Integration of Wet and Dry Bench Processes Optimizes Targeted Next-generation Sequencing of Low-quality and Low-quantity Tumor Biopsies

Note: This protocol describes the simultaneous processing of samples using a MiSeq NGS System but can be adapted for the Personal Genome Machine (PGM) instrument. For the recommended minimum DNA input of 400 amplifiable template copies, the assay is capable of producing at least 3,000x median coverage for each of 96 samples per NGS run, and equivalent coverage depth for 24 samples using the PGM on a 318 chip. The method also requires the use of a real-time PCR instrument.

1. DNA Functional Quantification and Quality Control (QC)

1. Thaw reagents: 2x Master Mix, Primer Probe Mix, Inhibition Primer Probe Mix, 6-carboxy-X-rhodamine (ROX), Diluent, and the four human genomic DNA calibration curve standards (DNA Standard (50 ng/µl), DNA Standard (10 ng/µl), DNA Standard (2 ng/µl), and DNA Standard (0.4 ng/µl)) (Table 1). Vortex all reagents for 10 sec and centrifuge at maximum speed for 10 sec to collect contents. Keep the 2x master mix on ice.
2. Prepare a sufficient amount of master mix for the total number of samples to be tested and include 10% more volume to avoid shortages due to pipetting. Prepare the master mix in a microcentrifuge tube using the following volumes per sample: 5 µl 2x Master Mix, 0.5 µl Primer Probe Mix, 0.5 µl Inhibition Primer Probe Mix, 0.05 µl ROX and 2.95 µl Diluent. Vortex for 10 sec and centrifuge at maximum speed for 10 sec to collect contents.
3. Add 9 µl master mix into wells of a 96-well plate.
4. Add 1 µl of the DNA Standards in duplicate to generate a calibration curve. Mix by pipetting up-and-down 5 times.
5. Ensure the nucleic acid sample is well mixed before use. Add 1 µl sample to the master mix and mix by pipetting up-and-down 5 times.
6. Seal the plate, vortex for 10 sec and centrifuge at maximum speed for 10 sec to collect contents.
7. Place the plate into the PCR System. Assign both FAM (functional quantification) and VIC (functional inhibition) detectors for each sample according to the manufacturer's instructions. Perform PCR cycles of 10 min at 95 °C, and 40 cycles (15 sec at 95 °C, 1 min at 60 °C).
8. Analyze the qPCR data by generating a linear regression plot for each of the duplicate DNA standards using software protocols.
9. Plot the Log₁₀ of the copy number for each DNA standard on the x-axis and the corresponding FAM C_q value on the y-axis.
10. Confirm that the results from the nucleic acid sample fall within the dynamic range of the DNA Standards calibration curve, and then calculate the concentration of the unknown DNA in "functional" or amplifiable copy number per µl from its corresponding position on the reference standard curve. Figure 1 shows examples of calibration curves that passed and failed.
11. Determine if amplification occurred in each reaction by checking for the presence of the non-human target amplicon in the VIC channel.
    Note: As a positive control, the Inhibition Primer Probe Mix contains primers specific for a non-human exogenous target and the corresponding template. The Inhibition Primer Probe Mix is a component of the master mix which is added to each reaction, including the no-template control (NTC). In absence of an inhibitor, the PCR product for the non-human target should always be detected in the VIC channel. An "undetected" C_q for a sample in the VIC channel indicates the presence of PCR inhibitors that may benefit from subsequent clean-up of the sample prior to further processing.

2. Library Preparation: Gene-specific (GS) PCR

1. Prepare a sufficient amount of master mix for the total number of samples to be tested and include 10% more volume to avoid shortages due to pipetting. Prepare the GS PCR master mix in a microcentrifuge tube using the following volumes per sample: 5 µl 2x Amplification Master Mix (Table 1) and 1 µl Pan Cancer Primer Panel (Table 1). Mix by pipetting up-and-down, vortex for 10 sec and centrifuge at maximum speed for 10 sec to collect contents.
2. Aliquot the 6 µl GS PCR master mix into wells of a 96-well plate. Add 4 µl of each nucleic acid sample into individual wells. To other wells, add 4 µl of the FFPE Control (Table 1), 4 µl of the Multi-Variant control (Table 1), and 4 µl of nuclease-free water for a procedural NTC. For each addition, mix by pipetting up-and-down 5 times.
3. Seal the plate, vortex for 10 sec and centrifuge at maximum speed for 10 sec to collect contents.
4. Place the plate in the thermocycler for the following PCR cycles: 5 min at 95 °C, 2 cycles (15 sec at 95 °C, 4 min at 60 °C), 23 cycles (15 sec at 95 °C, 4 min at 72 °C), and a final extension of 10 min at 72 °C. Hold at 4 °C.
   Note: After completion of step 2.4, the plate will be referred to as the GS PCR plate.

3. Library Preparation: Tag PCR

1. Thaw reagents: 2x Index Master Mix (Table 1) and Index Codes (Table 1). Vortex for 10 sec and centrifuge at maximum speed for 10 sec to collect contents.
   Note: Index Codes are premixed to provide a unique set of pairwise indices (barcodes) for each sample.
2. In a 96-well plate, add 7.5 µl of the 2x Index Master Mix and 5.5 µl of an Index Code to a specified well and mix by pipetting up-and-down 5 times.
3. Carefully open the GS PCR plate, and add 2 µL GS PCR product to the new plate with the master mix. Mix by pipetting up-and-down 5 times. For each sample, record the Sample ID and the corresponding pairwise Index Codes. Seal the plate, vortex for 10 sec and centrifuge at maximum speed for 10 sec to collect contents.
4. Place the plate in the thermocycler and PCR for 5 min at 95 °C, 10 cycles (30 sec at 95 °C, 30 sec at 55 °C, 1 min at 72 °C), and a final extension of 10 min at 72 °C. Hold at 4 °C.
   Note: After completion of step 3.4, the plate will be referred to as the Tag PCR plate.

4. Library Purification and Size Selection

1. Remove the Library Pure Prep magnetic beads (Table 1) and Elution Buffer (Table 1) from 2-8 °C and allow to equilibrate to RT for 30 min. Add 9.6 ml 100% ethanol to the Wash Buffer (Table 1) container, cap and mix by inverting the bottle several times.
2. Vortex the magnetic beads for 10 sec and add 11 µl into separate wells of a 96-well plate.
3. Open the Tag PCR plate and add 10 µl of Tag PCR product to the beads and pipet mix 5 times. Incubate the mixture for 4 min at RT.
4. Place the 96-well plate on the magnetic stand (Table 1) for 4 min. With the 96-well plate still on the stand, remove and discard the supernatant with a pipette.
5. Remove the 96-well plate from the magnetic stand and add 100 µl ethanol-containing Wash Buffer to each well and mix by pipetting up-and-down 5 times. Incubate for 2 min.
6. Place the 96-well plate on the magnetic stand for 2 min, then remove and discard the supernatant with a pipette.
7. Repeat step 4.5, for a total of 2 ethanol washes, removing as much wash solution possible after the second wash.
8. With the 96-well plate on the magnetic stand, dry the beads for 2 min at RT, then remove the plate from the stand.
9. Resuspend the beads by adding 20 µl Elution Buffer to each well and pipet up-and-down 5 times.
10. Incubate for 2 min at RT.
11. Place the 96-well plate back on the magnetic stand for 4 min, and carefully remove and transfer 18 µl of the clear supernatant to a new well.
    Note: The procedure may be safely stopped at this step and samples stored at -15 to -30°C. To restart, thaw frozen samples on ice before proceeding.

5. Library Quantification

1. Thaw Library Quant (LQ) reagents: 2x LQ Master Mix, LQ Primer / Probe Mix, LQ Standard, LQ Positive Control, LQ Diluent, and LQ ROX (Table 1). Vortex for 10 sec and centrifuge at maximum speed for 10 sec to collect contents.
2. Using purified library products, perform a serial dilution of each individual sample in LQ Diluent.
   1. Add 2 µl (library purified product) to 198 µl LQ Diluent and mix up-and-down with a pipet 10 times.
   2. Add 2 µl (1:100 dilution) to 198 µl LQ Diluent and mix up-and-down with a pipet 10 times.
3. Prepare a sufficient amount of LQ master mix for the total number of samples to be tested and include 10% more volume to avoid shortages due to pipetting. Prepare the LQ Master Mix in a microcentrifuge tube using the following volumes per sample: 5 µl 2X LQ Master Mix, 2 µl LQ Primer/Probe Mix, 0.5 µl LQ Standard and 0.5 µl LQ ROX. Mix by pipetting up-and-down, vortex for 10 sec and centrifuge at maximum speed for 10 sec to collect contents.
4. Add 8 µl LQ master mix to a well of an optical 96-well plate.
5. In separate wells, add 2 µl diluted library, 2 µl LQ Positive Control and 2 µl LQ Diluent (NTC) and mix by pipetting up-and-down 5 times. Seal the plate with optical adhesive film, vortex for 10 sec and centrifuge at 400 x g for 10 sec to collect contents.
6. Assign both FAM and VIC detectors for each sample. Perform PCR amplification using cycling conditions of 5 min at 95 °C, and 40 cycles (15 sec at 95 °C, 1 min at 60 °C).
7. Determine the concentration of each sample (nM) using the comparative C_q method. Calculate the difference of the known LQ Standard (C_q VIC) to the unknown library (C_q FAM). The concentration of the Diluted Sample (pM) is calculated using the following equation:
      [Conc_lib]_nM = 12.5 x 2^ΔCq
   If using a dilution factor other than 10,000, calculate the ratio of the target dilution factor to 10,000, and multiply this factor by the result of the equation in 5.7.

6. Library Normalization and Sample Pooling

1. Determine the median concentration (nM) across all samples (each containing a unique pairwise index) to be pooled.
2. Determine the individual sample volume (µl) to pool by multiplying the median concentration across all samples by 5, then dividing by its individual concentration (nM). Round the resulting value to the nearest integer. Round volumes with values of <2 µl to 2 µl, and volumes >15 µl to 15 µl.
3. Add the normalized volume (µl) for each sample to a single microcentrifuge tube to create the sample pool.
4. Calculate the new concentration for each sample using rounded integer values and record the results.
5. To determine the concentration of the sample pool, calculate the sum of all individual concentrations and record the resulting value (nM).
6. Dilute the sample pool to 1.25 nM using Sequencing Diluent (Table 1).
   Note: The procedure may be safely stopped at this step and samples stored at -15 to -30 °C. To restart, thaw frozen samples on ice before proceeding.

7. Sequencing

1. Denature the sample pool in the presence of PhiX Control v3 (Table 1) by adding the following: 15 µl of 1.25 nM Sample Pool, 3 µl of 0.5 nM PhiX and 2 µl of 1 N NaOH. Vortex briefly followed by a brief centrifugation and incubate for 5 min at RT.
2. Place the denatured sample pool on ice.
3. Add 8 µl of denatured library to 992 µl of pre-chilled HT1-Hyb buffer to a microcentrifuge tube. Vortex briefly to mix, followed by a brief centrifugation to collect contents. Keep on ice.
4. Add 600 µl of the denatured and diluted library to position #17 of the reagent cartridge.
5. Thaw Read 1 Sequencing Primers (Table 1), Index Read Sequencing Primers (Table 1), and Read 2 Sequencing Primers (Table 1). In microcentrifuge tubes, separately dilute 4 µl Sequencing Primers with 636 µl HT1-Hyb buffer. Note: HT1-Hyb buffer is provided with the sequencing reagent kit (Table 1).
6. Mix by vortexing for 10 sec and centrifuge at maximum speed for 10 sec to collect contents. Add 600 µl of diluted Read 1 Sequencing Primers to position #18 of the sequencing reagent cartridge, 600 µl of diluted Index Read Primers to position #19, and 600 µl of diluted Read 2 Sequencing Primers to position #20.
7. Load the reagents on the NGS instrument (Table 1) and sequence according to the manufacturer's instructions. Perform a paired-end 2 x 150 cycle sequencing run.

8. Data Analysis

Note: The NGS instrument software converts cluster images to base calls and quality scores, and demultiplexes pairwise indices to generate individual gzip-compressed FASTQ (*.fastq.gz) files for each sample. Prior to analyzing the demultiplexed files, the reader must download and install the associated bioinformatics software (Table 1). The software can be installed on a consumer-grade Windows PC and does not require specialized computing hardware or an internet connection to perform the data analysis.

1. Double-click the software desktop icon.
2. Login to the system using the username and password provided in the software manual.
3. Open the project dashboard, and click "New Project".
   1. Name the project and provide an optional project description. Select the targeted NGS panel type and NGS instrument type. Click "Save and continue".
   2. Upload the compressed FASTQ files for the forward and reverse reads. Do not upload the "unassigned" FASTQs, which contain reads that failed to demultiplex. Click "Save and continue".
   3. Input the number of functional input copies used to prepare each library as determined by the DNA functional quantification assay (see Step 1 above). Manually add values or copy and paste values from a spreadsheet into the annotation table. Click "Save and continue."
   4. Review the annotated libraries uploaded for analysis and click "Submit analysis" to initiate the analysis.
4. Monitor the progress of the analysis displayed through the project dashboard.
   Note: An analysis complete status indication is presented when the results are ready for review.
5. Review the analyzed results for sample QC metrics including total coverage per library, percentage of reads passing filters, amplicon coverage depth and uniformity. Review the variant calls for each sequenced library with dbSNP, COSMIC, 1,000 genomes and other sources of functional and population level annotation.
6. Export the raw results as summary spreadsheet tables, *.bam files and *.vcf files for long-term storage or downstream analysis with complementary informatics tools.