Spotlight 59 Oncology Panel

Targeted NGS Amplicon Library Prep for Illumina Platform

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INTENDED USE

The Spotlight 59T Panel for Illumina platforms enables the preparation of high quality targeted Next Generation Sequencing (NGS) libraries from a variety of liquid biopsy sample types, including CTCs (circulating tumor cells) enriched from peripheral blood, and circulating, and cell-free DNA (cDNA). For formalin-fixed, paraffin-embedded tissue (FFPE) samples, the sensitivity/specificity will be lower due to DNA damage, and computational workflow changes are needed for best performance. Please contact Fluxion for more information. Adapters are included for dual indexing and multiplexing up to 48 samples (96 reactions) on a sequencing run.

The kit utilizes Illumina-compatible adapter sequences and has been validated on Illumina platforms only. The table below lists key characteristics and typical performance.

Spotlight 59™M protocol is optimized for 10-25ng of genomic or cDNA per reaction (i.e. a minimum total of 20ng input DNA per sample is required) Quantifying the starting genomic material is highly recommended.

Spotlight 59(TM) contains 277 amplicons with an average size of 140bp that cover hotspots and contiguous regions of 59 genes, and an additional 104 amplicons with an average size of 145bp that cover exonic SNPs with high minor allele frequency and gender identification targets. These Sample_ID primers have been manufactured as spiked-in to the panel primer pool at a low percentage to account for only 2-4% of total reads. A sequencing depth of minimum 7,500X for the main panel targets is recommended.

INTRODUCTION

This protocol explains how to prepare and sequence DNA libraries using the
Spotlight 59 panel.

Before you start

  • Upon receipt, store the kit at -20 °C.
  • Separate pre-PCR reagent box (Box 1 of 2) from post-PCR reagents (Box 2 of 2) to prevent contamination
  • Store PEG solution at room temperature. (PEG solution may freeze during
    shipping, but testing showed that freezing and thawing cycle does not
    negatively impact quality of the reagent. If immediately used upon arrival,
    allow it to reach room temperature before use)

Kit Contents

Kit contains enough reagents for the preparation of either 24 samples or 48 samples, depending on kit size (10% excess volume provided).

Required Materials Not Supplied

  • SPRIselect beads (Beckman Coulter, Cat. No. B23317/B23318/B23319) or AMpure XP bead(Beckman Coulter, Cat. NO A63881)
  • Invitrogen DynaMag™, Agencourt® SPRIPlate® or similar magnetic rack for magnetic bead clean-ups
  • High quality input DNA (see cDNA Extraction and Quantification Guide)
  • PCR-based library quantification assay for Illumina libraries (NEB™M,
  • E7630S/L or Kapa Biosystems, KK4824)
  • Microcentrifuge
  • Programmable thermocycler operating within manufacturer's specifications
  • 0.2mL PCR tubes or 96-well plate
  • Aerosol-resistant tips and pipette ranges from 1-1000uL
  • 200-proof/absolute ethanol (molecular biology grade)
  • Nuclease-free water (molecular biology grade)
  • Sequencing kit for validated instruments (MiSeq, HiSeq or MiniSeq); Note: read lengths must be ≥150bp per read, so ≥300 cycle kits need to be used.

Spotlight 59 amplicons, like any amplicon enrichment technology, poses a risk of contamination of surfaces and other samples following the amplification step. Please use extreme caution when opening your sample tubes following the Multiplex PCR step. It is highly recommended that separate workspaces and pipettes be maintained for pre PCR and post-PCR steps. A negative pressure hood should be used for post-PCR steps if available. Clean lab areas using 0.5% sodium hypochlorite (10% bleach) and use specialty barrier pipette tips. Dispose of pipette tips and other disposables in sealed plastic bags.

PROTOCOL

Introduction

This section describes the Spotlight 59™M Library Prep protocol. Follow the protocol in the order described using the specified parameters. Before proceeding, verify kit contents and make sure that you have the required equipment and consumables. See Required Materials Not Supplied

Prepare for Pooling

If you plan to pool libraries, record information about your samples before beginning library prep

Tips and Techniques

Avoiding Cross-Contamination

  • When adding or transferring samples, change tips between each sample.
  • When adding adapters or primers, change tips between each sample.
  • Remove unused index adapter tubes from the working area.

Handling Beads

  • Pipette bead suspension slowly.
  • When mixing, mix thoroughly.
  • If beads are aspirated into the pipette tips, dispense back to the plate on the magnetic stand and wait until the liquid is clear.
  • When washing beads: 
    • Use the appropriate magnet for the plate.
    • Dispense liquid so that beads on the side of the wells are wetted.
    • Keep the plate on the magnet until the instructions specify to remove it.
    • Do not agitate the plate while on the magnetic stand. Do not disturb the
      bead pellet.

PROTOCOL OVERVIEW

  • This protocol contains a Multiplex PCR step for the simultaneous production of hundreds of amplicon targets in a single tube and an Indexing step for the addition of dual indexed adapters, enabling multiplexing of up to 96 unique libraries.
  • Bead-based SPRI clean-ups are used to purify the sample by removing
    unused oligonucleotides and changing buffer composition between steps.

LIBRARY PREP

For best results, please follow these instructions:

  • To maximize efficient use of enzyme reagents, remove enzyme tubes from -20 °C storage and place on ice, NOT in a cryocooler, for at least 10 minutes to allow reagents to reach 4 °C prior to pipetting. Attempting to pipette enzymes at -20 °C may result in a shortage of enzyme reagents.
  • After thawing reagents, briefly vortex all reagents except the enzymes in the Indexing Step (Y2, Y3, Y4) and spin down in a microcentrifuge.
  • Separate the Multiplex PCR Reagents (keep in pre-PCR area) and Indexing Reagents (keep in post-PCR area).
  • Prepare reactions on ice before adding to samples and performing incubations.
  • Before starting, prepare a fresh 80% ethanol solution using 200- proof/absolute ethanol and nuclease-free water (approximately 1mL will be used per sample).
  • Wait 5 minutes each time you put tubes on a magnetic stand.
  • A calculator tool to help scale up your reactions is available by visiting
    www.swiftbiosci.com on a product page.

Multiplex PCR Step

  1. Save the following program on the thermal cycler (confirm lid heating is turned ON).
  2. Load 10uL of sample DNA (adjust with Pre-PCR TE) into each PCR tube.
  3. Prepare 2 separate tubes per sample. Label and use reagent tubes A and B for the first half of the samples (either 12 or 24, depending on the kit), and tubes C and D for the second half of the samples to be processed.
  4. Assemble on ice. Components A, B, C or D (For example: Tube A will have reagent A only and not B, C or D) along with G2, and G3 should be gently vortexed first and may be master-mixed when running multiple samples in parallel.
  5. Mix well and then add 20uL of the Multiplex PCR Reaction Mix to each of the PCR tube containing 10uL DNA. Place in the thermocycler and run the program.
    Treat PCR products with care to avoid contaminating work areas and other samples. It is highly recommended that separate workspaces and pipettes be maintained for pre-PCR and post-PCR steps. A negative pressure hood should be used for post-PCR steps if available. Clean lab areas using 0.5% sodium hypochlorite (10% bleach) and use specialty barrier pipette tips. Dispose of pipette tips and other disposables in sealed plastic bags.
  6. Move samples to post-PCR area before opening tubes. Keep samples at room temperature. At no time should 'with bead' samples be stored on ice, as this affects binding to SPRI beads.
  7. Make the Indexing Reaction Mix with the following components. Assemble this reaction mix on ice and keep cold until adding it to samples in the Indexing Step, but leave the samples themselves at room temperature in preparation for SPRI cleanup. All components may be master-mixed when running multiple samples in parallel.

SPRI Step 1

Ensure beads and samples are at room temperature. Briefly vortex beads to homogenize before use.

  1. Add 36uL of SPRIselect beads to each 30uL sample (ratio: 1.2).
  2. Mix by vortexing. (Ensure no bead-sample suspension droplets are left on the sides of the tube.)
  3. Incubate at room temperature for 5 minutes.
  4. Briefly spin the samples in a microcentrifuge.
  5. Place on a magnetic stand and wait until the liquid is clear (~5 minutes).
  6. Remove and discard all supernatant from each tube (approximately 5uL may be left behind).
  7. Leave tubes on the magnet stand. Wash 2 times as follows:
    1. Add 200uL of fresh 80% ethanol to the pellet
    2. Incubate for 30 seconds
    3. Remove the ethanol solution.
  8. Briefly spin the samples in a microcentrifuge.
  9. Place back onto the magnet.
  10. Using a 20uL pipette, remove residual 80% ethanol from each well.
  11. Air-dry the pellet briefly, watching the pellet to avoid cracking or over- drying. Leave tubes on the magnet. Proceed to the Indexing Step for resuspension without delay.

Indexing Step

Continue working in the post-PCR area.

  1. Add a unique combination of 5uL Index D50X + 10uL Index D7XX to each
    sample bead pellet.
  2. Add 35uL of the cold Indexing Reaction Mix to each sample and resuspend the pellet (total volume 50uL).
  3. Place in the thermal cycler and incubate at 37 °C for 20 minutes (Lid
    heating OFF).

SPRI Step 2

Ensure PEG NaCl solution is at room temperature. Briefly vortex the PEG NaCl solution to
homogenize before use.

  1. Add 42.5uL of PEG NaCl solution to each 50uL sample (ratio: 0.85).
  2. Mix by vortexing. (Ensure no bead-sample suspension droplets are left on the sides of the tube.)
  3. Incubate at room temperature for 5 minutes.
  4. Briefly spin the samples in a microcentrifuge.
  5. Place on a magnetic stand and wait until the liquid is clear (~5 minutes).
  6. Remove and discard all supernatant from each tube (approximately 5uL may be left behind).
  7. Leave tubes on the magnet stand. Wash 2 times as follows:
    1. Add 200uL of fresh 80% EtOH to the pellet
    2. Incubate for 30 seconds
    3. Remove the ethanol solution.
  8. Briefly spin the samples in a microcentrifuge,
  9. Place back onto the magnet.
  10. Using a 20uL pipette, remove residual 80% EtOH from each well.
  11.  Air-dry the pellet briefly, watching the pellet to avoid cracking or over-drying, while on the magnet.
  12. Take tubes off the magnet.
  13. Add 20uL of Post-PCR TE buffer and re-suspend the pellet, mixing well by pipetting up and down until homogenous.
  14. Incubate at room temperature for 2 minutes off of the magnet.
  15. Place on a magnetic stand and
  16. Transfer 20uL library eluate to a fresh tube.
  17. Ensure that eluate does not contain magnetic beads. If magnetic beads are present, pipette eluate into a new tube, place on magnet, and transfer eluate again.

LIBRARY QUANTIFICATION

Quantify a 1:10,000 dilution of your library in triplicate using a qPCR assay based upon a library size of 265bp. Upon calculating library concentration, be sure to adjust for proper library size of the standards in your library quantification kit. Variation in length of DNA in the standards from the kit and your library size may lead to improper estimation of DNA concentration.

Improper library quantification by other methods will lead to uneven pooling and suboptimal cluster density, impacting sequencing data.

It is not recommended to use a Bioanalyzer for quantifying libraries because:

  • As there is no PCR enrichment of the library following the Indexing Step, the Bioanalyzer will not accurately quantify fully adapted library vs. other DNA.
  • Library adapters have secondary structure which exhibits migration artifacts on the Bioanalyzer.

It is not recommended to use a fluorometric method (such as Qubit) for quantifying
libraries because:

  • As there is no PCR enrichment of the library following the Indexing Step, a fluorometric method will not accurately quantify fully adapted library VS. other DNA.

Illumina Sequencing

The panel should be sequenced on Illumina machines in a 2x150 bp run. For MiSeq v3, we recommend using the 2x300 kit and stopping at cycle 151 on each side. For MiSeq we recommend spiking 1% phiX and loading the libraries at 12pM. For maximum resolution, we recommend sequencing each replicate to >=8000x. On MiSeq v3 this allows for multiplexing of 16-24 libraries (8-12 duplicate samples) depending on uniformity and yield, and 8-12 libraries (4-6 duplicate samples) on MiSeq v2.


APPENDIX A:

Sample quantities for Spotlight 59 panel and ERASE-Seq Variant Caller

Spotlight 59 utilizes the ERASE-Seq algorithm (https://www.ncbi.nlm.nih.gov/pmc/articles/PMC5890993/) to provide high-resolution variants calls at ultralow allele frequencies in NGS data. Part of this algorithm involves the incorporation of separate molecular amplification pools (MAPs) in order to increase the accuracy of variant calls. The protocol is optimized for realistic cDNA yields and sequencing resources, and therefore the standard protocol involves using two MAPs per sample (10ng per MAP, 20ng total input) sequenced at >= 7500x each (>=15000x total for the combined MAPs). This provides accurate detection of clinically-relevant variants to 0.1% allele frequency.

When DNA input is limited (<20ng per sample) we still recommend dividing the sample into two reactions, unless total DNA is < 10ng, at which point we recommend loading the entire sample into a single reaction to avoid dropout of low frequency alleles.

Sample Sheet - Special Considerations for MiSeq

  • Open Illumina Experiment Manager and create a sample sheet.
  • On the Instrument selection page, select "MiSeq".
  • In the MiSeq Application Selection page, select category "Other" and select application "FASTQ Only"
  • On the workflow parameter page:
  • Enter the Reagent Cartridge barcode.
  • Select "TruSeq HT" as the Sample Prep Kit.
  • Index Reads: "2".
  • Read Type: "Paired End".
  • Cycles Read 1: "151", Cycles Read 2: "151".
  • Make sure the "Use Adapter Trimming" and "Use Adapter Trimming Read 2" are selected.

Troubleshooting Common Problems

Library analysis on Agilent Bioanalyzer or TapeStation

Please note that qPCR-based methods are most accurate for quantifying Spotlight 59 Amplicon libraries. Despite that, this section provides an overview of expected results when using a Bioanalyzer. The secondary structure of Spotlight 59 Amplicon libraries exhibits two features, which should be understood if analyzed using electrophoretic methods such as Agilent Bioanalyzer or TapeStation:

  1. If using high quality DNA, "extended amplicons" can be observed. They are formed from the forward primer and the reverse primer of two adjacent amplicons. Note that these extended amplicons are not formed when using fragmented or cross-linked (FFPE) DNA, or cell-free DNA. Coverage uniformity is not affected by the presence or absence of extended amplicons.
  2. After indexing, the library is partially single-stranded and the migration is impaired, causing the library to appear large on the Bioanalyzer; therefore, the traces should not be used to accurately determine the size or the quantity of the library.

Indexed Adapter Sequences

During the Indexing Step in the protocol, you must use a unique combination of Index Adapters to re-suspend and label each library. Libraries made with uniquely indexed adapter combinations may be multiplexed during cluster generation and co-sequenced on the same Illumina flow cell.

CONTENTS: Unique indexed adapters, which should be used where this manual calls for 5 or 10uL of each Index Primer:

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