Jul 03, 2026

Sample Preparation for PacBio SMRT-UMI Viral Sequencing for HIV Drug Resistance

DOI
https://dx.doi.org/10.17504/protocols.io.x54v9b3nql3e/v1
  • Ceejay Boyce1
  • 1Seattle Children's Research Institute
Ceejay Boyce
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DOI: https://dx.doi.org/10.17504/protocols.io.x54v9b3nql3e/v1
Protocol Citation: Ceejay Boyce 2026. Sample Preparation for PacBio SMRT-UMI Viral Sequencing for HIV Drug Resistance. protocols.io https://dx.doi.org/10.17504/protocols.io.x54v9b3nql3e/v1
License: This is an open access  protocol  distributed under the terms of the  Creative Commons Attribution License,  which permits unrestricted use, distribution, and reproduction in any medium, provided the original author and source are credited
Protocol status: Working
We use this protocol and it's working
Created: January 29, 2026
Last Modified: July 03, 2026
Protocol  Integer ID: 241822
Keywords: HIV drug resistance, PacBio sequencing
Abstract
This Standard Operating Procedure describes how to recover, amplify, quantify, and prepare amplicon libraries derived from HIV RNA for drug resistance genotyping (codons PR19-IN270) conducted using the Pacific Biosciences (PacBio) Single-Molecule Real-Time platform and universal molecular identifiers (SMRT-UMI).
Guidelines
Sample Integrity
Extracted RNA samples are removed from the BSL2 facility (extraction room) and immediately moved to pre-PCR benches for use in cDNA synthesis reactions, then stored in an ULT freezer for longer-term storage. cDNA preparations are stored in -20 °C freezers in pre-PCR for immediate use in 1st round PCR reactions and ULT freezers for longer-term storage. 1st round PCR reactions are prepared in pre-PCR and then permanently moved to post-PCR for amplification and further handling. 1st round PCR reactions are stored in -20 °C freezers in post-PCR. The master mix for 2nd round PCR reactions is prepared in pre-PCR and then permanently moved to post-PCR for addition of 1st round PCR product and subsequent PCR amplification. 2nd round PCR reactions are stored in -20 °C freezers in post-PCR.

Materials
Reagents and Materials
Recommended vendors are listed. Unless otherwise specified, products of equal or better quality than those listed can be used.
  1. Supply Sources
AB
ProductSource(s)
dNTPsGenscript, ThermoFisher
PrimeSTAR GXL (with dNTP and 5x Buffer) Takara
QIAamp Viral RNA mini kitQiagen
High Pure Viral Nucleic Acid kit Roche
RNasIN Plus (40U/ul)VWR
Superscript III (200U/ul) (with DTT and 5x Buffer) ThermoFisher
RNaseHThermoFisher
Agencourt AMPure XPBeckman Coulter
Nuclease-free waterSigma Aldrich, ThermoFisher
1M Tris-HCl (pH 8.0)Boston BioProducts
SMRTbell Express Template Prep Kit 2.0 (or v3.0)Pacific Biosciences
Barcoded Overhang Adapter Kit–8A (or 3.0 Plate)Pacific Biosciences
SMRTbell Enzyme Clean up KitPacific Biosciences
AMPure PB BeadsPacific Biosciences
Ethanol, Absolute (200 Proof)Fisher Scientific
1.5mL DNA LoBind tubesEppendorf
0.2 mL 8 & 12Strip Tubes; CapsVWR Scientific
96-well PCR PlateVWR Scientific
0.75% Agarose, 1-10 kb size selections, S1Sage Science
GeneRuler Express DNA LadderThermoFisher Scientific
AgaroseFisher Scientific
Ethidium Bromide (10mg/mL)Sigma Aldrich
8e5 (1x10 copies/5uL)Prepared in Frenkel Lab
1x SBPrepared in Frenkel Lab
6x Loading DyePrepared in Frenkel Lab
PCR primers
  • The primer to be used for cDNA synthesis for HIV pol is illustrated in Table 1 cDNA primers contain, from the 5’ end: sequences to be used as templates for reverse primers in 1st and 2nd round PCR, an 8-nucleotide random sequence universal molecular identifier (UMI), and an HIV-specific sequence.
  • Table 2 indicates the approximate size of each amplicon. This information is used to calculate extension times for PCR and quantify DNA concentrations.
  • Ordering Instructions: Oligos are supplied by Integrated DNA Technologies (IDT) with standard desalting and delivered LabReady (Normalized to 100uM in IDTE pH 8.0). cDNA primers with UMIs are ordered with handmixed bases. Primers without UMIs ordered as 100 nmole DNA Oligos
  • Preparation: Working aliquots are diluted from stock to 20uM, 10uM, or 2.5uM concentration using 1X TLE in 500uL aliquots. 50ul of working stock can be aliquoted into PCR strip tubes for easy multi-channel pipetting.
  • Storage: 100uM stocks and working aliquots are stored in -20°C freezer.
  • Stability: Stocks stable for many years if at high concentration (e.g., 100uM). Working aliquots should be used within 6 months.

dNTPs
  • Preparation: Working aliquots are diluted from stock to 10mM (each) using nuclease free water. 10mM working stocks are prepared in 1ml aliquots.
  • Storage: Stocks and 10mM working aliquots are stored in a -20°C freezer. 10mM working aliquots can be stored temporarily at 4°C during frequent usage (e.g., when used daily).
  • Stability: Stock and working solutions stable at -20°C. Avoid freeze-thaw cycles and frequent temperature changes.

10mM Tris-HCl (pH 8.0)
  • Preparation: Working aliquots are diluted from 1M stock to 10mM using nuclease free water. 10mM stocks are prepared in 15ml aliquots.
  • Storage: 1M stock and working aliquots stored at room T.
  • Stability: Refer to expiration date on package.

Positive Control 8e5
  • Preparation: Working aliquots are diluted from stock to 10 copies/5uL using 1X TLE + 10ng/ul tRNA. 10 copy/5uL working stocks are prepared in 50ul aliquots.
  • Storage: stock and 10 copy/5uL working aliquots are stored in -20°C freezer.
  • Stability: stock stable for years, working aliquots should not be used for >1 year.

1x Gel loading Buffer
  • Preparation: working aliquot is diluted 1:1 using nuclease free water (5mL)
  • Working aliquot is used to replenish 0.2mL strip tubes for ease of multi-channel pipetting.
  • Storage: undiluted and working stock are stored at room temperature.
  • Stability: Refer to expiration date on package.

1% agarose gel
  • Preparation: For small/medium/large gel (40/60/100mL), add 2/3/5ul EtBr (10mg/mL) and swirl to mix. Pour into casting tray and add desired combs. Allow to set at room T for ~30 minutes.
  • Storage: Melted agarose stocks – combine 2g Agarose with 200mL 1x SB and microwave to melt Agarose. Swirl to mix and store in bead bath.

GeneRuler Express DNA ladder (100-5000bp)
  • Preparation: For non-ready-made version, concentrated DNA ladder is combined with 100ul of TriTrack 6X loading buffer, and 400ul Nuclease-free water. Ready-made ladder requires no preparation aside from thawing.
  • Storage: Stock is kept at -20°C and working aliquots are stored at room T.
  • Stability: Refer to expiration date on package.
Safety warnings
All plasma samples should be handled in a BSL2 facility by trained and approved personnel.

This protocol requires prior approval by the users' Institutional Review Board or equivalent ethics committee for the use of human plasma specimens.
Viral RNA Extraction
Determine volume of plasma to extract.
Volume extracted is dependent on viral load:
  • If VL > 50,000 c/mL, extract 0.14 mL of plasma
  • If VL = 15,000 to 50,000 c/mL, extract 0.28 mL of plasma
  • If VL = 10,000 to 14,999 c/mL, extract 0.50 mL of plasma
  • If VL = 3,000 to 9,999 c/mL, extract 1 mL of plasma
  • If VL < 3,000 c/mL, extract 2 mL or more (if available, otherwise 1 mL)
Extract viral RNA using either the QIAmp Viral RNA mini kit or MiniMag extraction kit following the manufacturer's protocol.
  • Qiagen kit can be used for 0.14 and 0.28 mL extractions
  • MiniMag kit can be used for any volume extractions
  • Elute RNA in 60 µL of kit elution buffer
cDNA Synthesis
1h 22m
To amplify each gene region for sequencing, follow steps below using the appropriate primer sets from Table 1.
Table 1: cDNA & PCR Primers
ABC
Primer: 5’ --> 3’ Sequence:
cDNA
PB2_INTR1 CCCGCGTGGCCTCCTGAATTATCCGCTCCGTCCGACGACTCACTATACACTCANNNNNNNNCTTTTCCATGTTYTAATCYTCATCCTGTC Alternative cDNA primer
PB2_INTR2 CCCGCGTGGCCTCCTGAATTATCCGCTCCGTCCGACGACTCACTATACACTCANNNNNNNNCAATCAKCACCTGCCATCTGTTTTCCAT Primary cDNA primer
Forward
MozFO CCTACACCTGTCAACATAATTGG 1st Rd, for RT+IN
PB-F_modMozFI TCGTCGGCAGCGTCCTGTACCAGTAAAATTAAAGCCAGGAATGGATGG 2nd Rd, for RT+IN
modPRA GRAAAAARGGCTGTTGGAAATGTGG 1st Rd, for PR+RT+IN
PB-F_NEF11 TCGTCGGCAGCGTCCAAATCACTCTTTGGCARCGACC 2nd Rd, for PR+RT+IN
Reverse
PB-R1-alt1 CCCGCGTGGCCTCCTGAATTAT 1st Rd
PB2_INTR2_noUMI CCGCTCCGTCCGACGACTCACTATACAATCAKCACCTGCCATCTGTTTTCCAT 1st Rd for 8e5 PCR control
PB-R2-alt1_ill GTCTCGTGGGCTCGGCCGCTCCGTCCGACGACTCACTATA 2nd Rd
Key:
Sample Index: We have added Sample Index 02 CACTCA into primer to utilize the PORPID pipeline without issue. However, Sample Indexes can be added to make unique cDNA primers in lieu of downstream sample indexing for DNA library multiplexing.
(N)8: random template ID (UMI)
Green: HIV-1 gene specific sequence for cDNA synthesis
Blue: 5’ adapter sequence to add forward indexing primer
Purple: 3' adapter sequence to add reverse indexing primer
In thin-walled PCR tube, combine the following reagents and mix well (1st mix):
  • 25 µL RNA eluate
  • 2.5 µL dNTP mixture (10mM each)
  • 1 µL cDNA primer (20uM)
In PCR cycler, heat the mixture for 00:05:00 at 65 °C .
5m
Remove from PCR cycler and snap-chill on ice for 00:02:00 or chill at 4 °C in PCR cycler.
2m
In 0.5ml tube, prepare the SSIII master mix below, in this order:
  • 4 µL Nuclease free water
  • 10 µL 5x buffer
  • 2.5 µL 100mM DTT
  • 2.5 µL RnasIN plus (40U/ul)
  • 2.5 µL Superscript III (SSIII, 200U/ul)
Remove 1st mix from ice or PCR cycler, add the SSIII master mix, mix well and incubate at 50 °C for 01:00:00 in a PCR cycler.
1h
Inactivate SSIII by heating to 70 °C for 00:15:00 .
15m
Hold at 4 °C .
Add 1 µL (2U) RNase H and incubate at 37 °C for 00:20:00 .
Hold at 4 °C .
cDNA Purification
1h 6m 30s
cDNA must be purified to remove any excess UMI primer before downstream amplification. Note the following:
  • If two or more cDNA synthesis reactions were performed with the same primer, combine them together for cDNA cleanup
  • Use Agencourt AMPure XP or RNAclean XP beads for purification
  • Use 0.2mL PCR strip tubes or 96-well PCR plate
  • Make a fresh stock of room temperature 80% EtOH (320uL needed per sample)
Allow working stock tube of beads to warm to room temperature for 30-60 minutes.
30m
Resuspend beads by inverting the tube several times or vortexing until thoroughly mixed.
1m
Add equal volume of beads to cDNA (1:1 ratio)
4m
Pipette mix in tube/well 10-15 times until solution is uniform in color
2m
Incubate at Room temperature away from magnet for 00:10:00 , pipette mix (15x) after 00:05:00 .

  • Note: Sample-bead mixture can be incubated longer (~20 minutes total); however, 10 minutes is the minimum incubation time.
10m
Place tubes/plate on magnet and let sit for 00:05:00 or until solution is clear.

5m
Slowly remove and discard the supernatant with pipette (do not disturb beads). Leaving a small volume of supernatant (2-3uL) to avoid the beads is acceptable.
Dispense 100 µL freshly made Room temperature 80% EtOH with tubes still on the magnet without disturbing the beads.
  • Ethanol must completely cover the bead mass on the side of the tube for effective washing.


Incubate bead pellet with EtOH for 00:00:30 at Room temperature , then remove EtOH by pipette.

30s
Repeat steps 3.8 & 3.9 for a total of 3 washes.
  • If necessary, briefly centrifuge tubes/plate to collect EtOH at the bottom after final wash.

2m
Remove the last drop of EtOH by pipette. Incubate the tube/plate uncovered on magnet at Room temperature for 00:01:00 .
  • Do not over dry bead pellet (monitor for cracking) as this makes it difficult to resuspend for elution


1m
Away from the magnet, add nuclease-free water and gently pipette mix until the beads are completely resuspended.

Incubate at Room temperature for at least 00:05:00 .

5m
Place tube on magnet for 00:05:00 or until solution is clear.

5m
Transfer 30 µL of supernatant to 0.2mL PCR tube without disturbing beads.
  • If cDNA is not to be used immediately for 1st round PCR, prepare a separate 4.5 µL aliquot to be used for qPCR quantification to avoid additional freeze-thaw between qPCR and 1st round PCR.

1m
Store cDNA in -20 °C freezer for short-term and in ULT freezer for long-term storage.
cDNA Quantification
1h 19m
To quantify the number of HIV templates reverse transcribed, we perform a qPCR assay targeting a short region of HIV integrase.
  • For all samples with a viral load <5,000 c/mL, .
Setup qPCR reaction: 25uL rxn in duplicate for each cDNA sample (23uL mastermix + 2uL cDNA).
cDNA Mastermix (MM) recipe:
  • Nuclease-free water: 6 µL
  • iTaq universal probes supermix: 12.5 µL
  • INT Forward Primer Mix 20 micromolar (µM) : 0.5 µL
  • INT Reverse Primer Mix 20 micromolar (µM) : 0.5 µL
  • INT Probe 20 micromolar (µM) : 0.5 µL
  • Total Volume: 20 µL
Pipette 20 µL of MM into each well of the qPCR plate.
2m
Add 5 µL of standard curve DNA (8e5 HIV DNA stored at 10^4, 10^3, 10^2, and 10^1 copies/5uL in TLE+tRNA) in duplicate, with four replicates total for the 10^1 copies standard.
2m
Add 3 µL of nuclease-free water to all the unknown sample wells, then2 µL of cDNA sample to each well in duplicate.
5m
Seal plate and spin down contents. Place plate in thermocycler and run with the qPCR program conditions below:
ABC
CyclesTempTime
1X95C3:00
45X95C0:15
60C0:30

1h 10m
QC standard curve: look at threshold, Ct values, efficiency, R2
If the quantified amount is >10% of the expected amount based on viral load, continue to next step. Otherwise, check in with supervisor about whether to continue or troubleshoot to obtain more templates before amplifying.
1st Round PCR Amplification
Table 2: Target Amplicon Sizes and PCR Extension Times
ABC
AmpliconApproximate Size (bp)PCR Extension Time (min: sec)
RT+IN 1st Rd – primary set 2650 2:30
RT+IN 2nd Rd – primary set 2539 2:30
RT+IN 2nd Rd – alternate 2nd Rd F 2500 2:30
RT+IN 1st Rd – alternate cDNA primer 2600 2:30
RT+IN 2nd Rd – alternate cDNA primer 2500 2:30
RT+IN 2nd Rd – alternate cDNA & 2nd Rd F 2500 2:30
PR+RT+IN 1st Rd – primary set 3119 3:00
PR+RT+IN 1st Rd – alternate cDNA primer 3100 3:00
PR+RT+IN 2nd Rd – primary set 2852 3:00
For samples with plasma viral loads ≥5,000 copies/mL, add no more than 50 copies of cDNA to each 1st round PCR reaction (based on estimated copies in RNA extracted/cDNA reaction). Perform as many reactions as needed for a total of ~500 templates. Do not exceed 5uL of cDNA per reaction.
  • For samples >50c/uL by qPCR, dilute to 25c/uL and add 2uL of cDNA dilution to 16 replicate reactions.
  • For samples between 14-50c/uL, add the volume of cDNA corresponding to 50 copies total in 16 replicate reactions.
  • For samples <14c/uL, divide all of the cDNA equally into reactions (do not exceed 5ul/reaction) and repeat cDNA reaction to get as close to 800 templates as possible.

For samples with plasma viral loads <5,000 copies/mL, divide all of the cDNA equally into 12-24 replicate reactions (~2.5uL/reaction).
  • If repeating a sample and purified cDNA volume is >30uL, increase reaction number proportionally.
1st Rd PCR Mastermix (MM) recipe:
  • 5X buffer (Mg2+ 1mM final): 5 µL
  • dNTP 200 micromolar (µM) each final : 2 µL
  • Forward primer 20 micromolar (µM) : 0.3125 µL
  • PB-R1-alt1 20 micromolar (µM) : 0.3125 µL
  • PrimeSTAR GXL DNA Polymerase, 1.25U/uL: 0.5 µL
  • Nuclease-free water: QS total volume to 25uL

Add up to 5 µL of cDNA per reaction for a total reaction volume of 25 µL .

Positive control: 10 copies of 8e5 HIV DNA (be sure to spike in 0.5uL of HIV-specific reverse primer)
Negative control: nuclease-free water
2nd Rd PCR Screen (Samples with viral load <5,000 c/mL ONLY)
In order to only pool the PCR reactions that amplified in 1st round, we perform a screen to see which replicate reactions are positive.
2nd Rd PCR Mastermix (MM) recipe:
  • 5X buffer (Mg2+ 1mM final): 5 µL
  • dNTP 200 micromolar (µM) each final : 2 µL
  • Forward primer 20 micromolar (µM) : 0.3125 µL
  • PB-R2-alt1 20 micromolar (µM) : 0.3125 µL
  • PrimeSTAR GXL DNA Polymerase, 1.25U/uL: 0.5 µL
  • Nuclease-free water: 15.875 µL

Add 1 µL of 1st rd PCR product for a total reaction volume of 25 µL .
Place tubes or plate in a thermocycler and run with the PCR program conditions listed below:
ABC
CyclesTempTime
1X98C2:00
35X98C0:10
62C0:15
68C~1:00/kb
1X68C7:00
4Chold
Visualize PCR products on a 1% agarose gel, using Generule Express DNA ladder to judge amplicon size.
  • Reference Table 2 for expected sizes of full-length amplicons and positive control amplicons.
Assess results:
  • If 50% or more of the reactions are positive, move to Step 7.
  • If <50% of the reactions are positive, check in with supervisor to confirm whether the remaining plasma should be extracted to generate additional templates for sequencing before moving to Step 7.
  • If there are no positives, discuss with supervisor to decide next course of action.
2nd round PCR reactions from this screen can be discarded after the gel image and results are recorded in the processing worksheet.
Pooling 1st Round PCR Reactions
1st round replicate PCR reactions are pooled prior to purification.
For samples with viral load ≥5,000 c/ml, pool all 1st round PCR reactions into 0.5ml lo-bind tube and mix well. Label with PCR reaction code and “P” at the end to indicate pooled product.
For samples with viral load <5,000 c/ml, use the gel results from Step 6.3 to pool only the 1st round PCR reactions that have a positive band at the expected size. Label with PCR reaction code and “P” at the end to indicate pooled product.
Make 30 µL aliquot of pooled amplicon in 0.2mL PCR strip tubes for purification by BluePippin and store at 4 °C

Save control reactions for use with 2nd round. Store in 0.2ml PCR strip tubes.
Pooled 1st round PCR and control reactions stored in 4 °C if for a short term (e.g., 48 hours) and in a -20 °C freezer for longer term storage.

Purification of Pooled 1st Round PCR Products
7h 35m
The preferred method of purification is the Blue Pippin instrument as described in the following steps. If purification by Blue Pippin is not possible, bead purification can be substituted using 0.5X bead:amplicon volume ratio and eluting in 30 µL of 10mM TrisHCl, pH 8.0.
Refer to the BluePippin Quick Guide or Operations Manual for detailed descriptions.
Sample preparation:
  • Bring samples, loading solution and DNA Marker S1 to Room temperature
  • For up to 4 samples, combine 30 µL of pooled 1st round PCR with 10 µL of loading solution
  • Mix samples thoroughly, then briefly centrifuge to collect
20m
Create the following protocol on the instrument:
  • Define Cassette: 0.75% DF Low Voltage 1-6kb
  • Reference marker lane assignment 5 – Apply to all lanes
  • Select lanes for DNA capture and select “Tight” capture
  • Reference Table 2 to set Capture Size to match size of 1st round amplicon
  • Save protocol with initials and date
1m
Calibrate optics using the calibration fixture.
  • If calibration fails, shut down the machine and unplug the power cord. Leave unplugged for 30-60s and then start up the machine. Attempt calibration again.
  • Note: Multiple reset cycles of the machine may be required to fully calibrate the machine.
2m
Inspect the cassette for:
  • Buffer levels in all reservoirs is sufficient (add more buffer, if needed)
  • No breakage or bubbles in gel columns
  • If any defects in cassette, take a picture to document the issue and report to the manufacturer
1m
Prepare cassette for loading:
  • Dislodge bubbles from elution wells by tipping the sample side down
  • Place cassette into optical nest and remove adhesive strips
  • Remove all buffer from elution wells and replace with 40 µL fresh electrophoresis buffer (be careful not to introduce bubbles into elution wells as this can lead to failure of continuity test)
  • Seal elution wells with new adhesive strips
  • Check buffer levels in sample wells and top-up if needed
  • Perform continuity test. If any elution well fails, remove adhesive strip and repeat previous three steps. If the well repeatedly fails, do not use this channel.
  • Re-check buffer levels in sample wells and top-up if needed
  • Remove 40 µL buffer from each Sample Well
3m
Load samples:
  • Add 40 µL of DNA Marker S1 to sample well 5
  • Add 40 µL of pooled 1st round amplicon to sample wells 1-4. For consistency, load the PCR amplicon with the lowest reaction number in well 1 and the highest in well (e.g., ABC3P in well 1, ABC4P in well 2, ABC5P in well 3, and ABC6P in well 4)
3m
Run:
  • Close lid and press Start – the run will automatically stop when collection is complete
  • After run is complete, allow time before removing samples from elution chamber to increase recovery: At least two hours, although overnight is preferable.
7h
Collect DNA fractions – should be ~40ul per sample. Purified 1st round PCR products are stored at 4 °C for short-term and long-term in a -20 °C freezer.
5m
2nd Round PCR
1h 53m
Purified 1st round PCR products are used as template in four 25 µL 2nd rd replicate PCR reactions per sample. Use the mastermix recipe and thermocycling conditions below.
2nd Rd PCR Mastermix (MM) recipe:
  • 5X buffer (Mg2+ 1mM final): 5 µL
  • dNTP 200 micromolar (µM) each final : 2 µL
  • Forward primer 20 micromolar (µM) : 0.3125 µL
  • PB-R2-alt1 20 micromolar (µM) : 0.3125 µL
  • PrimeSTAR GXL DNA Polymerase, 1.25U/uL: 0.5 µL
  • Nuclease-free water: 15.875 µL

Add 1 µL of 1st rd PCR product for a total reaction volume of 25 µL .
8m
Place tubes or plate in a thermocycler and run with the PCR program conditions listed below:
ABC
CyclesTempTime
1X98C2:00
22X98C0:10
62C0:15
68C~1:00/kb (See Table 2)
1X68C7:00
4Chold
1h 45m
Pool 2nd round reactions together into one PCR strip tube and add "P" to the label to denote it is a pooled product. If not proceeding to step 10 directly, store at 4 °C until use.

Visualize 2nd Rd PCR Product (Agarose Gel Screen)
1h
Confirm amplification of 2nd rd PCR with a 1% agarose gel screen.
Visualize PCR products on a 1% agarose gel, using Generule Express DNA ladder to judge amplicon size.
  • Reference Table 2 for expected sizes of full-length amplicons and positive control amplicons.
  • If the gel band is faint, a replicate 2nd round PCR can be performed to ensure adequate concentration of DNA for pooling. Repeat steps 9.1-9.3.
1h
If there is no amplification, log this information in the processing worksheet and notify supervisor.
  • If sample was <5,000 c/mL viral load, and it is known that the 1st rd PCR was positive (from 2nd rd screen), repeat steps 9.1-9.3.
  • If sample was ≥5,000 c/mL viral load, perform a single 2nd round reaction as described in Step 6 (35 cycle PCR) using the pooled 1st round (unpurified) PCR product as template. If this PCR is positive, it is certain that the 1st round PCR worked to repeat Steps 8 & 9. If this PCR is negative, it is likely that there was no amplification in the 1st round PCR and the sample prep must be repeated starting from Step 5 if possible (with alternative primers) or if there is insufficient cDNA, Step 2.
2nd Round PCR Purification
41m
Notes:
  • Use Agencourt AMPure XP magnetic beads
  • Use PCR strip tubes
  • Make a fresh working stock of Room temperature 80% EtOH (320 µL per sample)
Allow AMPure XP beads to come to room temperature then add 0.7X volume to each 2nd round PCR sample (~ 95 µL of pooled amplicon + 66.5 µL of beads) and pipette-mix well.
Incubate DNA+bead mixture for 00:10:00 at Room temperature .
10m
95 µL Place the DNA+bead mixture onto magnetic rack for 00:10:00 .
10m
While on rack, carefully remove the liquid without disturbing the bead pellet.
Add 300 µL of 80% EtOH to the tube, wait 10-30 seconds, then remove, taking care not o disturb the beads.
Repeat Step 11.5 two more times for a total of three washes.
Remove traces of EtOH from the tube (quick spin in minifuge, if needed, to collect EtOH) and allow to air dry for 00:01:00 (can dry for longer, but be careful not to over-dry the pellet).
1m
Remove the tube from the magnetic rack and elute DNA from the beads using 40 µL of 10mM Tris-Cl, pH 8.0.
  • Allow elution to proceed for 00:10:00 away from magnetic rack
  • Place tube back into magnetic rack and wait 00:10:00 for beads to collect
20m
Carefully remove DNA without disturbing the bead pellet and transfer to a new 0.2mL PCR tube with "+" added to the reaction code (e.g., ABC53+) to denote purification.
Store purified 2nd round products at -20 °C .
Index PCR of Purified Amplicons
There are 96 unique combinations of Index primers that can be used to label each positive 2nd round PCR reaction. There are 24 forward primers and 4 reverse primers (Table 3).
  • These primers are based off the Illumina Nextera XT Index Kit v2 Adapters
  • The 5’ end of the Nextera adapters were removed as there is no need to include when sequenced by PacBio

Table 3: Indexing Primers
ABC
PrimerSequenceCorresponding Nextera Adapter
Index _F01 CTACACTCGCCTTATCGTCGGCAGCGTC N701
Index_F02 CTACACCTAGTACGTCGTCGGCAGCGTC N702
Index _F03 CTACACTTCTGCCTTCGTCGGCAGCGTC N703
Index_F04 CTACACGCTCAGGATCGTCGGCAGCGTC N704
Index _F05 CTACACAGGAGTCCTCGTCGGCAGCGTC N705
Index_F06 CTACACCATGCCTATCGTCGGCAGCGTC N706
Index _F07 CTACACGTAGAGAGTCGTCGGCAGCGTC N707
Index_F08 CTACACCAGCCTCGTCGTCGGCAGCGTC N710
Index _F09 CTACACTGCCTCTTTCGTCGGCAGCGTC N711
Index_F10 CTACACTCCTCTACTCGTCGGCAGCGTC N712
Index_F11 CTACACTCATGAGCTCGTCGGCAGCGTC N714
Index_F12 CTACACCCTGAGATTCGTCGGCAGCGTC N715
Index_F13 CTACACTAGCGAGTTCGTCGGCAGCGTC N716
Index_F14 CTACACGTAGCTCCTCGTCGGCAGCGTC N718
Index_F15 CTACACTACTACGCTCGTCGGCAGCGTC N719
Index_F16 CTACACAGGCTCCGTCGTCGGCAGCGTC N720
Index_F17 CTACACGCAGCGTATCGTCGGCAGCGTC N721
Index_F18 CTACACCTGCGCATTCGTCGGCAGCGTC N722
Index_F19 CTACACGAGCGCTATCGTCGGCAGCGTC N723
Index_F20 CTACACCGCTCAGTTCGTCGGCAGCGTC N724
Index_F21 CTACACGTCTTAGGTCGTCGGCAGCGTC N726
Index_F22 CTACACACTGATCGTCGTCGGCAGCGTC N727
Index_F23 CTACACTAGCTGCATCGTCGGCAGCGTC N728
Index_F24 CTACACGACGTCGATCGTCGGCAGCGTC N729
Index_R01 CGAGATCTCTCTATGTCTCGTGGGCTCGG S502
Index _R02 CGAGATTATCCTCTGTCTCGTGGGCTCGG S503
Index_R03 CGAGATGTAAGGAGGTCTCGTGGGCTCGG S505
Index_R04 CGAGATACTGCATAGTCTCGTGGGCTCGG S506
Index_R05 CGAGATAAGGAGTAGTCTCGTGGGCTCGG S507
Index_R06 CGAGATCTAAGCCTGTCTCGTGGGCTCGG S508
Index_R07 CGAGATCGTCTAATGTCTCGTGGGCTCGG S510
Index_R08 CGAGATTCTCTCCGGTCTCGTGGGCTCGG S511
Index_R09 CGAGATTCGACTAGGTCTCGTGGGCTCGG S513
Index_R10 CGAGATTTCTAGCTGTCTCGTGGGCTCGG S515
Index_R11 CGAGATCCTAGAGTGTCTCGTGGGCTCGG S516
Index_R12 CGAGATGCGTAAGAGTCTCGTGGGCTCGG S517
Index_R13 CGAGATCTATTAAGGTCTCGTGGGCTCGG S518
Index_R14 CGAGATAAGGCTATGTCTCGTGGGCTCGG S520
Index_R15 CGAGATGAGCCTTAGTCTCGTGGGCTCGG S521
Index_R16 CGAGATTTATGCGAGTCTCGTGGGCTCGG S522
Each purified 2nd round PCR should be used in only one Index PCR reaction. Plan so that a multichannel pipette can be used to easily add the purified 2nd round amplicons can be added from strip tubes or a
96-well plate to the index PCR plate.
Set up the 3rd round Index PCR.
  • Set-up the PCR master mixes, one with each Index_R# primer and aliquot 40 µL per reaction to a 96-well plate according to the plate map in the indexing and library prep processing worksheet.

Index PCR Mastermix Recipe:
  • 5X buffer (Mg2+ 1 millimolar (mM) final): 10 µL
  • dNTP, 2.5mM each (200 micromolar (µM) each final): 4 µL
  • Reverse Index Primer, 10uM (250 micromolar (µM) final):1.25 µL
  • Nuclease-free water: 31.75 µL
  • PrimeSTAR GXL DNA Polymerase, 1.25U/uL: 1 µL

Here is an example of the primer map layout for a 96-well plate.

Add 5 µL of Index_F# primers to appropriate wells with a 12-channel pipettor.
Add 5 µL of purified 2nd round PCR product to each well for a total reaction volume of 50 µL .
Place plate in a thermocycler and run with the PCR thermocycling conditions below:
ABC
CyclesTempTime
1X98C2:00
5X98C0:10
55C0:30
68C~1:00/kb (See Table 2)
1X68C7:00
4Chold
If not proceeding directly to Step 13, store at 4 °C until use.

Visualize Indexed PCR Products to Determine Relative Concentration
Using gel electrophoresis, screen amplification success and judge DNA concentration relative to DNA ladder.
Run 5 µL of indexed PCR product on a 1% agarose gel with the Generuler Express DNA ladder to judge relative concentration.

If the gel band is faint, a replicate index PCR can be performed to ensure adequate concentration of DNA for pooling. Repeat Steps 12 & 13, ensuring that the same index primer pair is used for the replicate reaction.
DNA Quantification of Indexed PCR Product
DNA quantification can be performed using the Qubit dsDNA high-sensitivity kit and protocol or the Quant-iT Picogreen dsDNA assay (which is detailed below).
Take the Picogreen reagent and Lambda standard from 4 °C and equilibrate to Room temperature in a dark place.
Refer to the gel image of the indexed PCR products and determine which bands are as bright or brighter than the 500bp DNA ladder band.
  • These samples should be diluted 1:4 in TLE to ensure the sample(s) will quantify within the standard curve of the picogreen assay (0.004-1 ng/uL) and that hte volume for pooling into the DNA library later is >1uL.
Set up an aliquot plate where:
  • less bright bands have 20 µL of undiluted indexed PCR samples added to the plate
  • bright bands have 5 µL of undiluted indexed PCR sample + 15 µL of nuclease-free water added to the plate

Keep the samples in the same order as the index PCR plate to avoid mix-ups.
In pre-PCR, prepare the appropriate volume of 1xTE (calculated by processing worksheet) with 20X concentrate provided with the kit (use nuclease-free water). Aliquot enough 1xTE into a separate tube to make the Picogreen in the next step.
Allow the Quant-iT Picogreen reagent to warm to room temperature before opening the vial (contains DMSO) and prepare a 200-fold dilution in 1xTE:
  • For example, 125ul PG reagent + 24.875ml TE → use a plastic container and protect from light to avoid photodegradation.
  • Invert to mix; do NOT vortex dyes. Use within a few hours of preparation.

Put PG aliquot inside a black glove and store in cabinet to protect from light.
Prepare DNA standard curve by making the following serial dilutions, pulse-vortexing 10X between each dilution. Transfer 750uL each dilution.



Bring standards, diluted PG reagent and remaining 1xTE to post-PCR. Place PG reagent in drawer to protect from photodegradation.
Add 100 µL of each standard to duplicate wells in rows A and B of a 96-well black fluorometer plate, as shown in example plate map below:



Add 98ul 1xTE to each non-standard well (test samples in duplicate).
Add 2ul of each sample to the corresponding duplicate wells, starting with rows C & D.
Prep fluorescence microplate reader for use (e.g., login, set sample map) BEFORE adding PG to the plate.
Using “Gemini” fluorescence microplate reader (SpectraMax, i3x, 4th floor): Open the desktop program “SoftMax Pro 6.4.2”
  • From the topscreen menu, select Protocols -> Protocol Manager -> Protocol Library -> Nucleic Acids -> PicoGreen Fluorescence
  • Select the Template Editor icon from the Plate 1 toolbar and enter in an appropriate plate set-up
  • Assign standard concentrations (ug/mL) to appropriate wells. Series -> bottom to top -> set dilution to correct value.
  • Assign a plate blank in duplicate for standard 10. Enter duplicate unknown wells for each sample. Series -> x=2, y=1. Set unknown dilution factor to 1/50 (and make sure the serial dilution factor is set to / “1” (i.e., no serial dilution)
Add 100 µL diluted PG reagent to each well with a repeater pipette and mix well (pipet mix 10+ times) using multi-channel pipette. Incubate 2-5 min at Room temperature protected from light.
Place the plate in the plate reader and click on “Read” to initiate measurement.
  • Verify that the R2 value is at least 0.98 and that sample readings fall within the standard curve.
  • Export data by selecting “Export” from the drop-down menu in the upper left corner of the plate screen (icon is 96-well plate)
  • Check box for Expt 1, and deselect intro
  • Export data in raw, plate format
  • Save as .xls file
Record the concentration of each sample and log this information in the processing worksheet.
  • If sample is above the limit of quantitation, make a dilution of the DNA and test again.
If the sample concentration is less than 0.5ng/ul repeat steps 9-11, combining the original purified sample with the new reactions before performing step 12.
Determine Amount of DNA to be Pooled in DNA Library
Note: One Revio SMRT cell can generate sufficient reads for 100,000 cDNA copies (as estimated by qPCR), so we typically pool amplicons from 100-120 plasma specimens at the most to ensure adequate sequencing depth per template.
Target at least 500ng total DNA for each pool to ensure ample material for library preparation.
  • For target of 500ng DNA, divide 500ng by the total number of estimated cDNA copies from all the samples in that pool (e.g., 500ng DNA / 9000 copies in pool 1 = 0.056ng DNA / copy)
For each sample, multiply the number of estimated copies of each sample by the mass of DNA per copy determined in Step 15.1 to determine the mass of DNA needed per sample.
  • e.g., 300 copies Sample A x 0.056 ng/copy = 16.8ng DNA needed
Determine the volume of purified DNA to pool for each sample by dividing the mass of DNA needed from Step 15.2 by the concentration of the purified DNA from Step 14.15.
  • e.g., 8ng DNA needed / 10ng/ul DNA concentration = 1.68ul volume required
  • When pooling, samples with different amplicon sizes must be accounted for to ensure equimolar ratios. Consult Table 2 for 2nd round PCR amplicon sizes and reduce input proportionally for smaller amplicons.
If sample does not have sufficient volume, repeat steps 9-11, combining the original purified sample with the new reactions before step 12 and repeating the calculations in Step 15.3.
DNA Library Pooling & Purification
20m 10s
Notes:
  • Use Agencourt AMPure XP beads
  • Use low-bind 1.5mL microcentrifuge tubes
  • Make a fresh stock of room temperature 80% EtOH (3mL per pool)
For each pool, combine all samples into 1.5ml tube using the volumes determined in Step 15.3.
00:00:00 Mix each DNA pool with 0.6x volume of AMpure Beads and incubate for 00:10:00 .
10m
Place the DNA + Bead mixture into the magnetic rack for 00:10:00 .
10m
While in the rack, carefully remove the liquid without disturbing the pellet of beads.
Add 1 mL of 80% EtOH to the tube, wait 00:00:10 , then remove, taking care not to disturb the beads.
10s
Repeat Step 16.5 two more times for a total of three washes.
Remove traces of EtOH from the tube (quick spin in minifuge, if needed, to collect EtOH) and allow to air dry for 00:01:00 (can dry for longer, but be careful not to over-dry the pellet).
Remove the tube from the magnetic rack and elute DNA from the beads using 50 µL of 10mM Tris-Cl, pH 8.0.
  • Allow elution to proceed for 00:10:00 away from magnetic rack
  • Place tube back into magnetic rack and wait 00:10:00 for beads to collect
Carefully remove DNA without disturbing the bead pellet and transfer to a new 1.5mL PCR tube and label with pool name and "+" to denote purification.
Store pooled libraries temporarily at 4 °C and then long-term at -20 °C .
DNA Quantification of Pooled Libraries
2m
To verify the amount of DNA in each pool -- important for the SMRTbell Template Prep step -- we quantify the DNA with the Qubit dsDNA HS kit.
Allow 1x dsDNA HS Working Solution, standards, and purified samples from Step 16 to equilibrate to Room temperature Note: 1x dsDNA HS Working Solution should be protected from light.
Set out two more tubes than the number of samples you wish to examine, these will be used for standards.
  • Label one tube S1 and one S2
  • Number or label the tubes for your samples
  • Only mark the tops of the tubes, not the sides
Add 190 µL of the 1x dsDNA HS Working Solution to S1 and S2 tubes.
  • Add 10 µL of Standard 1 to tube S1 and 10 µL of Standard 2 to tube S2
Add 198 µL of the 1x dsDNA HS Working Solution to each sample tube.
  • Add 2 µL of each purified sample from Step 16 to corresponding tube
Mix each sample by vortexing for 3–5 seconds.
Allow all tubes to incubate at Room temperature for 00:02:00 .
2m
On the Home screen of the Qubit select DNA, then 1X dsDNA HS. Press Read Standards to proceed.
Insert Standard S1 and then Standard S2 as prompted.
Select Run Samples and select 2 µL as the sample volume.
  • Record the concentration of each sample and log this information in the Specimen Preparation worksheet.
  • If sample is above the limit of quantitation, make a 10x dilution of the DNA and test again.
DNA concentration should be above 10ng/ul.
  • If DNA concentration is below 10ng/ul, but above 5ng/ul, repeat Step 16 with only one wash step and elute into 15ul 10mM Tris-HCl, pH 8.0
  • If DNA concentration is below 5ng/ul, repeat Step 16 with greater sample volumes from 15.3 to generate more DNA
Preparing DNA Libraries for PacBio Sequencing
Follow the PacBio SOP "Preparing multiplexed amplicon libraries using SMRTbell prep kit 3.0" to add SMRTbell adapters to the DNA libraries and pool everything into one library for submission to the sequencing core.

https://www.pacb.com/wp-content/uploads/Procedure-checklist-Preparing-multiplexed-amplicon-libraries-using-SMRTbell-prep-kit-3.0.pdf