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

The overall goal of this procedure is to describe a comprehensive system for sequencing challenging cancer specimens that combines wet lab reagents, standardized controls, and a stand along bioinformatics suite. This method can help answer key questions in the cancer field, such as how to sequence low quantity and low quality tumor biopsies to achieve accurate DNA variant analyses and interpretations. The main advantage of this method is that samples that are difficult to sequence are those that simply have limiting DNA quantities available can be quickly and reliably analyzed using single source reagents and controls and fully integrated bioinformatics software.

Generally individuals new to this method will struggle because of the complexity of targeted NGS, which this method simplifies. However, the procedures for library quantification and sample pooling require special attention to achieve uniform coverage across all sample libraries. This protocol integrates wet and dry bench processes for the sequencing of challenging cancer specimens.

The first step is quantification and qualification of DNA to determine the number of DNA template copies that can be amplified by real time PCR. Begin the procedure for sample quantification and quality control by preparing a sufficient amount of a master mix in a microcentrifuge tube. Use the following volumes per sample, 5 microliters of 2x master mix, 0.5 microliters of primer probe mix, 0.5 microliters of inhibition primer probe mix, 0.05 microliters of ROX, and 2.95 microliters of diluent.

Add nine microliters of the master mix into each well of a 96 well plate. Add one microliter of the DNA standards in duplicate and mix by pipetting up and down five times. After ensuring that the nucleic acid sample is well mixed, add one microliter of the sample to the master mix and mix by pipetting up and down five times.

Place the sealed plate into the PCR system. Perform PCR cycles of 10 minutes at 95 degrees Celsius followed by 40 cycles of 15 seconds at 95 degrees Celsius and one minute at 60 degrees Celsius. Analyze the qPCR data by generating a linear regression plot for each of the duplicate DNA standards.

The concentration of the unknown DNA in functional or amplifiable copy number per microliter is then calculated as described in the protocol text. The next step after sample quantification and quality control, is the enrichment of cancer hot spot sequences by single tube multiplex PCR. Gene specific or gsPCR, will be performed to enrich for 46 loci in 21 cancer genes.

Prepare a sufficient amount of gsPCR master mix in a microcentrifuge tube by using the following volumes per sample, five microliters of 2x amplification master mix, and one microliter of pan-cancer primer panel. Aliquot six microliters of the gsPCR master mix into each well of a 96 well plate. Add four microliters of each nucleic acid sample into individual wells.

Mix by pipetting up and down five times. Include the appropriate controls. Place the sealed plate in the thermocycler for the following PCR cycles.

Fives minutes at 95 degrees Celsius. 15 seconds at 95 degrees Celsius followed by four minutes at 60 degrees Celsius for two cycles. 15 seconds at 95 degrees Celsius followed by four minutes at 72 degrees Celsius for 23 cycles.

A final extension of 10 minutes at 72 degrees Celsius, and followed by holding at four degrees Celsius. After gene specific PCR, perform 10 cycles of tag-PCR to incorporate platform specific adapters for compatibility with next generation sequencing. Add 7.5 microliters of the 2x index master mix and 5.5 microliters of an index code to a specified well in a 96 well plate and mix by pipetting up and down five times.

Carefully open the gsPCR plate and add two microliters of gsPCR product to the new plate with the master mix. Place the sealed plate in the thermocycler for the following PCR cycles, five minutes at 95 degrees Celsius, 30 seconds at 95 degrees Celsius, 30 seconds at 55 degrees Celsius, and one minute at 72 degrees Celsius for 10 cycles, and a final extension of 10 minutes at 72 degrees Celsius. Followed by holding at four degrees Celsius.

Following tag-PCR, library purification and size selection is accomplished using magnetic bead chemistry. Begin this procedure by adding 11 microliters of library pure prep magnetic beads into separate wells of a 96 well plate. Open the tag-PCR plate and add 10 microliters of tag-PCR product to the beads and mix by pipetting up and down five times.

Incubate the mixture for four minutes at room temperature. Place the 96 well plate on a magnetic stand for four minutes. With the 96 well plate still on the stand, remove and discard the supernatant with a pipette.

After washing the beads twice with ethanol containing wash buffer as described in the protocol text dry the beads for two minutes at room temperature and then remove the plate from the stand. Resuspend the beads by adding 20 microliters of elution buffer to each well and pipetting up and down five times. Incubate for two minutes at room temperature.

Place the 96 well plate back on the magnetic stand for four minutes and carefully remove and transfer 18 microliters of the clear supernatant to a well in a new 96 well plate. The next step in this protocol is the quantification of each purified sample library by relative competitive real time PCR. To begin this procedure, prepare a sufficient amount of the LQ master mix in a microcentrifuge tube using the following volumes per sample, five microliters of 2x LQ master mix, two microliters of LQ primer probe mix, 0.5 microliters of LQ standard, and 0.5 microliters of LQ ROX.

Add 8 microliters of the LQ master mix to a well of an optical 96 well plate. In separate wells, add two microliters of a serial dilution of the library, two microliters of LQ positive control, and two microliters of LQ diluent and mix by pipetting up and down five times. Place the sealed plate into the PCR system and assign both FAM and VIC detectors for each sample.

Perform PCR amplification using the following cycling conditions, five minutes at 95 degrees Celsius, and 15 seconds at 95 Celsius, followed by one minute at 60 degrees Celsius for 40 cycles. The concentration of each sample is then calculated as described in the protocol text. Following library quantification, each sample library is normalized based on it’s measured concentration and pooled into a single tube for sequencing.

A simplified analysis module is used to facilitate the computations for library normalization and sample pooling. To normalize the library, first determine the median concentration in nanomolar across all samples each containing a unique pair-wise index to be pooled. Next, determine the individual sample volume in microliters to pool by multiplying the median concentration across all samples by five, then dividing by it’s individual concentration.

Add the normalized volume for each sample to a single microcentrifuge tube to create the sample pool. Dilute the sample pool to 1.25 nanomolar using sequencing diluent. Prepare the sample pool for sequencing by denaturing it in the presence of phix control V3.Combine the following, 15 microliters of 1.25 nanomolar sample pool, three micorliters of 0.5 nanomolar phix, and two microliters of 1N sodium hydroxide.

After a brief vortex and centrifugation, incubate for five minutes at room temperature. Add 8 microliters of the denatured library to 992 microliters of prechilled HT1 high buffer in a microcentrifuge tube. Add 600 microliters of the denatured and diluted library to position number 17 of the reagent cartridge.

In microcentrifuge tubes, separately dilute four microliters of each sequencing primer with 636 microliters of HT1 high buffer. Add 600 microliters of diluted read-1 sequencing primers to position number 18 of the sequencing reagent cartridge. 600 microliters of diluted index read primers to position number 19 and 600 microliters of diluted read-2 sequencing primers to position number 20.

Load the reagents on the next generation sequencing instrument and perform a paired end, two by 150 cycle sequencing run according to the manufacturers instructions. Finally analyze the sequencing data using the companion bioinformatics software. A total of 90 samples could be processed in a single NGS run using a bench top sequencer.

For the data set shown, samples included cell line, residual clinical formal and fixed parafin embedded, or FFPE, and synthetic template DNA and resulted in high depth sequencing and uniform coverage. Outliers were comprised of no template controls, a dilution series of one cell line DNA with a large copy number amplification, and one FFPE DNA that was flagged for PCR inhibition by the preanalytical qPCR based QC assay. Coverage uniformity across the 46 amplicons was maintained using three different operators and for different low quality FFPE DNA samples.

In FFPE tumor DNA control mixture comprised on a known 5%BRAF V600E mutation was reported to have the target BRAF mutation at an average abundance of 5.2 plus or minus 1.3 percent by three operators using an input of 400 amplifiable copies. Dilutions of cell line and FFPE DNA samples with copy number amplifications demonstrated does dependence for amplification of EGFR and KRAS. Imporantly, FFPE DNA input could be reduced to as few as 50 amplifiable copies or 1.2 nanograms of bulk DNA.

While preserving the detection of known mutations without any false positive calls. Once mastered, this technique can be done in 11 hours with about three and half hours of hands on time for a batch size of 24 samples at a time, if it is performed properly.