2. MinION Mk1C IT requirements

   The MinION Mk1C contains fully-integrated compute and screen, removing the need for any accessories to generate and analyse nanopore data. For more information refer to the MinION Mk1C IT requirements document.

3. MinION Mk1D IT requirements

   Sequencing on a MinION Mk1D requires a high-spec computer or laptop to keep up with the rate of data acquisition. For more information, refer to the MinION Mk1D IT requirements document.

4. Software for nanopore sequencing

   MinKNOW

   The MinKNOW software controls the nanopore sequencing device, collects sequencing data and basecalls in real time. You will be using MinKNOW for every sequencing experiment to sequence, basecall and demultiplex if your samples were barcoded.

   For instructions on how to run the MinKNOW software, please refer to the MinKNOW protocol.

   EPI2ME (optional)

   The EPI2ME cloud-based platform performs further analysis of basecalled data, for example alignment to the Lambda genome, barcoding, or taxonomic classification. You will use the EPI2ME platform only if you would like further analysis of your data post-basecalling.

   For instructions on how to create an EPI2ME account and install the EPI2ME Desktop Agent, please refer to this link.

5. Check your flow cell

   We highly recommend that you check the number of pores in your flow cell prior to starting a sequencing experiment. This should be done within 12 weeks of purchasing for MinION/GridION/PromethION or within four weeks of purchasing Flongle Flow Cells. Oxford Nanopore Technologies will replace any flow cell with fewer than the number of pores in the table below, when the result is reported within two days of performing the flow cell check, and when the storage recommendations have been followed. To do the flow cell check, please follow the instructions in the Flow Cell Check document.

   Flow cell	Minimum number of active pores covered by warranty
   Flongle Flow Cell	50
   MinION/GridION Flow Cell	800
   PromethION Flow Cell	5000

1. Consumables and reagent quantities:

   Consumables	X24, X48 and X96 samples
   Hard-Shell® 96-Well PCR Plates	1

   Reagents	X24 samples	X48 samples	X96 samples
   LunaScript™ RT SuperMix (5x)	96 µl	192 µl	384 µl

2. Important Keep the RNA sample on ice as much as possible to prevent nucleolytic degradation, which may affect sensitivity.
3. In a clean pre-PCR hood, mix together the following components in each well of a 96-well plate on ice or in a PCR cool rack, such as the Eppendorf PCR-Cooler:

   Reagent	Volume per well
   RNA sample	16 µl
   LunaScript RT SuperMix (5x)	4 µl
   Total	20 µl

   Note: We recommend using up to 16 µl of RNA sample. Use nuclease-free water to make up the final volume to 16 µl if required.

4. Mix gently by pipetting, and spin down. Return the plate to ice.
5. Preheat the thermal cycler to 25°C.
6. Incubate the samples in the thermal cycler using the following program:

   Step	Temperature	Time	Cycles
   Primer annealing	25°C	2 min	1
   cDNA synthesis	55°C	10 min	1
   Heat inactivation	95°C	1 min	1
   Hold	4°C	∞

7. End of Step While the reverse transcription reaction is running, prepare the primer pools as described in the next section.

1. Consumable and reagent quantities:

   Consumables	X24, X48 and X96 samples
   1.5 ml Eppendorf tubes	4
   Bio-Rad Hard-Shell® 96-Well PCR Plates	3

   Reagents	X24 samples	X48 samples	X96 samples
   Q5® Hot Start High-Fidelity 2X Master Mix	300 µl	600 µl	1200 µl
   Primer pool at 10 µM (A or B)	88.8 µl	177.6 µl	355.2 µl
   Nuclease-free water	91.2 µl	182.4 µl	364.8 µl

2. Primer design

   To generate tiled PCR amplicons from the SARS-CoV-2 viral cDNA, primers were designed by Josh Quick using Primal Scheme. These primers are designed to generate 400 bp amplicons that overlap by approximately 20 bp. These primer sequences can be found here. Where we show example data outputs in this protocol, the same parameters were used to design primers to the human coronavirus 229E to provide guideline statistics.

3. Important We recommend ordering the required primers from IDT in a lab-ready format at 100 µM. However, if primers have been ordered lyophilised, they should be resuspended in water or low-EDTA TE buffer to a final concentration of 100 µM.
4. Important We recommend handling the primer stocks and derivatives in a clean template-free PCR hood.
5. If primers are supplied individually, add 5 µl of each primer from pool A per sample to a 1.5 ml Eppendorf DNA LoBind tube to give a 100 µM stock primer pool.
6. If primers are supplied individually, add 5 µl of each primer from pool B per sample to a 1.5 ml Eppendorf DNA LoBind tube to give a 100 µM stock primer pool.
7. Dilute each 100 µM stock 1 in 10 with nuclease-free water to form a working stock of each pool at 10 µM.

   Note: To achieve the desired final concentration of each primer in the pool at 0.015 µM in the PCR reaction, 3.7 µl of the 10 µM working stock is needed for each PCR reaction. Two separate PCR reactions will be performed per sample, one for pool A primers and one for pool B. This results in tiled amplicons that have approximately 20 bp overlap.

8. In a clean pre-PCR hood, set up two individual reactions for primer pool A and primer pool B in clean 5 ml centrifuge tubes:

   Reagent	X24 samples	X48 samples	X96 samples
   Q5® Hot Start High-Fidelity 2X Master Mix	300 µl	600 µl	1200 µl
   Primer pool at 10 µM (A or B)	88.8 µl	177.6 µl	355.2 µl
   Nuclease-free water	91.2 µl	182.4 µl	364.8 µl
   Total	480 µl	960 µl	1920 µl

9. In a clean 96-well plate, aliquot 20 µl of pool A reaction to each well per sample. Repeat in a new 96-well plate with pool B reaction. Add 5 µl of the reverse-transcribed samples per well in both plates.
10. Important Carry forward the negative control from the reverse transcription reaction to monitor cross-contamination events.

    We recommend having a single negative for every plate of samples and a standard curve of positive controls.

11. Mix well by pipetting and spin down in a centrifuge.
12. Incubate using the following program, with the heated lid set to 105°C:

    Step	Temperature	Time	Cycles
    Initial denaturation	98°C	30 sec	1
    Denaturation Annealing and extension	98°C 65°C	15 sec 5 min	25–35
    Hold	4°C	∞

    Note: Cycle number should be varied for low or high viral load samples. Guidelines provided by Josh Quick suggest that 25 cycles should be used for Ct 18–21 up to a maximum of 35 cycles for Ct 35, however this has not been tested here.

13. Important If available, a clean post-PCR hood should be used for all steps that involve handling amplified material. Decontamination with UV and or DNAzap between sample batches is recommended.
14. Combine the 25 µl reaction from pool A and the 25 µl reaction from pool B per sample, into a clean hard-shell fully skirted PCR plate to load on the NGS Star; one well per sample.

    Note: Ensure the sample from pool A corresponds to the sample from pool B.

15. End of Step Take forward the plate into the automated clean-up step.

1. Consumables and equipment quantities:

   Consumables	X24 samples	X48 samples	X96 samples
   Hamilton 50 µl CO-RE tips with filter	72	144	288
   Hamilton 300 µl CO-RE tips with filter	104	200	392
   Hamilton 1000 µl CO-RE tips with filter	24	24	24
   Hamilton 60 ml Reagent Reservoir, Self-Standing with Lid	2	2	2
   Bio-Rad Hard-Shell® 96-Well PCR Plate	2	2	2
   Roche Diagnostics MagNA Pure LC Medium Reagent Tubs 20	1	1	1
   Abgene™ 96 Well 0.8mL Polypropylene Deepwell Storage Plate	1	1	1

2. Reagent quantities:

   Reagents	X24 samples	X48 samples	X96 samples
   AMPure XP Beads	3.3 ml	4.6 ml	7.2 ml
   80% ethanol	17.5 ml	28.1 ml	49.2 ml
   Nuclease-free water	7.4 ml	7.7 ml	8.5 ml

3. Switch on the Hamilton NGS STAR 96 robot and open 'Hamilton Run Control' on the computer by clicking the icon:

   

4. Select 'File' and 'Open' to choose the method to run: PCRtilingLSK109_v0.8.3.4MPH.med.

   After a couple minutes, the software will connect to the instrument and the deck layout should appear.

   Note: Ensure the Trace View and MLSTAR deck layout are selected at the top of the page with the instrument and the HHS, CPAC and ODTC are connected.

   

5. Important Ensure the instrument is set to 'Instrument' mode and highlighted green rather than 'Simulation' mode.

   

6. Tip The ODTC and CPAC are not required for this automated step and can be switched off.
7. From Run Control, click the green play button to begin setting up the deck.

   

8. Select 'Process03: Tiling PCR Cleanup' as the start process.

   

9. Select 'Process03: Tiling PCR Cleanup' as the Stop Process.

   

10. Click 'Browse' to upload the Input File Worklist for the specific samples with the relevant information in the run and click 'OK'.

    

    Relevant information required includes sample identification, well location and target well location.

    Example sample sheet:
    

11. Insert plates in their corresponding positions displayed on screen. Click 'Ok' to continue.

    

12. Load a full deck of 50 µl tips into the positions on screen. Click 'Ok' to continue.

    

13. Update the first and last 50 µl tips available to use on the 'Edit Tip Count' window and click 'Ok' to continue.

    Note: Click 'Remove All' twice if all the positions are not brown and update the tips available. There must be a tip rack (empty, partial or full) in every position. All tips in the tip racks also need to be input into the GUI to prevent a hardware crash.

    

14. Load a full deck of 300 µl tips in the positions on screen. Click 'Ok' to continue.

    

15. Update the first and last 300 µl tips available to use on the 'Edit Tip Count' window and click 'Ok' to continue.

    

16. Freshly prepare 80% ethanol in nuclease-free water.

    Reagents	X24 samples	X48 samples	X96 samples
    80% ethanol	17.5 ml	28.1 ml	49.2 ml

17. Insert the trough of 80% ethanol in the position on screen and click 'Ok' to continue.

    

18. Prepare the nuclease-free water and resuspend the AMPure XP beads by vortexing.

    Reagents	X24 samples	x48 samples	x96 samples
    Nuclease-free water	7.4 ml	7.7 ml	8.5 ml
    AMPure XP beads	3.3 ml	4.6 ml	7.2 ml

19. Important Ensure the AMPure XP beads are well mixed before use by vortexing.
20. Insert the troughs of AMPure XP beads and nuclease-free water in their positions on screen. Click 'Ok' to continue.

    

21. Load 1000 µl tips and insert the Tiling PCR Plate containing 50 µl DNA from the previous step into the positions on screen. Click 'Ok' to continue.

    

22. Update the first and last 1000 µl tips available to use in the 'Edit Tip Count' window and click 'Ok' to continue.

    

23. Click 'Ok' to start the run.

    

24. When the run is complete, unload the output plate. Click 'Ok' to continue.

    

25. Do not unload the carrier by clicking 'No'.

    

26. Click 'Ok' on the summary box to continue.

    

27. Quantify 1 µl of each eluted sample from the output plate using a Qubit fluorometer plate reader off deck.
28. Update the worklist with the measured concentration of purified PCR product.
29. End of Step Take forward the output plate containing the post PCR clean-up samples, into the next step.

1. Consumables and equipment quantities:

   Consumables	X24 samples	X48 samples	X96 samples
   Hamilton 50 µl CO-RE tips with filter	215	411	803
   Hamilton 300 µl CO-RE tips with filter	2	4	8
   Hamilton 1000 µl CO-RE tips with filter	42	60	96
   Hamilton 60 ml Reagent Reservoir, Self-Standing with Lid	2	2	2
   Hamilton PCR ComfortLid	1	1	1
   Bio-Rad Hard-Shell® 96-Well PCR Plate	3	3	3
   Roche Diagnostics MagNA Pure LC Medium Reagent Tubs 20	2	2	2
   Sarstedt Inc Screw Cap Micro Tube 2ml	2	4	6
   Abgene™ 96 Well 0.8mL Polypropylene Deepwell Storage Plate	2	2	2

2. Reagent quantities:

   Reagents	X24 samples	X48 samples	X96 samples
   NEBNext Ultra II End-prep reaction buffer	87.5 µl	175 µl	315 µl
   NEBNext Ultra II End-prep enzyme mix	37.5 µl	75 µl	135 µl
   NEBNext Blunt/TA Ligase Master Mix (M0367)	350 µl	700 µl	1350 µl
   AMPure XP beads	2317 µl	2633.6 µl	3.2 ml
   80% ethanol	7.1 ml	7.2 ml	7.2 ml
   Nuclease-free water	8.4 ml	8.8 ml	11 ml
   Short Fragment Buffer (SFB)	2550 µl	3.1 ml	4.2 ml

3. Important For optimal efficiency of the end-prep reaction, use 75 ng of cDNA from the previous step.
4. Important We recommended carrying the RT negative control through this step until sequencing.
5. Tip All connected modules are required for this step and must be switched on.
6. Prepare the NEBNext Ultra II End repair / dA-tailing Module reagents in accordance with manufacturer’s instructions, and place on ice.

   For optimal performance, NEB recommend the following:

   1. Thaw all reagents on ice.
   2. Flick and/or invert reagent tube to ensure they are well mixed.
   3. Always spin down tubes before opening for the first time each day.
   4. The Ultra II End prep buffer and FFPE DNA Repair buffer may have a little precipitate. Allow the mixture to come to room temperature and pipette the buffer up and down several times to break up the precipitate, followed by vortexing the tube for several seconds to ensure the reagent is thoroughly mixed.
   5. The FFPE DNA repair buffer may have a yellow tinge and is fine to use if yellow.

7. Thaw the native barcodes at room temperature, enough for one barcode per sample. Individually mix the barcodes by pipetting and load a minimum of 5 µl per well into a 96-well HSP plate. Store on ice.
8. Thaw the tube of Short Fragment Buffer (SFB) at room temperature, mix by vortexing, spin down and place on ice.
9. From Run Control, click the green play button to begin setting up the deck.

   

10. Select 'Process04: cDNA End Prep' to start.

    

11. Select 'Process08: Barcode Ligation Cleanup' as the Stop Process.

    

12. Click 'Browse' to upload the Input File Worklist with the updated sample concentrations and click 'OK'.

    

    Note: If fewer than 96 samples are to be processed, ensure the positions of the barcodes in the barcode plate match those in the worklist.

    

13. Select 'Input samples require normalisation'.

    

14. Fill in the normalisation settings as suggested below:

    

    Note: If samples fall outside the recommended concentrations, a warning will appear. Click 'Ok' to continue.
    

15. Prepare the End Prep Mastermix and Native Barcode Ligation Mastermix with the following reagents according to the Hamilton user interface. Select 'Yes' or 'No' to continue.

    Note: It is user preference whether to save and print the instructions.

    End-prep Mastermix:

    Reagent	X24 samples	X48 samples	X96 samples
    Ultra II End Prep Buffer	87.5 µl	175 µl	315 µl
    Ultra II End Prep Enzyme Mix	37.5 µl	75 µl	135 µl

    
    
    Native Barcode Mastermix:

    Reagent	X24 samples	X48 samples	X96 samples
    NEB Blunt/TA Ligase Master Mix	350 µl	700 µl	1350 µl

    

16. Important Ensure the mastermixes are well mixed and homogenous before loading the 2 ml Sarstedt tubes. Mixing in the robot is not effective.
17. Insert the ComfortLid Rest Position as displayed on screen. Click 'Ok' to continue.

    

18. Insert plates to their corresponding positions on screen. Click 'Ok' to continue.

    

19. Load a full deck of 50 µl tips into the positions on screen. Click 'Ok' to continue.

    

20. Select the first and last tip positions to use in the 'Edit Tip Count' window. Click 'Ok' to continue.

    

21. Load a full deck of 300 µl tips in the positions on screen. Click 'Ok' to continue.

    

22. Select the first and last tip positions to use in the 'Edit Tip Count' window. Click 'Ok' to continue.

    

23. Freshly prepare 80% ethanol in nuclease-free water.

    Reagents	X24 samples	X48 samples	X96 samples
    80% ethanol	7.1 ml	7.2 ml	7.2 ml

24. Insert the trough of freshly prepared 80% ethanol in the position on screen. Click 'Ok' to continue.

    

25. Resuspend the AMPure XP beads by vortexing.
26. Important Ensure the AMPure XP beads are well mixed before use by vortexing.
27. Insert the nuclease-free water, Short Fragment Buffer (SFB) and AMPure XP beads in the positions on screen. Select 'Ok' to continue.

    Reagent	X24 samples	X48 samples	X96 samples
    Beads	2317 µl	2633.6 µl	3.2 ml
    Nuclease-free water	8.4 ml	8.8 ml	11 ml
    Short Fragment Buffer (SFB)	2550 µl	3.1 ml	4.2 ml

    

28. Spin down the Post PCR Clean Up Samples and the Native Barcode plate (referenced as 'Input Barcode HSP plate').
29. Insert 1000 µl tips, Post PCR Clean Up Samples plate and the Input Barcode HSP plate to their correct positions on screen. Select 'Ok' to continue.

    

30. Select the first and last tip positions to use in the 'Edit Tip Count' window. Click 'Ok' to continue.

    

31. Mix and load the tubes of prepared mastermixes into their positions on screen. Select 'Ok' to start the run.

    The status window in the bottom right of the screen will illustrate what the robot is doing.

    

32. Tip If all the barcodes on the Native Barcode plate are not required, the remaining barcodes can be unloaded from the deck as soon as the native barcode ligation reaction has finished. Ensure the unused barcodes are sealed and stored at -20°C for future use.

    

33. When the run is complete, unload the deck following the on screen instructions:

    Output plate:
    

    
    
    Barcode plate (if not previously removed):
    

    
    
    Input sample plate:
    

    
    
    Normalised sample plate:
    

34. Do not unload the carrier by clicking 'No'.

    

35. Click 'Ok' on the summary box to continue.

    

36. Quantify 1 µl of the pooled sample in the output plate using a Qubit fluorometer off deck.
37. Update the worklist with the measured concentration of the pooled sample for Processes 9-11.
38. End of Step Take the output plate containing the normalised pooled barcoded samples in A2, into the Adapter Ligation and Clean-up step.

1. Consumables quantities:

   Consumables	X24, X48 and X96 samples
   Hamilton 50 µl CO-RE tips with filter	6
   Hamilton 300 µl CO-RE tips with filter	2
   Hamilton 1000 µl CO-RE tips with filter	25
   Hamilton 60 ml Reagent Reservoir, Self-Standing with Lid	1
   Bio-Rad Hard-Shell® 96-Well PCR Plate	1
   Roche Diagnostics MagNA Pure LC Medium Reagent Tubs 20	2
   Sarstedt Inc Screw Cap Micro Tube 2ml	2
   Abgene™ 96 Well 0.8mL Polypropylene Deepwell Storage Plate	1

2. Reagents required:

   Reagents	X24, X48 and X96 samples
   Adapter Mix II (AMII)	18.75 µl
   Short Fragment Buffer (SFB)	2250 µl
   Elution Buffer (EB)	35 µl
   NEBNext Quick Ligation Buffer	37.5 µl
   NEBNext T4 DNA Ligase	18.75 µl
   AMPure XP Beads	2022 µl

3. Tip Native barcoded pooled samples from different runs may be processed together in this step. Transfer all the pooled samples into clean wells of the output plate from the previous step, now referenced as the Normalised Pool plate.
4. Important The CPAC and HHS units are required and must be switched on.
5. Thaw the Elution Buffer (EB), Short Fragment Buffer (SFB), and NEBNext Quick Ligation Reaction Buffer (5x) at room temperature and mix by vortexing. Then spin down and place on ice. Check the contents of each tube are clear of any precipitate.
6. Spin down the T4 Ligase and the Adapter Mix II (AMII), and place on ice.
7. From Run Control, click the green play button to begin setting up the deck.

   

8. Select 'Process10: Adapter Ligation' to start.

   

9. Select 'Process11: Adapter ligation cleanup' to stop the automated run and quantify the samples before sequencing.

   

10. Click 'Browse' to upload the Input File Worklist with the updated sample concentrations and well identifications and click 'OK'.

    

    Update the worklist for the Source_Well and Target_Well match the position of the pooled samples in the input plate.

    Note: It is possible to transfer pools from multiple runs into a single plate to run adapter ligation (process 10 and 11), up to a maximum of 8 source wells.

    

11. Prepare the Adapter Ligation Mastermix with the following reagents according to the Hamilton user interface. Click either 'Yes' or 'No' to continue.

    Note: It is user preference whether to print and save the instructions.

    Take care when pipetting the viscous AMII.

    Reagents	X24, X48 and X96 samples
    Adapter Mix II (AMII)	18.75 µl
    Quick T4 DNA Ligase	18.75 µl
    NEBNext Quick Ligation Reaction Buffer	37.5 µl

    

12. Insert plates to their corresponding positions on screen. Select 'Ok' to continue.

    

13. If required, reload a full deck of 50 µl tips into the positions on screen. Select 'Ok' to continue.

    Note: If there are enough tips left, there is no need to reload tips.

    

14. Select the first and last tip positions to use in the 'Edit Tip Count' window. Click 'Ok' to continue.

    

15. If required, reload a full deck of 300 µl tips into the positions on screen. Select 'Ok' to continue.

    Note: If there are enough tips left from the previous run, there is no need to reload tips.

    

16. Select the first and last tip positions to use in the 'Edit Tip Count' window. Click 'Ok' to continue.

    

17. Resuspend the AMPure XP beads by vortexing.
18. Important Ensure the AMPure XP beads are well mixed before use by vortexing.
19. Insert the Short Fragment Buffer (SFB) and AMPure XP beads in their positions on screen. Click 'Ok' to continue.

    Reagents	X24, X48 and X96 samples
    Short Fragment Buffer (SFB)	2250 µl
    Beads	2022 µl

    

20. Insert the Elution Buffer (EB) in their position on screen. Click 'Ok' to continue.

    Reagents	X24, X48 and X96 samples
    Elution Buffer (EB)	35 µl

    

21. Load the 1000 µl tips and the Normalised Pool Plate to the correct positions on screen. Click 'Ok' to continue.

    Note: The Normalised Pool Plate is the output plate from the previous step, containing the pooled barcoded samples.

    

22. Highlight the 1000 µl tips available to use on the 'Edit Tip Count' window. Select 'Ok' to continue.

    

23. Mix and insert the prepared Adapter Ligation Mastermix into their positions on screen. Click 'Ok' to start the automated run.

    

24. When the run is complete, unload the robot following the Hamilton user interface as below:

    Eluted DNA samples:
    

    
    
    Input Sample plate:
    

25. Do not unload the carrier by clicking 'No'.

    

26. Click 'Ok' on the summary box to continue.

    

27. Quantify 1 µl of each eluted sample in the Eluted DNA Samples plate using a Qubit fluorometer off deck.
28. End of Step Seal the plate once the library is prepared and store on ice until ready to load onto the flow cell.

    We do not recommend running the liquid handling robot overnight as the plate must be sealed and stored on ice as soon as library preparation is finished.

29. Tip Library storage recommendations

    We recommend storing libraries in Eppendorf DNA LoBind tubes at 4°C for short-term storage or repeated use, for example, re-loading flow cells between washes.
    For single use and long-term storage of more than 3 months, we recommend storing libraries at -80°C in Eppendorf DNA LoBind tubes.

30. Optional Action

    If quantities allow, the library may be diluted in Elution Buffer (EB) for splitting across multiple flow cells.

    Additional buffer for doing this can be found in the Sequencing Auxiliary Vials expansion (EXP-AUX001), available to purchase separately. This expansion also contains additional vials of Sequencing Buffer (SQB) and Loading Beads (LB), required for loading the libraries onto flow cells.

1. Thaw the Sequencing Buffer (SQB), Loading Beads (LB), Flush Tether (FLT) and one tube of Flush Buffer (FB) at room temperature before mixing the reagents by vortexing, and spin down at room temperature.
2. To prepare the flow cell priming mix, add 30 µl of thawed and mixed Flush Tether (FLT) directly to the tube of thawed and mixed Flush Buffer (FB), and mix by vortexing at room temperature.
3. Open the MinION device lid and slide the flow cell under the clip.

   Press down firmly on the flow cell to ensure correct thermal and electrical contact.

   

   

4. Optional Action

   Complete a flow cell check to assess the number of pores available before loading the library.

   This step can be omitted if the flow cell has been checked previously.

   See the flow cell check instructions in the MinKNOW protocol for more information.

5. Slide the priming port cover clockwise to open the priming port.

   

6. Important Take care when drawing back buffer from the flow cell. Do not remove more than 20-30 µl, and make sure that the array of pores are covered by buffer at all times. Introducing air bubbles into the array can irreversibly damage pores.
7. After opening the priming port, check for a small air bubble under the cover. Draw back a small volume to remove any bubbles:
   1. Set a P1000 pipette to 200 µl
   2. Insert the tip into the priming port
   3. Turn the wheel until the dial shows 220-230 µl, to draw back 20-30 µl, or until you can see a small volume of buffer entering the pipette tip
      

   Note: Visually check that there is continuous buffer from the priming port across the sensor array.

   

8. Load 800 µl of the priming mix into the flow cell via the priming port, avoiding the introduction of air bubbles. Wait for five minutes. During this time, prepare the library for loading by following the steps below.

   

9. Thoroughly mix the contents of the Loading Beads (LB) tubes by vortexing.
10. Tip Using the Loading Beads

    Demo of how to use the Loading Beads.

11. In a new tube, prepare the library for loading as follows:

    Reagent	Volume per flow cell
    Sequencing Buffer (SQB)	34 µl
    Loading Beads (LB), mixed immediately before use	25.5 µl
    Nuclease-free water	4.5 µl
    DNA library	11 µl
    Total	75 µl

    Note: Load the library onto the flow cell immediately after adding the Sequencing Buffer (SQB) and Loading Beads (LB) because the fuel in the buffer will start to be consumed by the adapter.

12. Important The Loading Beads (LB) tube contains a suspension of beads. These beads settle very quickly. It is vital that they are mixed immediately before use.
13. Complete the flow cell priming:
    1. Gently lift the SpotON sample port cover to make the SpotON sample port accessible.
    2. Load 200 µl of the priming mix into the flow cell priming port (not the SpotON sample port), avoiding the introduction of air bubbles.

    

    

14. Mix the prepared library gently by pipetting up and down just prior to loading.
15. Add 75 μl of the prepared library to the flow cell via the SpotON sample port in a dropwise fashion. Ensure each drop flows into the port before adding the next.

    

16. Gently replace the SpotON sample port cover, making sure the bung enters the SpotON port and close the priming port.

    

    

17. Important Install the light shield on your flow cell as soon as library has been loaded for optimal sequencing output.

    We recommend leaving the light shield on the flow cell when library is loaded, including during any washing and reloading steps. The shield can be removed when the library has been removed from the flow cell.

18. Place the light shield onto the flow cell, as follows:

    1. Carefully place the leading edge of the light shield against the clip.
       Note: Do not force the light shield underneath the clip.

    2. Gently lower the light shield onto the flow cell. The light shield should sit around the SpotON cover, covering the entire top section of the flow cell.

    

19. Caution The MinION Flow Cell Light Shield is not secured to the flow cell and careful handling is required after installation.
20. End of Step Close the device lid and set up a sequencing run on MinKNOW.

1. Overview of nanopore data analysis

   For a full overview of nanopore data analysis, which includes options for basecalling and post-basecalling analysis, please refer to the Data Analysis document.

2. How to start sequencing

   The sequencing device control, data acquisition and real-time basecalling are carried out by the MinKNOW software. Please ensure MinKNOW is installed on your computer or device. There are multiple options for how to carry out sequencing:

   1. Data acquisition and basecalling in real-time using MinKNOW on a computer

   Follow the instructions in the MinKNOW protocol beginning from the "Starting a sequencing run" section until the end of the "Completing a MinKNOW run" section.

   2. Data acquisition and basecalling in real-time using the MinION Mk1B/Mk1D device

   Follow the instructions in the MinION Mk1B user manual or the MinION Mk1D user manual.

   3. Data acquisition and basecalling in real-time using the MinION Mk1C device

   Follow the instructions in the MinION Mk1C user manual.

   4. Data acquisition and basecalling in real-time using the GridION device

   Follow the instructions in the GridION user manual.

   5. Data acquisition and basecalling in real-time using the PromethION device

   Follow the instructions in the PromethION user manual or the PromethION 2 Solo user manual.

   6. Data acquisition using MinKNOW on a computer and basecalling at a later time using MinKNOW

   Follow the instructions in the MinKNOW protocol beginning from the "Starting a sequencing run" section until the end of the "Completing a MinKNOW run" section. When setting your experiment parameters, set the Basecalling tab to OFF. After the sequencing experiment has completed, follow the instructions in the Post-run analysis section of the MinKNOW protocol.

3. Important Required settings in MinKNOW

   When setting the sequencing parameters in MinKNOW, in the Basecalling set barcoding as Enabled, and in the barcoding options, toggle Barcode both ends, Mid-read barcodes and Override minimum mid barcoding score to ON and set Minimum mid barcoding score to 50.
   Optional: basecalling and/or demultiplexing of sequences can be performed using the stand-alone Guppy software.

   

1. Recommended analysis pipeline

   The recommended workflows for the bioinformatics analyses are provided by the ARTIC network and are documented on their web pages at https://artic.network/ncov-2019/ncov2019-bioinformatics-sop.html.

   The reference guided genome assembly and variant calling are also performed according to the bioinformatics protocol provided by the ARTIC network. Their best practices guide uses the software contained within the FieldBioinformatics project on GitHub.

   This workflow uses only the basecalled FASTQ files to perform a high-quality reference-guided assembly of the SARS-CoV-2 genome. Sequenced reads are re-demultiplexed with the requirement that reads must contain a barcode at both ends of the sequence (this only applies to the Classic and Eco PCR tiling of SARS-CoV-2 protocols but not the Rapid Barcoding PCR tiling of SARS-CoV-2), and must not contain internal barcodes. The reads are mapped to the reference genome, primer sequences are excluded and the consensus sequence is polished. The Medaka software is used to call single-nucleotide variants while the ARTIC software reports the high-quality consensus sequence from the workflow.

2. To further simplify the installation of the coronavirus bioinformatics protocols, the workflows have been packaged into two EPI2ME products

   The FieldBioinformatics workflow for SARS-CoV-2 sequence analysis is provided as a Jupyter notebook tutorial in the EPI2ME Labs software. The coronavirus workflow has been augmented to include additional steps that help with the quality control of individual libraries, and aid in the presentation of summary statistics and the final sets of called variants.

   The FieldBioinformatics workflow for SARS-CoV-2 sequence analysis is also provided as an EPI2ME workflow – this provides a more accessible interface to a bioinformatics workflow and the provided cloud-based analysis also performs some secondary interpretation by preparing an additional report using the Nextclade software.

3. Expected results

   Here, results are shown based on human coronavirus 229E spiked into 100 ng of human RNA derived from GM12878 cell line. 10 pg–0.001 pg of viral RNA obtained from ATCC was spiked into the human RNA and human-only and reverse transcription negative controls were carried through the prep to sequencing. Every sample underwent 30 and 35 cycles of PCR to determine sensitivity and specificity guidelines, as well as the expected amplicon drop-out rate for each sample.

   Note: The viral RNA from ATCC is generated from cell lines infected with human coronavirus 229E. The RNA supplied is total RNA extracted from the cell lines and includes both human and viral RNA. Therefore, the levels of sensitivity are likely to be higher than those reported here.

4. Sample balancing

   The graph below shows the expected sequence balancing if the protocol is followed. Here, equal masses went into the end-prep and native barcode ligation prior to pooling by equal mass for adapter ligation.

   
   Figure 3. Number of reads per sample after native barcode demultiplexing in MinKNOW. All 14 samples were run on a single flow cell.

5. On-target rate

   Sequences from each demultiplexed sample were aligned to the human coronavirus 229E genome using minimap2. The proportion of primary alignments per sample are reported below.

   
   Figure 4. Proportion of reads for each sample aligning to the human coronavirus 229E reference genome.

6. Assessment of negative controls

   After 12 hours of sequencing, the number of reads from the negative control samples aligning to the viral reference genome is shown in the graph below and is compared with the absolute number of sequences aligning to the lowest input (0.001 pg).

   
   Figure 5. Absolute number of reads aligning to the human coronavirus 229E reference genome in the negative controls compared with the lowest input of viral RNA. Sequencing was carried out for 12 hours to pick up low levels of sequences assigned to barcodes representing these samples.

7. Target coverage for different PCR cycles and viral load

   To assess the impact of PCR dropout with lowering input viral load and increasing PCR cycles, Mosdepth was used to calculate the proportion of the viral genome covered to different depth levels. These numbers were calculated after 12 hours of sequencing with 14 samples multiplexed.

   
   Figure 6. Coverage and depth of the human coronavirus 229E genome for different input quantities of viral RNA and different cycle numbers after 12 hours of sequencing on a single flow cell.

8. How much sequencing is required?

   This is unknown in real clinical samples. The graph below can be used to determine the proportion of the genome that could be covered to a given depth with different numbers of reads (30 cycles) at different input amounts in a background of 100 ng human RNA.
   Note: this is absolute depth.

   
   Figure 7. Subsampled sequences to give an indication of the depth of sequencing achievable covering different amounts of the human coronavirus 229E genome. Input quantities and cycle number titrations show that high cycle numbers should be avoided where possible to minimise amplicon drop out.

   This protocol provides amplification of low copy number viral genomes in a tiled method with low off-target amplification and minimal cross-contamination between samples. With <60 copies per reaction (0.001 pg viral input) in 100 ng background human RNA, under ideal circumstances, one should expect to cover >75% of the targeted genome at a depth of 200X within under 50,000 reads in the samples with the lowest viral titre and <20,000 reads in those with a higher viral titre.

1. After your sequencing experiment is complete, if you would like to reuse the flow cell, please follow the Flow Cell Wash Kit protocol and store the washed flow cell at +2°C to +8°C.

   The Flow Cell Wash Kit protocol is available on the Nanopore Community.

2. Tip We recommend you to wash the flow cell as soon as possible after you stop the run. However, if this is not possible, leave the flow cell on the device and wash it the next day.
3. Alternatively, follow the returns procedure to send the flow cell back to Oxford Nanopore.

   Instructions for returning flow cells can be found here.

4. Important If you encounter issues or have questions about your sequencing experiment, please refer to the Troubleshooting Guide that can be found in the online version of this protocol.

1. Please contact your automation vendor FAS and/or Nanopore FAS if you have any issues.

1. Below is a list of the most commonly encountered issues, with some suggested causes and solutions.

   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.

2. Fewer pores at the start of sequencing than after Flow Cell Check

   Observation	Possible cause	Comments and actions
   MinKNOW reported a lower number of pores at the start of sequencing than the number reported by the Flow Cell Check	An air bubble was introduced into the nanopore array	After the Flow Cell Check it is essential to remove any air bubbles near the priming port before priming the flow cell. If not removed, the air bubble can travel to the nanopore array and irreversibly damage the nanopores that have been exposed to air. The best practice to prevent this from happening is demonstrated in this video.
   MinKNOW reported a lower number of pores at the start of sequencing than the number reported by the Flow Cell Check	The flow cell is not correctly inserted into the device	Stop the sequencing run, remove the flow cell from the sequencing device and insert it again, checking that the flow cell is firmly seated in the device and that it has reached the target temperature. If applicable, try a different position on the device (GridION/PromethION).
   MinKNOW reported a lower number of pores at the start of sequencing than the number reported by the Flow Cell Check	Contaminations in the library damaged or blocked the pores	The pore count during the Flow Cell Check is performed using the QC DNA molecules present in the flow cell storage buffer. At the start of sequencing, the library itself is used to estimate the number of active pores. Because of this, variability of about 10% in the number of pores is expected. A significantly lower pore count reported at the start of sequencing can be due to contaminants in the library that have damaged the membranes or blocked the pores. Alternative DNA/RNA extraction or purification methods may be needed to improve the purity of the input material. The effects of contaminants are shown in the Contaminants Know-how piece. Please try an alternative extraction method that does not result in contaminant carryover.

3. MinKNOW script failed

   Observation	Possible cause	Comments and actions
   MinKNOW shows "Script failed"		Restart the computer and then restart MinKNOW. If the issue persists, please collect the MinKNOW log files and contact Technical Support. If you do not have another sequencing device available, we recommend storing the flow cell and the loaded library at 4°C and contact Technical Support for further storage guidance.

4. Pore occupancy below 40%

   Observation	Possible cause	Comments and actions
   Pore occupancy <40%	Not enough library was loaded on the flow cell	Ensure you load the recommended amount of good quality library in the relevant library prep protocol onto your flow cell. Please quantify the library before loading and calculate mols using tools like the Promega Biomath Calculator, choosing "dsDNA: µg to pmol"
   Pore occupancy close to 0	The Ligation Sequencing Kit was used, and sequencing adapters did not ligate to the DNA	Make sure to use the NEBNext Quick Ligation Module (E6056) and Oxford Nanopore Technologies Ligation Buffer (LNB, provided in the sequencing kit) at the sequencing adapter ligation step, and use the correct amount of each reagent. A Lambda control library can be prepared to test the integrity of the third-party reagents.
   Pore occupancy close to 0	The Ligation Sequencing Kit was used, and ethanol was used instead of LFB or SFB at the wash step after sequencing adapter ligation	Ethanol can denature the motor protein on the sequencing adapters. Make sure the LFB or SFB buffer was used after ligation of sequencing adapters.
   Pore occupancy close to 0	No tether on the flow cell	Tethers are adding during flow cell priming (FLT/FCT tube). Make sure FLT/FCT was added to FB/FCF before priming.

5. Shorter than expected read length

   Observation	Possible cause	Comments and actions
   Shorter than expected read length	Unwanted fragmentation of DNA sample	Read length reflects input DNA fragment length. Input DNA can be fragmented during extraction and library prep. 1. Please review the Extraction Methods in the Nanopore Community for best practice for extraction. 2. Visualise the input DNA fragment length distribution on an agarose gel before proceeding to the library prep. In the image above, Sample 1 is of high molecular weight, whereas Sample 2 has been fragmented. 3. During library prep, avoid pipetting and vortexing when mixing reagents. Flicking or inverting the tube is sufficient.

6. Large proportion of unavailable pores

   Observation	Possible cause	Comments and actions
   Large proportion of unavailable pores (shown as blue in the channels panel and pore activity plot) The pore activity plot above shows an increasing proportion of "unavailable" pores over time.	Contaminants are present in the sample	Some contaminants can be cleared from the pores by the unblocking function built into MinKNOW. If this is successful, the pore status will change to "sequencing pore". If the portion of unavailable pores stays large or increases: 1. A nuclease flush using the Flow Cell Wash Kit (EXP-WSH004) can be performed, or 2. Run several cycles of PCR to try and dilute any contaminants that may be causing problems.

7. Large proportion of inactive pores

   Observation	Possible cause	Comments and actions
   Large proportion of inactive/unavailable pores (shown as light blue in the channels panel and pore activity plot. Pores or membranes are irreversibly damaged)	Air bubbles have been introduced into the flow cell	Air bubbles introduced through flow cell priming and library loading can irreversibly damage the pores. Watch the Priming and loading your flow cell video for best practice
   Large proportion of inactive/unavailable pores	Certain compounds co-purified with DNA	Known compounds, include polysaccharides, typically associate with plant genomic DNA. 1. Please refer to the Plant leaf DNA extraction method. 2. Clean-up using the QIAGEN PowerClean Pro kit. 3. Perform a whole genome amplification with the original gDNA sample using the QIAGEN REPLI-g kit.
   Large proportion of inactive/unavailable pores	Contaminants are present in the sample	The effects of contaminants are shown in the Contaminants Know-how piece. Please try an alternative extraction method that does not result in contaminant carryover.

8. Reduction in sequencing speed and q-score later into the run

   Observation	Possible cause	Comments and actions
   Reduction in sequencing speed and q-score later into the run	For Kit 9 chemistry (e.g. SQK-LSK109), fast fuel consumption is typically seen when the flow cell is overloaded with library (please see the appropriate protocol for your DNA library to see the recommendation).	Add more fuel to the flow cell by following the instructions in the MinKNOW protocol. In future experiments, load lower amounts of library to the flow cell.

9. Temperature fluctuation

   Observation	Possible cause	Comments and actions
   Temperature fluctuation	The flow cell has lost contact with the device	Check that there is a heat pad covering the metal plate on the back of the flow cell. Re-insert the flow cell and press it down to make sure the connector pins are firmly in contact with the device. If the problem persists, please contact Technical Services.

10. Failed to reach target temperature

    Observation	Possible cause	Comments and actions
    MinKNOW shows "Failed to reach target temperature"	The instrument was placed in a location that is colder than normal room temperature, or a location with poor ventilation (which leads to the flow cells overheating)	MinKNOW has a default timeframe for the flow cell to reach the target temperature. Once the timeframe is exceeded, an error message will appear and the sequencing experiment will continue. However, sequencing at an incorrect temperature may lead to a decrease in throughput and lower q-scores. Please adjust the location of the sequencing device to ensure that it is placed at room temperature with good ventilation, then re-start the process in MinKNOW. Please refer to this link for more information on MinION temperature control.

11. Guppy – no input .fast5 was found or basecalled

    Observation	Possible cause	Comments and actions
    No input .fast5 was found or basecalled	input_path did not point to the .fast5 file location	The --input_path has to be followed by the full file path to the .fast5 files to be basecalled, and the location has to be accessible either locally or remotely through SSH.
    No input .fast5 was found or basecalled	The .fast5 files were in a subfolder at the input_path location	To allow Guppy to look into subfolders, add the --recursive flag to the command

12. Guppy – no Pass or Fail folders were generated after basecalling

    Observation	Possible cause	Comments and actions
    No Pass or Fail folders were generated after basecalling	The --qscore_filtering flag was not included in the command	The --qscore_filtering flag enables filtering of reads into Pass and Fail folders inside the output folder, based on their strand q-score. When performing live basecalling in MinKNOW, a q-score of 7 (corresponding to a basecall accuracy of ~80%) is used to separate reads into Pass and Fail folders.

13. Guppy – unusually slow processing on a GPU computer

    Observation	Possible cause	Comments and actions
    Unusually slow processing on a GPU computer	The --device flag wasn't included in the command	The --device flag specifies a GPU device to use for accelerate basecalling. If not included in the command, GPU will not be used. GPUs are counted from zero. An example is --device cuda:0 cuda:1, when 2 GPUs are specified to use by the Guppy command.