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In This Article

  • Summary
  • Abstract
  • Introduction
  • Protocol
  • Results
  • Discussion
  • Disclosures
  • Acknowledgements
  • Materials
  • References
  • Reprints and Permissions

Summary

Here we describe a protocol that uses droplet polymerase chain reaction for target enrichment followed by next generation sequencing of blood plasma circulating tumor DNA. This technique was used to characterize mutations in the genes ESR1 (all coding regions), TP53 (all coding regions), PIK3CA (hotspots), PIK3R1 (hotspots) and POLE (exonuclease domain).

Abstract

The estrogen receptor gene (ESR1) is expressed in approximately two-thirds of breast cancers (BCs) and predicts sensitivity to endocrine therapy. Mutations in ESR1 have recently been associated with endocrine therapy resistance in patients with estrogen receptor positive metastatic breast cancer (ER+ MBC). Thus, monitoring the status of ESR1 mutations may facilitate personalized therapy decisions for ER+ MBC patients. Additionally, mutations in PIK3CA and TP53 are also prevalent in ER+ MBC and may influence therapeutic responses. Recent studies demonstrate mutational heterogeneity in metastatic breast cancer (MBC), highlighting a need to monitor for the emergence of new mutations over time. The analysis of blood plasma circulating tumor DNA (ctDNA) by next generation sequencing (NGS) has emerged as an attractive approach to address the mutation heterogeneity and evolution of MBC over time. However, the high costs and intensive bioinformatics required for plasma ctDNA NGS analysis limit its utility in clinical studies that require longitudinal monitoring. We have recently developed and validated an assay for plasma ctDNA mutation profiling that utilizes droplet PCR-based multiplexed target enrichment followed by NGS, which we have termed dPCR-Seq. Here we describe the protocol for dPCR-Seq, illustrating its relative simplicity in library preparation and bioinformatics analysis to detect ESR1 (all coding regions), TP53 (all coding regions), PIK3CA (hotspots), PIK3R1 (hotspots), and POLE (exonuclease domain) mutations in breast cancer patients. We have validated a subset of the ESR1 mutations identified by dPCR-Seq using allele-specific digital PCR (dPCR) assays, demonstrating exceptional concordance in the measurement of mutant allele frequency (MAF) in clinical plasma ctDNA specimens. We anticipate that dPCR-Seq may have practical utility in future studies that investigate longitudinal monitoring of plasma ctDNA mutations as potential biomarkers of therapeutic response in ER+ MBC patients.

Introduction

Breast cancer is the most common cancer diagnosed in women worldwide1. Endocrine therapy, which inhibits estrogen receptor signaling with tamoxifen, fulvestrant, or aromatase inhibitors (AIs), is a mainstay of treatment for ER+ MBC2,3,4,5. However, most metastatic breast cancer patients will develop resistance to endocrine therapy. Ligand-binding domain mutations in ESR1 have been identified as a key mechanism for acquired endocrine therapy resistance6,7. Mutations in four other genes (i.e., PIK3CA6,8,9, PIK3R110,11,12, POLE13 and TP5314,15,16,17,18,19,20) are also prevalent in breast cancer patients. The ability to noninvasively profile and monitor tumor-specific mutations in plasma circulating tumor DNA (ctDNA) has immense potential to guide systemic therapy decisions that are personalized and adaptive to an evolving landscape of gene mutations in metastatic cancer patients21,22. Indeed, recent studies suggest that monitoring ctDNA mutation dynamics in ER+ MBC patients may be predictive of therapeutic sensitivity23,24,25,26,27,28. However, the optimal strategy for analyzing plasma ctDNA mutations over time in clinical plasma samples has not been established.

A cost-effective multiplexed droplet digital polymerase chain reaction (dPCR) assay based on hydrolysis probes was found to be a quick and highly sensitive method to detect ESR1 hotspot mutations in ctDNA29,30,31. Although dPCR-based detection has excellent sensitivity and specificity, the assay is limited to detecting only the specific variant(s) for which the probes are designed. In contrast, plasma ctDNA next generation sequencing (NGS)-based assays have the potential to identify a broader genomic target region. However, concerns have arisen regarding their accuracy when measuring mutant allele fractions (MAF) of target alleles in plasma ctDNA32. Consequently, unique molecular identifiers were added to improve NGS accuracy when determining the absolute number of particular molecular species and their relative abundance during sequencing33.

The dPCR technology34,35 provides a platform for unified dPCR and NGS-based analyses of plasma ctDNA. We recently reported amplification of the complete coding sequences of ESR1 and TP53 and hotspot regions in PIK3CA, PIK3R1, and POLE from highly fragmented plasma ctDNA using a customized dPCR-based target enrichment assay. This was followed by NGS and bioinformatic analysis to accurately identify plasma ctDNA mutations with high sensitivity (79%) and specificity (100%) in a cohort of 58 breast cancer patients36. We estimate that the lower limit of detection for dPCR-Seq is 1.6% MAF. Significantly, we observed excellent concordance between MAF measured by dPCR-Seq versus allele-specific dPCR assays (R2+ = 0.96). This indicates that dPCR-Seq accurately maintains MAF during library preparation and subsequent NGS analyses. Our analyses indicate that dPCR-Seq is more cost-effective, simpler, and bioinformatic analyses are easier relative to alternative plasma ctDNA NGS assays, while also maintaining a favorable analytical performance profile. Furthermore, dPCR-Seq should be adaptable to alternative platforms for dPCR analyses that may comparably provide accurate multiplexed target amplification for NGS applications. Thus, we anticipate that dPCR-Seq will be useful in future studies of plasma ctDNA biomarkers of therapeutic sensitivity. Below, the dPCR-Seq protocol is described in detail.

Protocol

The collection and analysis of blood samples follows the ethics guidelines of the University of North Carolina at Chapel Hill. Informed written consent was obtained from each patient, and our study was approved by the Institutional Review Board of University of North Carolina at Chapel Hill.

1. Blood collection from breast cancer patients

  1. Collect 10 mL of blood from breast cancer patients in a glass blood collection tube. Please refer to Kumar et al. for patient characteristics36.
  2. After the blood collection, mix 8–10x by gentle inversion. Do not freeze specimens in the collection tube.
    NOTE: Process the collected blood ideally on the same day within 1–4 h of collection. However, if immediate availability of the sample is not possible, the cell-free DNA (cfDNA) is stable for up to 14 days if stored at 6 °C–37 °C.

2. Plasma extraction from peripheral blood

  1. Centrifuge the blood samples 2x, first at 1,800 x g followed by 2,000 x g, for 10 min each time and collect the plasma. Store the plasma at -80 ËšC until further use.
    NOTE: Care should be taken to avoid collecting the middle whitish layer below the plasma. Discard the blood by treating it with 10% bleach for 30 min or autoclave. Alternatively, follow any other institution or company-approved protocol to process medical waste.

3. Cell-free DNA extraction from peripheral blood

  1. Thaw the stored plasma samples in a 37 ËšC water bath for about 5 min.
  2. Use a commercial kit (see Table of Materials) to extract the cfDNA from the plasma using the manufacturer’s protocol with one modification at the elution step: Wait for about 15–60 min at room temperature (RT) after adding the elution buffer and then elute the sample. Process the sample using a vacuum manifold and elute the sample in a microfuge at 14,000 x g for 1 min.
    NOTE: A standard laboratory vacuum can be used for this step. If more concentrated cfDNA is required, then elute the sample in a small volume or concentrate it using a speed vacuum.
  3. Use 1–2 µL of cfDNA to quantify using a fluorimeter and store the eluted cfDNA at -20 ˚C until further use.

4. Designing the NGS cancer panel

  1. Design 272 primers to get 136 amplicons of 96 bp regions for targeted enrichment of ESR1 (all coding regions), TP53 (all coding regions), PIK3CA (hotspots), PIK3R1 (hotspots).
    NOTE: A complete list of the primers used in this assay is given in Appendix 1. Use any appropriate program to design the primers37,38,39.
  2. Synthesize each of the 272 individual primers as a 25 nM DNA oligonucleotide with standard desalted formulation. Mix sense and antisense oligonucleotides for each amplicon together to a final concentration of 100 µM in 60 µL of 10 mM Tris pH 8.0. Do this in a 96 well plate.
  3. Divide the oligonucleotides into two sets for targeted amplification of the 68 genomic regions in each set. So, 68 wells in both 96 well plates will have 60 µL of sense and antisense oligos at a concentration of 100 µM.
    NOTE: Both 96 well plates at this step will have 68 forward and 68 reverse primers.
  4. Treat both plates separately to make two sets of oligonucleotide mixtures. Take out 1.6 µL from each well on the plate and mix the oligonucleotides to prepare a mixture consisting of 0.16 nM of each primer (0.8 µM) in 200 µL of 10 mM Tris pH 8.0.
    NOTE: This step will lead to two tubes of oligonucleotide mixtures. Each tube will have 68 forward and 68 reverse primers. The primer sets for adjacent amplicons need to be separated into two distinct reactions to avoid template competition during dPCR.

5. Droplet generation and target enrichment by first round of PCR

  1. Thaw all reagents: genotyping master mix (2x), set 1 and set 2 primers, and ctDNA samples. Vortex all reagents for 10–15 s and quickly spin to collect the contents. Keep the reagents on ice.
  2. Prepare the master mix in a microcentrifuge tube using the following volumes per sample: 20 µL of 2x genotyping master mix, 1.6 µL of 25x droplet stabilizer, 2 µL of set 1 or set 2 primers.
    NOTE: Make 10% more volume of the amount of master mix required to avoid shortage due to potential pipetting errors.
  3. Add 5.0 ng of ctDNA and adjust the volume to 40 µL with water.
    NOTE: Because the amount of cfDNA is very limited, the protocol was standardized to use the minimum amount of cfDNA.
  4. Load the 40 µL reaction mixture on each well of the chip (e.g., RainDrop Source chip) for droplet generation.
    NOTE: Eight samples can be processed at one time using one chip. If more samples are required, then set up the reaction multiple times in a set of eight samples each time.
  5. Transfer the droplet emulsions from the chip to PCR tubes using a multichannel pipette.
  6. Set up a PCR reaction for the droplet emulsions with the set 1 and set 2 primers for each sample with the following conditions: 94 °C for 2 min; 55 cycles of 94 °C for 30 s, 54 °C for 30 s, 68 °C for 1 min; and finally one incubation at 68 °C for 10 min. Set a 1 °C/s temperature ramp speed between each step.

6. Recovery of DNA from the droplets after PCR amplification

  1. To break the emulsion, add 50 µL of droplet destabilizer to the post-amplification droplets and vortex them for 30 s. Centrifuge the preparation at 2,000 x g for 2 min to separate the aqueous and oil phase.
  2. Remove the oil phase from the bottom of the PCR tubes. To do so, insert the pipette tips carefully through the upper aqueous phase and carefully remove the bottom oil.
    NOTE: Once the oil is removed, the samples can be stored at 20 °C for up to 7 days. A total of 40 μL of sample yields approximately 33 μL of aqueous volume.
  3. Resuspend solid phase reversible immobilization (SPRI) magnetic beads by inversion. Add 39.6 μL of SPRI magnetic beads into each of the aqueous volumes to maintain 1.2x bead to reaction volume ratio. Mix the beads plus the aqueous sample by pipetting up and down about 10x so that the slurry appears uniform in density.
  4. Leave the PCR tubes at RT for 5 min. Firmly position the PCR strips with samples onto a 96 well magnetic plate. Leave it for 2–3 min or until pellets are formed and the supernatant is clear.
  5. Remove the supernatant using a multichannel pipette.
    NOTE: Avoid disturbing the pellet at this step. If a bead is also aspirated into the pipet tip, then redeposit it into the well and wait for about 30 s or until the bead pellet is re-formed and the supernatant is clear. Repipette the supernatant.
  6. Leave the PCR tubes on the magnet and add 180 μL of freshly made 85% ethanol to each tube. Mix by pipetting 5–6x. Leave for 1 min or until the supernatant is clear. Remove all the ethanol carefully.
    NOTE: The ethanol should be completely removed at this step. If any ethanol is visible, place the tubes on the magnet once again, wait for 15 s, and remove the remaining ethanol.
  7. Remove the PCR tubes from the magnet and leave at RT for 5 min maximum to dry the bead pellets. Add 20 μL of 10 mM Tris-HCl pH 8.0 per well with a multichannel pipette. Vigorously pipette 10x to resuspend the pellets.
  8. Place the tubes at RT for 2 min and then place on the magnet for 2 min to separate the beads. Use a multichannel pipette to collect 17 μL of eluent from each tube and deposit in a PCR tube.

7. Addition of adaptor and index sequences at the second round of PCR

  1. Thaw all reagents: 10x buffer, 50 mM MgSO4, dNTP (10 mM each), 4 M betaine, 5 µM universal forward primer, 5 µM index reverse primer and the first PCR template DNA from steps 5 and 6. Vortex all reagents for 10–15 s and quick spin to collect contents. Keep the reagents on ice.
  2. Prepare the master mix in a microcentrifuge tube using the following volume per sample: 3.25 µL of 10x buffer, 0.875 µL of 50 mM MgSO4, 1.124 µL of dNTP (10 mM each), 2.5 µL of 4 M betaine, 1.25 µL of DMSO, 1.25 µL of 5 µM universal forward primer, and 0.5 µL of high fidelity Taq polymerase.
    NOTE: Make 10% more volume of the amount of master mix required to avoid shortage due to potential pipetting errors.
  3. Add 1.25 µL of 5 µM index reverse primer and 13 µL of the first PCR template DNA to make 25 µL of secondary PCR reactions.
  4. Set up a PCR reaction with the set 1 and set 2 primers for each sample with the following conditions: 94 °C for 2 min; 10 cycles of 94 °C for 30 s, 56 °C for 30 s, 68 °C for 1 min; and finally one incubation at 68 °C for 10 min. Set a 1 °C/s temperature ramp speed between each step.

8. Recovery of DNA after the second round of PCR

  1. Resuspend the SPRI magnetic beads by inversion. Add 22.5 μL of SPRI magnetic beads into each of the aqueous volumes to maintain a 0.9x bead : reaction volume ratio for the second PCR.
  2. Mix the beads plus the aqueous sample by pipetting up and down about 10x so that the slurry appears uniform in density.
  3. Leave the PCR tubes at RT for 5 min.
  4. Firmly position the PCR strips with samples onto a 96 well magnetic plate. Leave it for 2–3 min or until pellets are formed and the supernatant is clear.
  5. Remove supernatant using a multichannel pipette.
    NOTE: Avoid disturbing the pellets at this step. If a bead is also aspirated into the pipet tip, then redeposit it into the well and wait for about 30 s or until the bead pellet is re-formed and the supernatant is clear. Repipette the supernatant.
  6. Leave the PCR tubes on the magnet and add 180 μL of freshly made 85% ethanol to each tube. Mix by pipetting 5–6x.
  7. Leave for 1 min or until the supernatant is clear.
  8. Remove all the ethanol carefully.
    NOTE: The ethanol should be completely removed at this step. If any ethanol is visible, place the tubes on the magnet once again, wait for 15 s, and remove the remaining ethanol.
  9. Remove the PCR tubes from the magnet and leave at RT for 5 min maximum to dry the bead pellets.
  10. Add 20 μL of 10 mM Tris-HCl pH 8.0 per well with a multichannel pipette. Vigorously pipette 10x to resuspend the pellets.
  11. Place the tubes at RT for 2 min and then place on the magnet for 2 min to separate the beads.
  12. Use a multichannel pipette to collect 17 μL of eluent from each tube and deposit in fresh PCR tubes.

9. DNA quantification on the bioanalyzer and pooling the libraries

  1. Check the quantity and quality of the libraries using automated electrophoresis in accordance with the manufacturer’s instructions.
  2. Run 1.0 μL of each library on the automated electrophoresis instrument.
  3. Add up and record all product yields with expected target amplicons between 280–320 bp along with other off target non-specific amplicons (e.g., <250 bp and >300 bp) as a function of molarity (nM/L) for each sample.
    NOTE: Figure 1 provides more details about selecting appropriate regions for DNA quantification.
  4. Dilute each library to 2 nM/L (2 nM) using nuclease-free water as a diluent. Mix an equal volume of each library in a tube to make an aggregate pooled sample concentration of 2 nM/L.
    NOTE: The pooled library can be stored at 4 °C for up to 3 days. Pool the DNA libraries prepared from the eight samples for each MiSeq run.
  5. Quantify the pooled library on a fluorometer and automated electrophoresis instrument with targeted sequencing.

10. Targeted sequencing protocol

  1. Sequence each pooled library using custom sequencing primers (Appendix 1) and the sequencing reagent kit (see Table of Materials) on a Next Generation Sequencer following the manufacturer’s instructions for 125 cycle paired-end sequencing.
  2. Trim the reads in the FASTQ files to remove the adaptors and any low-quality bases at the ends using the ea-utils module fastq-mcf. Use the default parameters except for k = 2.
  3. Align the sequence against the human reference genome [hg38] using Bowtie2 (bowtie2–2.2.4). Use default parameters except --local -N 1 -p 5).
  4. Select the two groups of aligned reads.
    NOTE: Filter the reads to be ≥60 nucleotides in length and select two groups of aligned reads. One group has both reads mapped to the same amplicon on the corresponding strands allowing a 1-nucleotide mismatch in the PCR primer region. The other group has both reads mapped to different amplicons within 1 kb of each other with primer sequences matching the amplicons to which they mapped allowing a 1-nucleotide mismatch.
  5. Using Samtools, create BAM files containing each of the two categories of reads in which the 5′- and 3′-primer sequences were soft-clipped, and the alignment positions were adjusted. Merge, sort, and convert the BAM files to mpileup files (samtools-1.19; mpileup with parameters -A -B -d 1000000 -Q 30 -q 20).
  6. Call variants using VarScan2 (VarScan.v2.3.5.jar mpileup2snp and mpileup2indel, --min-coverage 100 --min-reads2 1 --min-avg-qual 30 --min-var-freq 0 --strand-filter 1 --p-value 0.01 --output-vcf 1). Annotate the variants by snpEff (default parameters). Annotated variants unique to the patient samples are scored as true mutations.

Results

Detection of mutations ESR1, PIK3CA, TP53, PIK3RA, and POLE mutations
The dPCR-SEQ assay was used to detect mutations in 31 metastatic breast cancer patients (single time point samples from 24 patients, two time point samples from six patients and three time point samples from one patient). Mutations found in ESR1, PIK3CA, TP53, POLE, and PIK3RA genes by dPCR-Seq is shown in Figure 2

Discussion

Digital PCR has become an important tool in translational research that is used widely to track hotspot mutations in cancer patients. Genome-wide and targeted NGS of liquid and solid biopsy samples has also successfully been used to identify mutations in breast cancer patients36. We have developed a five gene panel to track mutations in ESR1, PIK3CA, PIK3R1, POLE, and TP53 mutations in plasma ctDNA isolated from metastatic breast cancer patients. Altern...

Disclosures

The authors declare no conflict of interest.

Acknowledgements

The authors thank RainDance Technologies for assistance in designing primers for target enrichment of ESR1, TP53, PIK3CA, PIK3R1, and POLE. We also thank M. Consugar and S. Guharaj (Raindance Technologies) for assistance with the Raindance Thunderbolts OpenSource platform. Authors thank all the funding agencies. The study was supported by the University Cancer Research Fund at the University of North Carolina at Chapel Hill and UNC Breast Cancer SPORE grant CA058223. G.P.G. holds a Career Award for Medical Scientists from the Burroughs Wellcome Fund.

Materials

NameCompanyCatalog NumberComments
10 μL Aerosol Barrier tipsVWR10017-062Can be replaced with other equivalent product
10 mL Serological PipettesVWR13-675-20Can be replaced with other equivalent product
10-100 μL 8-channel pipetteEppendorf3125000036Can be replaced with other equivalent product
1250 μL Aerosol Barrier tipsVWR10017-092Can be replaced with other equivalent product
2 μL Aerosol Barrier tipsVWR10010-364Can be replaced with other equivalent product
20 μL Aerosol Barrier tipsVWR10017-064Can be replaced with other equivalent product
200 μL Aerosol Barrier tipsVWR10017-068Can be replaced with other equivalent product
20-200 ul 8-channel pipetteDenville Scientific463230067Can be replaced with other equivalent product
2100 BioanalyzerAgilentG2939BA
5 mL Serological PipettesVWR13-675-22Can be replaced with other equivalent product
8-strip 0.2 mL PCR tubes & capsAxygenPCR-0208-CP-C
Absolute EthanolSigmaE7023-500ML
Agilent DNA 1000 KitAgilent5067-1504
Betaine AnhydrousSigmaB2629-100G
Bio-Rad’s C1000 Touch Thermal Cycler with 96–Deep Well Reaction ModuleBio-Rad1851197
Cell-Free DNA BCT tubes, RUOStreck218962
Centrifuge 5810REppendorf22625101Suitable for 15 ml conical tubes; Can be replaced with other equivalent instrument
DMSOSigmaD8418-100ML
dNTP Solution MixNew England BiolabsN0447L
DynaMag-96 magnetic plateLife Technologies12331D
Falcon 15ml Conical Centrifuge TubesCorning352096Can be replaced with other equivalent product
Falcon 50mL Conical Centrifuge TubesCorning352098Can be replaced with other equivalent product
Heating block (for 1.7 ml microcentrifuge tubes )Denville ScientificI0540
MicrocentrifugeEppendorf5424Suitable for 1.5-2.0 ml tubes; Can be replaced with other equivalent instrument
MiSeq Reagent Kit v3IlluminaMS-102-3003
MiSeq SequencerIlluminaSY-410-1003
Nuclease Free WaterIntegrated DNA Technologies11-05-01-04Can be replaced with other equivalent product
P10 pipettesDenville Scientific355022105Can be replaced with other equivalent product
P1000 pipettesDenville Scientific455060205Can be replaced with other equivalent product
P2 pipettesDenville Scientific455010336Can be replaced with other equivalent product
P20 pipettesDenville Scientific355032002Can be replaced with other equivalent product
P200 pipettesDenville Scientific45505009Can be replaced with other equivalent product
Platinum Taq DNA Polymerase High FidelityLife Technologies11304-029
Portable Pipet-Aid XP Pipette ControllerDrummond Scientific4-000-101
QIAamp Circulating Nucleic Acid KitQiagen55114
Qubit fluorimeterThermo Fisher ScientificQ33226
Qubit Assay TubesThermo Fisher ScientificQ32856
Qubit dsDNA HS Assay KitThermo Fisher ScientificQ32851
RainDrop System Source InstrumentRaindance20-04401
SPRIselect Reagent KitBeckman CoulterB23318
TaqMan Genotyping Master MixLife Technologies4371355
ThunderBolts Cancer Panel Consumables PackRaindance20-07205
Tris BaseThermo Fisher ScientificBP152-500
Vortex mixerDenville ScientificVortexer 59ACan be replaced with other equivalent product

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