TITLE: Monkeypox virus multiplexed PCR amplicon sequencing (PrimalSeq) V.1. 
AUTHORS: Nicholas      F.G. Chen*, Luc Gagne*, Matthew Doucette, Sandra Smole, Erika Buzby, Joshua Hall, Stephanie Ash, Rachel Harrington, Seana Cofsky, Selina Clancy, Curtis J Kapsak, Joel  Sevinsky, Kevin  Libuit, Chrispin Chaguza, Nathan D. Grubaugh, Daniel J. Park, Glen R. Gallagher#, Chantal B.F. Vogels#

[DESCRIPTION]
Background: The current global outbreak of Monkeypox virus (MPXV) concurrent with an ongoing SARS-CoV-2 pandemic has further highlighted the need for genomic surveillance and pathogen whole genome sequencing. While metagenomic and hybrid capture sequencing approaches were used to sequence many of the early MPXV cases, the viability of these methods is dependent on samples with high viral DNA concentrations. Given the atypical clinical presentation of cases associated with the current outbreak and uncertainty regarding viral load across both the course of infection and anatomical body sites, there is a strong need for a more sensitive and broadly applicable sequencing approach. Amplicon-based sequencing (PrimalSeq) was initially developed for sequencing of Zika virus, and later adapted as the main sequencing approach for SARS-CoV-2. Here, we used PrimalScheme to design a primer scheme for MPXV and we validated it with widely used SARS-CoV-2 sequencing protocols. Based on initial validation, our approach shows notably higher depth and breadth of coverage across the genome, particularly with higher PCR cycle threshold (Ct) samples, as compared to metagenomic sequencing. While further testing is needed, the early success of this approach has significant implications for sequencing efforts of the current MPXV outbreak and serves as a proof of concept of amplicon-based sequencing for use with other large-genome DNA viruses and potentially bacterial genomes.

Overview of Design: We used PrimalScheme (https://primalscheme.com/) to generate an MPXV primer scheme using a pre-outbreak A.1 clade reference genome (GenBank accession: MT903345). The primer scheme comprises a total of 163 primer pairs with an amplicon length ranging between 1597 and 2497 bp (average length of 1977 bp). The primer scheme is compatible with current ARTIC and COVIDSeq SARS-CoV-2 sequencing protocols, and while validated only with Illumina library prep methods and sequencing platforms, it would likely see a high degree of success with other sequencing platforms such as the Oxford Nanopore Technologies MinION. 

Initial Validation: We validated our MPXV primer scheme with clinical specimens at the Massachusetts Department of Public Health, Massachusetts State Public Health Laboratory under the IRB Project Titled “Rash Illness: Alternate Specimen Types and Sequencing” (protocol number 1917413). A total of 10 clinical specimens were included in this initial validation comprised of both throat and swabs of fluid from lesions from two individuals, belonging to the current outbreak-associated hMPXV B.1 clade as determined by prior characterization at the Centers of Disease Control and Prevention. Ct values were determined with the non-variola orthopox diagnostic qPCR assay developed for use by the Laboratory Response Network (Rapid Diagnostic Testing for Response to the Monkeypox Outbreak — Laboratory Response Network, United States, May 17–June 30, 2022 | MMWR (cdc.gov)). Clinical samples ranged in cycle threshold (Ct) values from 15.03 (high viral concentration) to 34.63 (low viral concentration), and each sample was sequenced in parallel using a metagenomics approach and the provided amplicon-based approach. Extractions and sequencing analysis were performed in accordance with current biosafety guidance including extraction in a BSL-3 setting. An evaluation of site-specific biosafety practices should be developed in consultation with your organization’s biosafety officer. Libraries were prepared with the Illumina DNA prep kit and sequenced on the MiSeq (v2 kit running 2x150 nt reads). Consensus genomes were generated at 10X coverage using the TheiaCoV_Illumina_PE Workflow Series on Terra.bio. A MPXV “fork” was developed for us by Curtis Kapsak from Theiagen, which included the MPXV reference sequence, primer scheme, and consensus genome length, and can currently be accessed at: https://github.com/theiagen/public_health_viral_genomics/tree/cjk-MPXV-theiacov. A non-MPXV specific TheiaCov workflow can also be found at: https://public-health-viral-genomics-theiagen.readthedocs.io/en/latest/theiacov_workflows.html with an accompanying protocols.io at: https://www.protocols.io/private/0E095FC2AC5211EC82F60A58A9FEAC02. We found comparable genome coverage between amplicon and metagenomic sequencing with low Ct (<18) samples, yet a significant increase in genome coverage with amplicon sequencing in higher Ct samples (>18; Figure 1.) Our findings highlight how amplicon-based approaches can significantly expand MPXV sequencing to a wider variety of samples. 

  
Amongst samples with a low Ct (<18), genome coverage via amplicon sequencing was consistently >97%, with minimal amplicon drop-outs (Figure 2). Amplicons 11, 75, and 118 showed consistent drop-out across the sequenced samples, while none of the primers had mismatches, except for a single nucleotide mismatch in the 11_RIGHT primer. We did obtain coverage for these amplicons when sequencing clade 2 DNA from cultured virus (strain USA-2003; NR-4928) obtained from BEI Resources (NIAID, NIH). This suggests that the dropouts of these primers may be a specific issue related to the current hMPXV B.1 clade genomes. As this protocol is still in development, we will further investigate performance as we continue to sequence additional samples. 

  
Conclusion: We developed an amplicon-based sequencing (PrimalSeq) approach for MPXV that improved the depth and breadth of genome coverage with low viral concentration specimens as compared to metagenomic sequencing.


This protocol represents the first iteration of development. Further versions will be uploaded to protocols.io with an accompanying description of changes as appropriate. 


[BEFORE_START]
This protocol is currently in the developmental phase. It is for research purposes only and should not be used in a diagnostic capacity. 

If using clinical samples, DNA will need to be extracted and purified prior to beginning library preparation.

Primer diluting and pooling requires several hours and may be conducted well in advance so long as the pooled primers are stored at -20°C.
Following primer dilution and pooling, this workflow can be completed in one day, however, it is recommended to be conducted across two, with the amplicon generation step on the first day and all subsequent wet-lab steps on the second day. 


[STEPS]
SECTION: Amplicon Generation
3. Reagents:
   





SECTION: Amplicon Generation
3.1. In two separate tubes, prepare the following master mixes: 

 
 



SECTION: Amplicon Generation
3.2. Label two sets of PCR tubes/plates for Pool 1 and Pool 2
SECTION: Amplicon Generation
3.3. Add the following: 
20 µL Pool 1 master mix to each Pool 1 tube/well
20 µL Pool 2 master mix to each Pool 2 tube/well

SECTION: Amplicon Generation
3.4. Add 5 µL DNA to each tube in both sets
Mix by pipetting up and down 10 times
Briefly centrifuge tubes/plates
 


SECTION: Amplicon Generation
3.5. Place on thermocycler and run the following program (choose preheat lid option):
   

SECTION: Amplicon Generation
3.6. *Safe Stopping Point: Amplified DNA may remain in the thermocycler at 4°C or stored at -20°C until ready to use*

SECTION: Amplicon Tagmentation and Clean Up
4. Reagents: 
  

SECTION: Amplicon Tagmentation and Clean Up
4.1. Spin down PCR tubes/plates
SECTION: Amplicon Tagmentation and Clean Up
4.2. Prepare the following master mix:

  

SECTION: Amplicon Tagmentation and Clean Up
4.3. In each PCR tube add:
30 µL  Master mix 
10 µL  Pool 1 amplicons
10 µL  Pool 2 amplicons

For a total reaction volume of 50 µL per PCR tube/well
 

SECTION: Amplicon Tagmentation and Clean Up
4.4. Mix by pipetting up and down and briefly centrifuge
SECTION: Amplicon Tagmentation and Clean Up
4.5. Place on thermocycler and run the following program (choose preheat lid option):

  
SECTION: Amplicon Tagmentation and Clean Up
4.6. Once the thermocycler reaches 10 °C , remove tubes/plates and spin down

SECTION: Amplicon Tagmentation and Clean Up
4.7. Add 10 µL  ST2 to each tube/well, mix by pipetting up and down, and briefly centrifuge

SECTION: Amplicon Tagmentation and Clean Up
4.8. Incubate at Room temperature for 5 min 

SECTION: Amplicon Tagmentation and Clean Up
4.9. Place on magnetic stand and wait until liquid is clear (a few minutes)
SECTION: Amplicon Tagmentation and Clean Up
4.10. Remove and discard all supernatant
SECTION: Amplicon Tagmentation and Clean Up
4.11. Remove tubes/plates from magnetic stand and add 100 µL TWB to each tube/well
Mix by pipetting up and down 10 times and spin down. Be careful to not introduce bubbles.


SECTION: Amplicon Tagmentation and Clean Up
4.12. Repeat steps 4.9 - 4.11. Leave the supernatant after the second wash so that the beads don’t dry out

SECTION: Amplify Tagmented Amplicons
5. Reagents: 
         
SECTION: Amplify Tagmented Amplicons
5.1. Prepare the following master mix:
   
SECTION: Amplify Tagmented Amplicons
5.2. Place the tubes/plates with tagmented amplicons on the magnetic stand and remove the supernatant once the liquid is clear
SECTION: Amplify Tagmented Amplicons
5.3. Use a 20µL pipette to remove any residual TWB from tubes/wells
SECTION: Amplify Tagmented Amplicons
5.4. Remove the tubes/plates from the magnetic stand
SECTION: Amplify Tagmented Amplicons
5.5. Add 40 µL master mix to each tube/well

SECTION: Amplify Tagmented Amplicons
5.6. Add 10 µL  dual-barcoded index adapters to each tube/well
   
 

SECTION: Amplify Tagmented Amplicons
5.7. Pipette up and down to mix and spin down
SECTION: Amplify Tagmented Amplicons
5.8. Place on thermocycler and run the following program (choose preheat lid option):
  
SECTION: Pool and Clean Up
6. Reagents: 
  
SECTION: Pool and Clean Up
6.1. Briefly centrifuge tubes/plates
SECTION: Pool and Clean Up
6.2. Place on magnetic stand and wait until liquid is clear (a few minutes)
SECTION: Pool and Clean Up
6.3. Pool libraries by equal volume:
  
SECTION: Pool and Clean Up
6.4. Pipette up and down to mix pooled libraries and briefly spin down
SECTION: Pool and Clean Up
6.5. Calculate the volume of ITB to reach a 0.6X beads:total pool volume ratio
 

SECTION: Pool and Clean Up
6.6. Mix by pipetting up and down, briefly centrifuge, and incubate at Room temperature for 5 min 

SECTION: Pool and Clean Up
6.7. Place on magnetic stand and wait until liquid is clear (a few minutes)
SECTION: Pool and Clean Up
6.8. Transfer supernatant to a new tube (do not discard, this is your final library). This volume will be slightly lower than the total combined volume
 

SECTION: Pool and Clean Up
6.9. Calculate the volume of beads to add to the supernatant to attain a second clean-up beads:sample ratio of 0.9X:

  
 

SECTION: Pool and Clean Up
6.10. Add beads to supernatant, mix by pipetting up and down, briefly centrifuge, and incubate at Room temperature for 5 min 
SECTION: Pool and Clean Up
6.11. Place on magnetic stand and wait until liquid is clear (a few minutes)
SECTION: Pool and Clean Up
6.12. Carefully discard supernatant
SECTION: Pool and Clean Up
6.13. Add 1000 µL 80% EtOH

SECTION: Pool and Clean Up
6.14. Wait 30 s 

SECTION: Pool and Clean Up
6.15. Repeat steps 6.12 - 6.14
SECTION: Pool and Clean Up
6.16. Remove supernatant using a 20µL pipette to remove all residual EtOH
SECTION: Pool and Clean Up
6.17. Add 55 µLof RSB to the tube, mix by pipetting up and down, and briefly centrifuge

SECTION: Pool and Clean Up
6.18. Incubate at Room temperaturefor 2 min

SECTION: Pool and Clean Up
6.19. Place on magnetic stand and wait until liquid is clear (a few minutes)
SECTION: Pool and Clean Up
6.20. Transfer 50 µL of final pooled library to a new 1.5mL tube

SECTION: Pool and Clean Up
6.21. Quantify library on a Qubit and obtain fragment distribution using a Bioanalyzer/Tape Station

 


SECTION: Dilute and Pool Primers
1. Reagents: 
  
 


SECTION: Dilute and Pool Primers
1.1. If not already done, separate odd and even numbered primer pairs into two separate boxes. These will constitute the two pools 

 

SECTION: Dilute and Pool Primers
1.2. Label 164, 8-strip tubes with the corresponding odd-numbered primer name (e.g. 3 left)
SECTION: Dilute and Pool Primers
1.3. To each tube add 90 µLof nuclease-free water 
 


SECTION: Dilute and Pool Primers
1.4. For each odd-numbered primer tube: 
Spin down 
Pipette 10 times to mix 
Add 10 µL to the corresponding labeled tube 
Pipette 10 times to mix 

SECTION: Dilute and Pool Primers
1.5. After all 164 primers have been aliquoted, combine 10uL from each tube into a 2mL tube. This will be the odd-numbered primer pool 
 

SECTION: Dilute and Pool Primers
1.6. Repeat steps 1.2-1.5 with the even -numbered primers (NOTE: There will be two fewer primers in the even set than the odd) 
SECTION: Dilute and Pool Primers
1.7. *Safe Stopping Point: Pooled Primers Can Be Stored at -20°C*
2. Library Preparation Method
SECTION: Amplicon Generation
3. Reagents:
   




SECTION: Amplicon Generation
3.1. In two separate tubes, prepare the following master mixes: 

  
 




SECTION: Amplicon Generation
3.2. Label two sets of PCR tubes/plates for Pool 1 and Pool 2
 

SECTION: Amplicon Generation
3.3. Add the following: 
20 µL Pool 1 master mix to each Pool 1 tube/well
20 µL Pool 2 master mix to each Pool 2 tube/well

SECTION: Amplicon Generation
3.4. Add 5 µL  DNA to each tube/well in both sets
Mix by pipetting up and down 10 times
Briefly centrifuge 
 


SECTION: Amplicon Generation
3.5. Place on thermocycler and run the MPx Tiling PCR program:
 
  

SECTION: Amplicon Generation
3.6. *Safe Stopping Point: Amplified DNA may remain in the thermocycler at 4°C or stored at -20°C until ready to use*

SECTION: Amplicon Clean Up and Size Selection
4. Reagents:
  

SECTION: Amplicon Clean Up and Size Selection
4.4. Spin down PCR tubes/plates
SECTION: Amplicon Clean Up and Size Selection
4.5. In each designated PCR tube/well containing the AMPure XP beads add: 
25 µL  Pool 1 PCR reaction amplicons 
25 µL  Pool 2 PCR reaction amplicons

Mix by pipetting 10 times 
 

 


SECTION: Amplicon Clean Up and Size Selection
4.1. Prepare  400 µL of 80% EtOH per sample

SECTION: Amplicon Clean Up and Size Selection
4.2. Allow AMPure XP beads 30 min to reach room temperature and vortex to resuspend

SECTION: Amplicon Clean Up and Size Selection
4.3. Add 50 µLof AMPure XP beads to new PCR tubes/designated wells of a new plate and mix by pipetting 10 times 

SECTION: Amplicon Clean Up and Size Selection
4.6. Ensure tube is capped (or plate is sealed with foil cover) and place on shaker at 1800 rpm for 10 min at RT 

SECTION: Amplicon Clean Up and Size Selection
4.7. Place tube/plate on magnetic stand and wait until supernatant is clear (a few minutes)
SECTION: Amplicon Clean Up and Size Selection
4.8. Carefully discard supernatant, being sure not to disturb the bead pellet 
SECTION: Amplicon Clean Up and Size Selection
4.9. Wash beads by covering pellet with 200 µL of 80% EtOH 

SECTION: Amplicon Clean Up and Size Selection
4.11. Remove EtOH with pipette and discard
SECTION: Amplicon Clean Up and Size Selection
4.12. Repeat steps 4.9-4.11 for a total of 2 washes 
SECTION: Amplicon Clean Up and Size Selection
4.10. Incubate for 30 s
SECTION: Amplicon Clean Up and Size Selection
4.13. Remove supernatant using clean pipette tips to remove all residual 80% EtOH
SECTION: Amplicon Clean Up and Size Selection
4.14. Keep tube/plate on the magnetic stand and allow the bead to dry for 1-5 minutes 
 

SECTION: Amplicon Clean Up and Size Selection
4.15. Remove tube/plate from the magnetic stand, add 32 µL of nuclease-free water to resuspend beads, and mix by pipetting 

SECTION: Amplicon Clean Up and Size Selection
4.16. Ensure tube is capped (or plate is sealed with foil cover) and place on orbital shaker at 1800 rpm for 5 min at RT 
SECTION: Amplicon Clean Up and Size Selection
4.17. Place tube/plate on magnetic stand until supernatant is clear and colorless (~2 min)
SECTION: Amplicon Clean Up and Size Selection
4.18. Remove 32 µL of supernatant and transfer into a new tube/well (this constitutes the purified sample) 

SECTION: Amplicon Clean Up and Size Selection
4.19. *Safe Stopping Point: Purified DNA may be stored at -20°C*

SECTION: Quantification of DNA
5. Quantify sample using the Qubit 1x dsDNA HS kit on the Qubit Flex Fluorometer
 

SECTION: Quantification of DNA
5.1. Record stock concentration (ng/µL) 
 

SECTION: Quantification of DNA
5.2. *Safe Stopping Point: DNA may be stored at -20°C*

SECTION: Library Clean Up
10. Allow Sample Purification beads (SPB) to adjust to room temperature for least 30 min
 

SECTION: Library Clean Up
10.1. Thaw Resuspension Buffer (RSB) to RT. Vortex and quick spin before using
SECTION: Library Clean Up
10.4. Remove tubes/plate from thermocycler and quick spin. Place tubes/plate on magnetic stand for 2-5 mins or until clear
SECTION: Library Clean Up
10.5. Transfer 45 µL of supernatant, without disrupting the beads, into the corresponding tubes/wells containing the IPB master mix. Discard the used tubes/plate containing the beads

SECTION: Library Clean Up
10.2. Using a reagent reservoir, combine the following volumes to prepare the IPB master mix. Vortex and quick spin:

  

SECTION: Library Clean Up
10.3. Add 85 µL of IPB master mix to new tubes/designated wells of a new plate

SECTION: Library Clean Up
10.6. Cap tubes/seal plate and shake at 1600 rpm for 1 min on orbital shaker

SECTION: Library Clean Up
10.8. During incubation, perform the following steps: 
Vortex the undiluted stock tube of IPB
Add 15 µL  of undiluted IPB to new tubes/wells of a new plate 
Cap tubes/seal plate and set aside until step 10.10

SECTION: Library Clean Up
10.9. Place tube/plate containing samples and IPB master mix on magnetic stand and let sit for ~2 minutes or until supernatant is clear 
SECTION: Library Clean Up
10.10. Transfer 125 µL of supernatant to tubes/wells containing undiluted SPB from step 10.8. Pipette to mix and discard tubes/plate that contained diluted IPB

SECTION: Library Clean Up
10.11. Cap tubes/ seal plate and shake at 1600 rpm for 1 min on orbital shaker
SECTION: Library Clean Up
10.13. Place tubes/plate on a magnetic stand for ~2 minutes or until supernatant is clear 
SECTION: Library Clean Up
10.15. Remove and discard the supernatant
SECTION: Library Clean Up
10.16. Wash beads by covering pellet with 200 µL of 80% EtOH and incubate for 30 s. Remove ethanol with pipette and discard

SECTION: Library Clean Up
10.17. Repeat step 10.16 for a total of two washes
SECTION: Library Clean Up
10.18. Remove residual 80% EtOH with a clean pipette tip
SECTION: Library Clean Up
10.19. Keep tubes/plate on magnetic stand and let bead dry for 1-5 minutes
 

SECTION: Pooling and Library Pool Quantification
11. In a new 1.5mL tube, add 5 µL of each designated sample to create a library pool for sequencing
SECTION: Library Clean Up
10.24. Libraries may be quantified via a Qubit prior to pooling or, alternatively, pooling may be done via equal volumes under the assumption of approximately equal library concentrations 

SECTION: Pooling and Library Pool Quantification
11.1. Quantify library on a Qubit and obtain fragment distribution using a Bioanalyzer/Tape Station

 

SECTION: DNA Preparation
6. Samples must be quantified, calculated, and diluted before proceeding
SECTION: DNA Preparation
6.1. In preparation for the Amplify Tag DNA step, thaw Enhance PCR Mix (EPM) and IDT for Illumina Nextera DNA UD Indexes plate on cold block. Gently vortex and quick spin before using
 

SECTION: DNA Preparation
6.2. Thaw samples, Bead-Linked Transposomes (BLT), & Tagmentation Buffer (TB1) on a cold block. Vortex and quick spin before using
SECTION: DNA Tagmentation
7. In a new 1.5ml tube prepare the following tag master mix. Vortex master mix gently, spin down, and place in a cold block: 
  
SECTION: DNA Tagmentation
7.1. In a new tube/designated wells of a new plate, add:
20 µL Tag master mix 
30 µL sample (purified DNA input with concentration adjusted to 100-500ng) 

Pipette 10 times to mix

 

SECTION: DNA Tagmentation
7.2. Ensure tube is capped (or plate is sealed with foil cover), gently vortex, and spin down 
SECTION: DNA Tagmentation
7.3. Incubate at 55 °C for 5 min and hold at 10 °C using the Flex Tagment Program: 
  

SECTION: DNA Tagmentation
7.4. Vortex and quick spin Tagment Stop Buffer (TSB) before using
SECTION: DNA Tagmentation
7.5. Remove tube/plate containing Tag master mix and DNA input from thermocycler. Quick spin, and place on a cold block
SECTION: DNA Tagmentation
7.6. Immediately add 10 µL of TSB to each tube/well to stop reaction. Pipette to mix. Cap tube (or seal plate) and place back in thermocycler

SECTION: DNA Tagmentation
7.7. Incubate the reaction at37 °Cfor 15 min and hold at 10 °C using the Flex Post-Tag program:
  

SECTION: Clean Up
8. Remove tube/plate from thermocycler and place on the magnetic stand and wait until supernatant is clear (~2 minutes)
SECTION: Amplify Tagmented DNA
9. In a new 1.5 mL lo-bind tube, combine the following volumes to prepare the Flex Amplify master mix in the table below. Vortex and quick spin and place on cold block
  

SECTION: Clean Up
8.1. Remove and discard supernatant
SECTION: Clean Up
8.2. Remove tube/plate from the magnetic stand and, using a deliberately slow pipetting technique, add 100 µL Tagment Wash Buffer (TWB) directly onto the beads and slowly pipette to mix 10 times 
 

SECTION: Clean Up
8.3. Place the tube/plate back on the magnetic stand and wait until the supernatant is clear (~2 minutes). Remove and discard supernatant
SECTION: Clean Up
8.4. Remove tube/plate from magnetic stand and add another 100 µL of TWB. Slowly pipette to mix

SECTION: Clean Up
8.5. Cap tube/seal plate and leave on magnetic stand. DO NOT remove TWB until step 9.1
SECTION: Amplify Tagmented DNA
9.1. Remove and discard supernatant from the tube/plate containing tagmented DNA and TWB
SECTION: Amplify Tagmented DNA
9.2. Remove tube/plate from magnetic stand and place on cold block
SECTION: Amplify Tagmented DNA
9.3. Immediately add 40 µL of Flex Amplify master mix to tube containing DNA
SECTION: Amplify Tagmented DNA
9.4. Add 10 µL of assigned Illumina Nextera DNA UD Indexes to designated tube/well
 


SECTION: Amplify Tagmented DNA
9.6. Incubate tubes/plate following the Flex Amplify program:
  
 



SECTION: Amplify Tagmented DNA
9.7. *Safe Stopping Point: Capped Tube/ Sealed Plate may be stored at 2-8°C for up to 30 days*

SECTION: Bioinformatics/Results Analysis
13. Sequencing results may be analyzed utilizing a standard amplicon sequencing bioinformatics pipeline, including those employed for SARS-CoV-2 sequencing. 
 

SECTION: Bioinformatics/Analysis
8. Sequencing results may be analyzed utilizing a standard amplicon sequencing bioinformatics pipeline, including those employed for SARS-CoV-2 sequencing. 
 

SECTION: Amplify Tagmented DNA
9.5. Shake tubes/plate at 1600 rpm for 1 min and quick spin 
SECTION: Library Clean Up
10.7. Incubate on benchtop at RT for 5 min

SECTION: Library Clean Up
10.12. Incubate tubes/plate on benchtop at RT for 5 min

SECTION: Library Clean Up
10.14.  Prepare fresh 80% EtOH
SECTION: Library Clean Up
10.20. Remove tube/plate from magnetic stand and resuspend beads with 32 µL of RSB. Cap tube/seal plate and shake at 1600 rpm for 1 min on orbital shaker
SECTION: Library Clean Up
10.21. Let plate incubate on benchtop at RT for 2 min
SECTION: Library Clean Up
10.22. Place tube/plate on magnetic stand for ~2 min until supernatant is clear. Remove 32 µL of supernatant and add to new tube/wells of a new plate
 

SECTION: Library Clean Up
10.23. *Safe Stopping Point: Capped Tube/ Sealed Plate may be stored at -20°C*
SECTION: Sequencing
7. Protocol validated on the MiSeq (v2 kit running 2x150 nt reads)
 

SECTION: Sequencing
12. Protocol validated on the MiSeq (v2 kit running 2x150 nt reads)