Amplification of Near Full-length HIV-1 Proviruses for Next-Generation Sequencing

All methods described here have been approved by the institutional review boards at the University of California San Francisco and the Western Sydney Local Health District, which includes The Westmead Institute for Medical Research.

1. Lysis of HIV-1-infected Cells

NOTE: Cells may be isolated from peripheral blood, leukapheresis samples, bone marrow biopsy, or tissue biopsy. Cell populations may be sorted using fluorescence-activated cell sorting (FACS).

1. Prepare a lysis buffer containing 10 mM Tris-HCl, 0.5% Nonidet P–40, 0.5% Tween-20, and 0.3 mg/mL proteinase K. To a cell pellet, add 100 µL of lysis buffer per 1 x 10⁶ cells. Pipette up and down to mix. Incubate at 55 °C for 1 hour followed by 85 °C for 15 min to lyse the cells and release genomic DNA for PCR amplification.
   NOTE: Cell lysis is sufficient to obtain genomic DNA for amplification. No DNA isolation or purification is required. The protocol can be paused at this point and genomic DNA can be stored at -20 °C indefinitely.

2. Amplification of Single HIV-1 DNA Proviruses via Nested PCR

1. Mix the reagents for the first round PCR (PCR1) listed in Table 1 (see Table of Materials). Add 38 µL of master mix to 85 wells (80 samples, 4 negative controls, 1 positive control) of a 96-well PCR plate (follow the layout in Figure 1B). Designate this plate ‘PCR1’.

Table 1: Reagents and volumes for PCR master mixes.

NOTE: The primers used for PCR1 are:
BLOuterF: 5’-AAATCTCTAGCAGTGGCGCCCGAACAG-3’ (HXB2 position 623-649)
BLOuterR: 5’-TGAGGGATCTCTAGTTACCAGAGTC-3’ (HXB2 position 9662-9686)

2. After preparing the master mix, move the PCR1 plate to a clean area designated for the addition of genomic DNA.
   1. Serially dilute genomic DNA from 1:3 to 1:81 with Tris-HCl (5 mM, pH 8), preparing 45 µL for each dilution (enough for 20 wells for each dilution). Add 2 µL of diluted genomic DNA to each sample well and 2 µL of Tris-HCl (5 mM, pH 8) to each negative control well (follow the layout in Figure 1B). Seal all the wells of the PCR1 plate, excluding the positive control well, using a clear adhesive seal (see Table of Materials).
      Note: The above recommended dilutions serve only as a starting guide for determining the end-point dilution. Dilutions will depend on the concentration of integrated HIV-1 DNA within the samples.
3. Move to an area designated for the addition of positive control. Add 2 µL of positive control (pNL4-3 diluted to 10⁵ copies/µL) to the positive control well of the PCR1 plate and seal the plate. Briefly spin the PCR1 plate in a PCR plate spinner or centrifuge (400 x g for 10 s at room temperature) to pull down any residual contents from the sides of the wells.
4. Run the PCR1 plate in a thermocycler: 94 °C for 2 min; then 94 °C for 30 s, 64 °C for 30 s, 68 °C for 10 min for 3 cycles; 94 °C for 30 s, 61 °C for 30 s, 68 °C for 10 min for 3 cycles; 94 °C for 30 s, 58 °C for 30 s, 68 °C for 10 min for 3 cycles; 94 °C for 30 s, 55 °C for 30 s, 68 °C for 10 min for 21 cycles; then 68 °C for 10 min (30 cycles total). Hold at 4 °C.
   NOTE: The protocol can be paused here and the PCR1 plate kept at 4 °C for up to 2 days.
5. Mix the reagents for the second round of PCR (PCR2) listed in Table 1. Add 28 µL to 85 wells (80 samples, 4 negative controls, 1 positive control) of a new 96-well PCR plate (follow the layout in Figure 1B). Designate this plate ‘PCR2’.

NOTE: The primers used for PCR2 are:
275F: 5’-ACAGGGACCTGAAAGCGAAAG-3’ (HXB2 position 646-666)
280R: 5’-CTAGTTACCAGAGTCACACAACAGACG-3’ (HXB2 position 9650-9676)

6. Briefly spin the PCR1 plate in a PCR plate spinner or centrifuge (400 x g for 10 s at room temperature) to pull down any residual contents from the sides of the wells. Add 80 µL of Tris-HCl (5 mM, pH 8) to each well of the PCR1 plate.
   1. Transfer 2 µL of the PCR1 plate to the PCR2 plate using a multichannel pipette. Ensure samples are transferred well to well (i.e., 2 µL from well A1 of PCR1 plate is transferred to well A1 of PCR2 plate). Seal the PCR2 plate using a clear adhesive seal (see Table of Materials).
   2. Briefly spin the PCR2 plate in a PCR plate spinner or centrifuge (400 x g for 10 s at room temperature) to pull down any residual contents from the sides of the wells. Seal the PCR1 plate with a heat sealing film for long term storage at -20 °C (see Table of Materials).
7. Run the PCR2 plate in a thermocycler: 94 °C for 2 min; then 94 °C for 30 s, 64 °C for 30 s, 68 °C for 10 min for 3 cycles; 94 °C for 30 s, 61 °C for 30 s, 68 °C for 10 min for 3 cycles; 94 °C for 30 s, 58 °C for 30 s, 68 °C for 10 min for 3 cycles; 94 °C for 30 s, 55 °C for 30 s, 68 °C for 10 min for 31 cycles; then 68 °C for 10 min (40 cycles total). Hold at 4 °C.
   NOTE: The protocol can be paused here and the PCR2 plate kept at 4 °C for up to 2 days.
8. Briefly spin the PCR2 plate in a PCR plate spinner or centrifuge to pull down any residual contents from the sides of the wells. Add 60 µL of Tris-HCl (5 mM, pH 8) to each well using a multichannel pipette.
   1. Run 15 µL of each well on 2 x 48-well precast 1% agarose gels containing ethidium bromide (0.1‒0.3 µg/mL, see Table of Materials). Use a ladder with a range up to 10 kb (see Table of Materials). Visualize to identify the wells containing the amplified product and their approximate sizes. Save the gel image.
      NOTE: Ensure the positive control well contains amplified product. If the negative control wells contain amplified product, consider contamination and disregard plate.
9. Determine the dilution at which no more than 30% of wells are positive for amplified product.
   NOTE: This is the end-point dilution in which a majority (80%) of wells contain amplified product from a single template. This dilution should be prepared and used in subsequent PCRs to obtain further proviral amplicons. Record the approximate size of each amplified product.

3. DNA Purification and Quantification

1. Briefly spin the PCR2 plate in a PCR plate spinner or centrifuge to pull down any residual contents from the sides of the wells. Transfer 40 µL from the wells containing amplified product (at or below the end-point dilution) to a new 96-well midi plate (with a well volume of 0.8 mL, see Table of Materials).
   NOTE: Write down the original and new well position of each amplified product in a spreadsheet as a record of each amplified product to be sequenced.
2. Purify the amplified DNA products using a magnetic bead based PCR purification kit (see Table of Materials) to remove primers, nucleotides, enzymes, oils and salts. Before starting, bring magnetic beads to room temperature and prepare fresh 80% ethanol. Use new pipette tips when appropriate to avoid cross contamination of DNA samples.
   1. Vortex magnetic beads to ensure they are thoroughly resuspended. Using a multichannel pipette, add 40 µL of beads to the 40 µL of amplified product in the 0.8 mL 96-well midi plate. Gently pipette up and down 10 times to mix. Alternatively, mix the solution by sealing the plate then shaking on a microplate shaker at 1,800 rpm for 2 min. Incubate at room temperature for 5 min.
   2. Place the plate on a magnetic stand (see Table of Materials) for 2 min. Remove and discard supernatant.
   3. Wash beads by adding 200 µL of 80% ethanol to each well with the plate on the stand. Incubate at room temperature for 30 s. Remove and discard the supernatant. Repeat once. Using a multichannel pipette with fine tips, remove any excess ethanol following the second wash. With the plate on the stand, allow the beads to air dry for 15 min.
   4. Remove the plate from the stand. Add 30 µL of elution buffer (see Table of Materials) to each well. Gently pipette up and down 10 times to mix. Alternatively, mix the solution by sealing the plate then shaking on a microplate shaker at 1,800 rpm for 2 min. Incubate at room temperature for 2 min.
   5. Place the plate on a magnetic stand for 2 min. Using a multichannel pipette, transfer 25 µL of supernatant (purified DNA) to a new 96-well PCR plate. Ensure that the samples are transferred well-to-well (i.e., supernatant of well A1 in 0.8 mL plate is transferred to well A1 of new 96-well plate).
      NOTE: 5 uL of the eluted sample is left behind to ensure no transfer of residual purification beads.
3. Determine and record the approximate concentration of each amplified DNA product following purification using a spectrophotometer (absorbance at a wavelength of 260 nm).
   NOTE: Measuring the approximate concentration of each amplified product is necessary at this stage to ensure no samples are lost during the purification steps and that the approximate concentration is within the range of the standard curve used in the next stage (0.001–1 ng/µL DNA in 100 µL of volume). The protocol can be paused here and cleaned samples can be stored at -20 °C for up to 6 months.
4. Quantify the concentration of DNA of each purified product using a dsDNA quantification kit (see Table of Materials).
   NOTE: This involves using a standard curve to determine the concentration of dsDNA in a sample. The kit includes buffer (10 mM Tris-HCl, 1 mM EDTA, pH 7.5), a lamda dsDNA standard, and a fluorescent dye. Keep all reagents on ice. Cover the tube containing dye with foil to avoid exposure to light. Measure the concentration of each amplified product in triplicate.
   1. Dilute buffer to 1x in sterile DNase free H₂O. Add 99 µL of buffer to an appropriate number of empty wells (3 times the number of amplified products to be measured, see Table 2 for an example layout) of a flat bottom tissue culture plate. Add 100 µL of buffer to 3 blank wells.
   2. For each amplified product to be measured, add 1 µL of purified DNA (from step 3.2.5) in triplicate to each well containing buffer (see Table 2 for an example layout).
   3. To prepare standards, dilute lamda dsDNA 10-fold from 2 ng/µL to 0.002 ng/µL. Add 100 µL of each dsDNA standard to 3 wells.
      NOTE: Following step 3.4.4, the final concentration of the standards will range from 1 to 0.001 ng/µL
   4. Dilute the fluorescent dye 1:200 with buffer. Quickly add 100 µL to each well containing sample, blanks, and standards. Mix up and down with a pipette. Cover the plate with foil to avoid contact with light.
   5. Read fluorescence emission on a microplate reader (excitation at 480 nm, emission at 520 nm). Record results in a spreadsheet.
   6. Determine the concentration of dsDNA in each sample using the fluorescence measurements recorded. Subtract the fluorescence measured in the blank wells from the sample and standards. Determine the average fluorescence for each sample and standard from the triplicates. Draw a standard curve based on the fluorescence measurements of the standards. Determine the concentration of the samples relative to the standards.

Table 2: Example layout of 96-well plate for quantification of dsDNA.

5. Dilute each purified product (obtained in step 3.2.5) with H₂O to 0.2 ng/µL.

4. Sequencing Library Preparation

1. Prepare amplified and purified proviral DNA for NGS using an NGS DNA library preparation kit (see Table of Materials). Follow the manufacturer’s instructions for all tagmentation, PCR amplification, and clean-up steps [except that the reaction volumes including input DNA (from step 3.5) can be halved to extend the use of library preparation reagents].
2. Normalize libraries manually using a qPCR-based NGS library quantification kit (see Table of Materials) to determine the individual concentration of provirus. Combine individual provirus libraries in equimolar amounts to a final concentration of 4 nM, or as specified by the sequencing service provider.
   NOTE: The protocol can be paused here and the pooled library stored at -20 °C.
3. Quantify the final pooled library using the same dsDNA quantification kit used in step 3.4. Follow the manufacturer’s instructions. Determine the average fragment lengths by running 1 µL of the pooled library on an automated electrophoresis system using an appropriate kit (see the Table of Materials). Use the concentration and average fragment lengths to determine the molarity of the pooled library.
4. Combine 5 µL of the pooled library with 5 µL of 0.2 N NaOH to denature the library. Add 5 µL of 200 mM Tris-HCl to neutralize. Dilute the final library to 12.5 pM with chilled hybridization buffer (available with DNA library preparation kit) immediately prior to sequencing.
5. Perform 2 x 150 nucleotide (nt) paired-end sequencing on an appropriate NGS platform (see Table of Materials).
   NOTE: When 96 proviral libraries are indexed per run, this yields approximately 20 million paired-end reads per run or 200,000 reads per individual provirus library for analysis following de-multiplexing. Steps 4.4 and 4.5 are typically performed by a sequencing facility.

5. De Novo Assembly of Sequenced HIV-1 Proviruses

NOTE: To obtain the genetic sequence of each amplified provirus, contigs are assembled de novo from the paired-end reads. Many platforms (e.g., CLC Genomics Workbench¹⁴), allow the design of custom workflows for de novo assembly. Other open source software such as FastQC¹⁵, Trimmomatic¹⁶, Cutadapt¹⁷, and FLASH¹⁸ can also be utilized for processing reads, as well as tools such as Bowtie2¹⁹ and SPAdes²⁰ for read mapping and de novo assembly. The steps for the de novo assembly of HIV-1 contigs using a specific commercial platform (see the Table of Materials) are outlined below (Figure 2). Customized workflow file is available upon request.

1. Import sequences and check quality: Import the paired sequence reads (in fastq.gz format) into the software, which will then be combined as single set of paired reads. Generate a sequence QC report and examine the quality of your data.
2. Quality control
   1. Perform read trimming according to the QC report. Remove any adapter sequences and ambiguous nucleotides. Trim fifteen 5’ and two 3’ terminal nucleotides. Discard reads less than 50 nucleotides in length. Use a stringent quality limit of 0.001 corresponding to a QC phred score of 30.
3. Merging overlapping pairs
   1. Form single extended reads by merging paired forward and reverse reads with overlapping regions.
4. De novo assembly
   1. Use the native CLC genomics de novo assembler with a word (or k-mer) size of 30 nt and a maximum bubble size of 65 nt to assemble a random subsample of 10,000 non-overlapping paired reads. The expected coverage for the de novo assembled contig is ~200x for a ~9 kb sequence.
      NOTE: This subsampling can reduce the computational burden such that most standard desktop computers can handle the assembly and analysis, and given the clonality of each provirus (one library is one provirus), this does not limit the diversity.
5. Re-mapping all reads to contigs
   1. To obtain the final majority consensus sequence of each provirus, map the full read set to the de novo assembled contig. Accept only contigs with a minimum average coverage of 1,000x and ensure the final contig length corresponds to the size of the band on the original agarose gel (step 2.8.1). Save the final majority consensus sequence as a .fasta file.
      NOTE: Most contigs can be assembled using the above steps. However, in some cases, for a single provirus, multiple contigs with a similar coverage are assembled. In these circumstances, contigs are aligned to a near full-length (~9 kb) consensus sequence from the same participant and manually assembled into a single contig. All reads are then mapped to the manually assembled contig and the final consensus accepted if read coverage is even throughout the assembly and no single nucleotide polymorphisms (SNPs) > 40% are present.
6. Further quality control
   1. To ensure each contig represents a single provirus, and is not due to the amplification of multiple proviruses within a single well, screen read coverage and variant calling of the final contig.
   2. Multiple proviral templates present during PCR are often identified by very uneven read coverage (due to the co-amplification of proviruses of different sizes) when mapping to a full-length consensus from the same participant, or by the presence of SNPs with a frequency of >40% (see representative results, Figure 4). Disregard mixed populations in subsequent analyses.
7. Alignment
   1. Import the final consensus of each proviral sequence into sequence viewing software such as Molecular Evolutionary Genetics Analysis (MEGA) 7²¹. Align each sequence manually to the HXB2 reference sequence. Trim the 5’ and 3’ ends to positions 666-9650 of HXB2 to remove primer sequences.
   2. Export the sequence list in fasta format and then align using MAFFT version 7²², with manual editing where appropriate to obtain the final alignment.
      NOTE: If any sequences do not align, first reverse complement the sequence. If the sequence still does not align perform a BLAST search to ensure the sequence is HIV-1. It is possible to amplify non-HIV-1 templates and these can be identified at this stage. If sequences are HIV-1 but do not align, consider the presence of inversions (see step 6.1).

6. Determining Potential Replication Competency of HIV-1 Proviral Sequences

NOTE: To identify sequences of genetically intact, and potentially replication-competent, HIV-1 proviruses a stringent process of elimination is followed (Figure 1C). Proviral sequences lacking inversions, large internal deletions, deleterious stop codons/ hypermutation, frameshift mutations, and/or defects in the MSD site or packing signal are considered genetically intact and potentially replication-competent.

1. Inversions
   1. During the alignment stage, identify inversions. Inversions are regions where the sequence does not align to the reference HXB2 unless the region is reverse complemented.
      NOTE: Depending on the application of the data, sequences containing inversions may need to be omitted from further analysis.
2. Large internal deletions
   1. Identify contigs with large internal deletions (>600 bp) in the alignment stage. Unless the deletion sits within nef, the sequence can be defined as defective.
      NOTE: Any sequences with internal deletions <600 bp will be identified by Gene Cutter (step 6.3.1) as having incomplete gene sequences.
3. Deleterious stop codons, frameshift mutations and deletions
   1. Check all contigs of length >8400 nucleotides for the presence of deleterious stop codons, frameshifts and incomplete gene sequences using the Los Alamos National Laboratory HIV Sequence Database Gene Cutter tool²³.
      NOTE: The Gene Cutter tool divides the proviral sequence into the genes gag, pol, vif, vpr, tat, rev, vpu, env, and nef, and translates them to amino acids. Gene Cutter then screens for the presence of stop codons and frameshift mutations (due to insertions or deletions). Contigs containing stop codons or frameshifts in any gene, excluding nef²⁴, are classified as defective. Gene Cutter also identifies incomplete gene sequences and any proviruses with a deletion <600 bp in a gene other than nef can be reclassified as defective due to a large internal deletion.
4. Hypermutation
   1. Generate a consensus of the remaining full-length proviral sequences using the Los Alamos National Laboratory HIV Sequence Database Consensus Maker tool (simple consensus maker)²⁵. Add the consensus sequence to the top of an alignment containing only the remaining full-length proviral sequences. Using this alignment and the Los Alamos National Laboratory HIV Sequence Database Hypermut tool²⁶ to identify APOBEC-induced G-A hypermutation in the remaining full-length proviral sequences.
5. Defects in the MSD and packaging signal
   1. Inspect each of the remaining contigs for defects in the MSD and the packaging signal (HXB2 region 670-810) which render the proviral sequence defective⁴. Look for a point mutation in the MSD site (sequence GT, HXB2 744-745) or any deletion in the four stem loops of the packaging signal (SL1 (HXB2 691-734), SL2 (HXB2 736-754), SL3 (HXB2 766-779), and SL4 (HXB2 790-810)).