Instructions for Use: Spotlight Amplicon Panels for Oncology Research (Normalase Technology)

Applies to: 910-0133

Updated: 2023-04-07
Revision: B

TABLE OF CONTENTS

OVERVIEW

CONSUMABLES AND EQUIPMENT

GUIDELINES

PROTOCOL

APPENDIX A: SEQUENCING RECOMMENDATIONS

APPENDIX B: SEQUENCING DATA ANALYSIS

APPENDIX C: INDEXED ADAPTER SEQUENCES

APPENDIX D: PRIMER SEQUENCES

APPENDIX E: MULTIPLEX SEQUENCING RECOMMENDATIONS

APPENDIX F: PLATE USAGE GUIDELINES

APPENDIX G: GEN AMPLICON UDI PRIMER PLATE SPECIFICATIONS AND DIMENSIONS

APPENDIX H: xGEN AMPLCON UDI PRIMER PLATE LAYOUTS

APPENDIX I: cfDNA EXTRACTION AND QUANTIFICATION RECOMMENDATIONS

OVERVIEW

Designed specifically for detection of mutations from liquid biopsy samples, Spotlight Amplicon Panels are turnkey solutions for Illumina sequencers, offering ultra-sensitive variant detection from low input quantities of DNA. The high sensitivity of Spotlight panels makes them ideal for cfDNA and CTCs. Incorporating ERASE-Seq, Fluxion’s statistically powered variant caller, Spotlight delivers exceptional sensitivity in liquid biopsy somatic variant detection. These kits are based on IDT xGen amplicon technology and leverage patented multiplex PCR technology, enabling library construction from a variety of sample types. The panels are targeted for ultra-sensitive somatic variant detection from circulating, cell-free DNA (cfDNA), using tiled, overlapping primer pairs within a single multiplexed pool.

See Appendix A for panel-specific information on panel content and size as well as multiplexed sequencing recommendations. See individual panel data sheets for details regarding target design and coverage.

* Customizable kit components available. Contact us for more information at sales@fluxionbio.com
+ Panel specific

Supported applications and sample types

• Applications: Variant discovery research, oncology research
• Sample types including, but not limited to: cell-free DNA (cfDNA), circulating tumor DNA (ctDNA), genomic DNA (gDNA) from circulating tumor cell samples (CTCs)

Spotlight Amplicon workflow

Spotlight Amplicon Panels utilize multiple overlapping amplicons in a single tube, using a rapid, 2-hour workflow to prepare ready-to-sequence libraries for research studies. The kit includes the Spotlight primers, as well as xGen Core kits and xGen sample indexes. The PCR1+PCR2 workflow generates comprehensive libraries, even from low input quantities. The libraries may be quantified with conventional methods such as Bioanalyzer ® (Agilent) and normalized by manual pooling, or normalized enzymatically with the included Normalase reagents.

This protocol includes instructions for the multiplex PCR step to enrich target sequences, an indexing PCR step to amplify and add combinatorial or unique dual indexed adapter sequences, and an optional, downstream Normalase step to produce an equimolar library pool.

Figure 1. Spotlight Amplicon workflow. A dual-indexed, amplicon library is prepared from DNA in three main steps: 1) multiplex PCR; 2) adapter attachment with indexing PCR; 3) an optional Normalase treatment step to produce equimolar library pools.

CONSUMABLES AND EQUIPMENT

These kits contain sufficient reagents for the preparation of 96 libraries for 48 samples, with 2 reactions for each sample (10% excess volume provided). Each Spotlight kit includes Spotlight amplicon primers, the xGen Core Reagents, and xGen Amplicon UDI plates for sample indexing. 

Spotlight kit contents

Consumables from IDT - Kit contents

Workflow stage	Component	96 rxn (µl)	Storage
Post-PCR	xGen Amplicon UDI (pre-mixed pairs)	12 per well	-20°C

Panel Reagents

Spotlight G1 Specific Panel Primers*

xGen Core Reagents

*Customizable kit components; UDI indexing primers are ordered as a separate line item at kit purchase. Each UDI primer set contains four 96-well plates. Each plate contains 96 pre-mixed primer pairs. 

Consumable from other suppliers

Equipment

*All Amplicon Panel libraries have been created with a Bio-Rad® T100 thermal cycler in our internal testing.

GUIDELINES

Reagent handling

• Upon receipt, store the Spotlight Amplicon Library Kit products at -25 to -15°C with the exception of the PEG solution and TE, which are stored at room temperature.
• Separate the multiplex Pre-PCR Reagents (keep in pre-PCR area) from the Indexing and Normalase Reagents (keep in post-PCR area).
• To maximize use of enzyme reagents when ready to use, remove enzyme tubes from -20°C storage and place on ice for 10 minutes prior to pipetting. Attempting to pipette enzymes at -20°C may result in shortage of enzyme reagents.
• After thawing reagents on ice, briefly vortex (except enzymes) to mix well, then pulse spin to collect contents before proceeding. Enzyme G3 is the only enzyme that may be vortexed.
• Always add reagents to the master mix in the specified order, as stated throughout the Protocol. The UD indexing primers are the only reagents that are added individually to each sample.
• Assemble all reagent master mixes and reactions ON ICE and scale volumes as appropriate, using 5% excess volume to compensate for pipetting loss. Neglecting to store master mixes and reagents on ice prior to incubations reduces yields and function of this product.

DNA input considerations

For next-generation sequencing (NGS), quantity and quality of circulating cell-free DNA (cfDNA) is essential to good library preparation. We recommend quantification by a qPCR-method, using a short amplicon to accurately determine the concentration of sample DNA [Simbolo M. et al. PLoS ONE (2013) 8(6): e62692].

The starting material should be quantified with the qPCR assay to determine usable DNA content and sample integrity. The panels are designed with amplicon size of <150 bp to ensure compatibility with cfDNA samples. For qPCR-based determination of sample quantity and integrity, xGen Input DNA Quant Primers are available (IDT Catalog No. 10009856). See Appendix I, cfDNA Extraction and Quantification Recommendations for more information.

Optimal coverage uniformity is achieved with input amounts in the 10-25 ng range per reaction. Between 25-100ng, coverage uniformity may be mildly reduced. Using less than 10 ng may reduce library yields and variant calling for low frequency alleles.

Consider the following example allele frequencies and detection limits. As shown below,
using at least 20ng per reaction (40ng total per sample) input for 0.1% allele detection limit ensures sufficient copy number of the allele of interest. If less than 20ng is available, the detection limits may be reduced. 

Avoid cross-contamination

To reduce the risk of DNA and library contamination, physically separate the laboratory space, equipment, and supplies where pre-PCR and post-PCR processes are performed, including appropriate reagent boxes for pre-PCR (multiplex) and post-PCR (Indexing and Normalase) reagents. Move samples to post-PCR area before opening tubes. This workflow, like any amplicon enrichment technology, poses a risk of contamination of surfaces and other samples following the amplification step. To help minimize this concern, use caution when opening your sample tubes after the multiplex PCR step. Follow these guidelines to avoid cross-contamination:

• Clean lab areas using 0.5% sodium hypochlorite (10% bleach).
• Use barrier pipette tips to avoid exposure to potential contaminants.
• Always change pipette tips between each sample.
• Perform pre-PCR reactions in a separate location from the post-PCR area, ideally in a PCR workstation.
• Separate the multiplex Pre-PCR Reagents (keep in pre-PCR area) from the Indexing and Normalase reagents (keep in post-PCR area).

Size selection during cleanups

SPRIselect beads from Beckman Coulter (B23317/B23318/B23319) are recommended with this protocol, however, they can be substituted with Agencourt® AMPure XP beads (Beckman Coulter, Cat. Nos. A63880/A63881/A63882). Make sure that the beads and samples are at room temperature. At no time should the "with bead" samples be stored on ice, as this affects binding to magnetic beads. Briefly vortex beads to homogenize before use. Make sure that the beads and samples never completely dry during processing.

Automation

This protocol is readily automatable. A 10% overage volume of reagents is supplied to accommodate automation.

PROTOCOL

Prepare Multiplex PCR

1. Load the Multiplex PCR program and allow the block to reach 98°C before loading samples (confirm lid heating is turned ON and has reached 105°C).
   
   Preprogram thermal cycler
   Important: Temperature at this step is panel-specific. Use the panel-specific annealing and extension temperatures specified in the table below:
   

   Reagents	Lid heating ON
   Multiplex PCR thermal cycler program	30 sec	98°C
   	10 sec	98°C	4 cycles
   	5 min	* °C
   	1 min	* °C
   	10 sec	98°C	18 cycles
   	1 min	64°C
   	1 min	65°C

   Panel	Annealing temperature (5 min)	Extension temperature (1 min)
   Spotlight Myeloid Amplicon Panel	66°C	66°C
   Spotlight Custom Amplicon Panel	Per spec	Per spec

2. Gently rock Enzyme G3 at room temperature for 5 minutes, or until any solutes appear to be in solution. Place back on ice for remainder of use.

3. Prepare 2 separate tubes per sample. Use Reagent G1A for one reaction and Reagent G1B for the other.

   Each sample has 2 reactions, one with G1A and one with G1B. Equal amount of input DNA from the same sample must be used in each of the two reactions.

4. Load 10 µL of DNA sample directly into each PCR tube.
5. Keep all tubes on ice during assembly of the master mix and the reaction, until placed in thermal cycler.

Multiplex PCR Master Mix

Before mixing, calculate the total volume of the Master Mix based on the number of reactions required, with appropriate overage for pipetting. Vortex components G1A, G1B, and G2, and pulse-spin tubes to collect contents. Make the Multiplex PCR Master Mix. Keep the prepared Master Mix on ice until ready to use.

* Reagent G1A/G1B are the panel-specific set of multiplex primers.

1. Combine the PCR Master Mix well and then add 20µL of the multiplex PCR reaction mix to each 10µL input DNA sample on ice. Mix well, then place in the thermal cycler and run the Multiplex PCR thermal cycler program.
   Important: Move samples to post-PCR area before opening tubes.
2. Near the completion of the thermal cycler program, prepare the indexing reaction mix (below) in the post-PCR area. Assemble the Indexing PCR Master Mix on ice and keep cold until you add it to samples in the indexing step. All components except indexes should be master-mixed when running multiple samples in parallel.

Prepare Indexing PCR Master Mix

Before mixing, calculate the total volume of the Indexing PCR Master Mix based on the number of reactions with appropriate overage for pipetting. To maintain a volume of Enzyme I3 that can be accurately pipetted, prepare at least 10 reactions at a time. Keep the prepared master mix on ice.

Keep prepared master mix on ice during Post-multiplex PCR size selection and cleanup.

Perform post-multiplex PCR size selection and cleanup

1. Ensure beads and samples are at room temperature. Briefly vortex beads to homogenize before use.
2. Add 30µL (ratio: 1.0) of magnetic beads to each 30µL sample. Mix by vortexing. Pulse-spin the samples in a microcentrifuge. Ensure no bead-sample suspension droplets are left on the sides of the tube.
3. Incubate the samples for 5 minutes at room temperature, off the magnet.
4. Place the sample tubes on a magnetic rack until the solution clears and a pellet is formed (<5 minutes).
5. While sample is on the magnet, remove and discard the supernatant without disturbing the pellet (approximately 5µL may be left behind). Leave tubes on the magnet.
6. Carefully add 180µL of freshly prepared 80% ethanol solution to the pellet while it is still on the magnet. Do not to disturb the pellet. Incubate for 30 seconds, and then slowly remove the ethanol solution.
7. Repeat for a second wash with the ethanol solution.
8. Pulse-spin the samples in a microcentrifuge, place back onto the magnet and remove any residual ethanol solution from the bottom of the tube with a small volume tip.
9. Resuspend each bead pellet in 17.4µL Post-PCR TE Buffer. Proceed to the Indexing PCR step. If an off-bead PCR is preferred, place the tubes back on the magnet and transfer the 17.4µL of eluate to a fresh tube.
   Important: Continue working in the post-PCR area. Keep samples at room temperature. At no time should the "with bead" samples be stored on ice, as this affects binding to magnetic beads.

Perform Indexing PCR

Set up the Indexing PCR thermal cycler program as follows:

* The PCR cycle number can be increased for samples that may give low yields.

1. Load the Indexing PCR program and allow the block to reach 37°C before loading samples (confirm lid heating is turned ON and has reached 105°C).
   Add 3.7µL of a premixed xGen Amplicon UD indexing primer pair to each sample, if you are using the single use plates (See Appendix F for UDI plate usage guidelines).
2. Add 28.9µL of the cold, Indexing PCR Reaction Mix to each sample and mix thoroughly (total volume 50µL).
3. Place in the thermal cycler and run the Indexing PCR program.

Perform post-indexing PCR cleanup

Note: Make sure the PEG NaCl solution is at room temperature before starting this section.

1. Briefly vortex the PEG NaCl solution to homogenize before use.
2. Add 42.5µL (ratio: 0.85X) of PEG NaCl solution to each 50µL sample. Mix by vortexing. Make sure that there are no bead-sample suspension droplets left on the sides of the tube. If performing an "off bead" PCR, use a 42.5µL (ratio: 0.85X) of fresh magnetic beads.
3. Incubate the samples for 5 minutes at room temperature, off the magnet.
4. Pulse-spin the samples in a microcentrifuge. Place the sample tubes on a magnetic rack until the solution clears and a pellet is formed (<5 minutes).
5. While leaving your sample on the magnet, remove and discard the supernatant without disturbing the pellet (approximately 5µL may be left behind). Leave tubes on the magnet.
6. Carefully add 180µL of freshly prepared, 80% ethanol solution to the pellet while it is still on the magnet. Do not disturb the pellet. Incubate for 30 seconds, then carefully remove the ethanol solution.
7. Repeat for a second wash with the 80% ethanol solution.
8. Pulse-spin the samples in a microcentrifuge, place back onto the magnet, and remove any residual ethanol solution from the bottom of the tube with a small-volume tip.
9. Proceed immediately to add 20µL of post-PCR TE buffer and resuspend the pellet, mixing well by pipetting up and down until homogenous. Incubate at room temperature for 2 minutes off the magnet, then place the sample back on the magnet and transfer the clear library eluate to a fresh tube. Make sure that the eluate does not contain magnetic beads (indicated by brown coloration in eluate). If magnetic beads are present, place back on magnet, and transfer eluate again.
   Safe Stop: Store freshly prepared libraries at -20°C.

Perform NGS library quantification

Accurate library quantification is essential to load the sequencing instrument properly. Libraries can be quantified using fluorometric-, electrophoretic-, or qPCR-based methods and normalized manually. Alternatively, libraries can be enzymatically normalized following the Normalase protocol below.

Note: For optimal normalization using Normalase reagents, a minimum of 12nM yield is needed per sample. If library yields are below 12nM, increase the number of PCR cycles to pass the 12nM threshold, or switch to the 6nM threshold Normalase protocol described below.

Introduction to Normalase Treatment

This guide provides instructions for optional enzymatic normalization of multiplexed xGen Amplicon libraries for equimolar pooling and balanced sample representation in sequencing. The protocol is designed for Spotlight amplicon libraries that produce consistent amplified library yields of ≥12nM following indexing PCR. Most samples processed with this protocol produce amplified library yields of 12nM or greater; however, if there is concern that not all samples will reach 12nM, adjusting Normalase chemistry to require a minimum of only 6nM can alternatively be performed.

Use this simple calculator for converting between ng/µL and nM. Finish with a library size of 285 bp, or a size appropriate to the panel (for use in the Base Pair Length column).

The workflow consists of three steps for libraries amplified to a minimum yield of 12nM during the adapter attachment and indexing PCR step:

1. Normalase I to enzymatically select a 4nM (or 2nM if using ≥6nM option) library fraction
2. Equal volume library pooling of samples for multiplexed sequencing
3. Normalase II to enzymatically generate an equimolar library pool

Figure 2. Workflow schematic. Normalase | Master Mix is added to samples and incubated at 30°C for 15 minutes. Sample pooling is performed, and then the Normalase II Master Mix is added to the pool and incubated at 37°C for 15 minutes. Reagent X1 inactivates the reaction and a final, equimolar pool is produced.

Normalase specification

The Normalase product specification is defined by cluster density of the Normalase pool when loaded on a MiSeq® v2 flow cell at 12pM to achieve a 1000-1200 K/mm2 cluster density and CV ≤15% within a pool. Across Illumina platforms, library types, and insert sizes, optimization of loading concentration may be required to achieve the optimal number of reads supported by the flow cell of choice.

Perform the Normalase 1 reaction

If you are concerned that the 12nM threshold has not been met for each library after indexing PCR:

• Spot check library yields using any fluorometric method (e.g., Qubit fluorometer) or electrophoretic method (e.g., Bioanalyzer machine).
• A Normalase workflow modification can be performed that requires a 6nM threshold to obtain a 2nM Normalase pool (see below).

1. Preset a thermal cycler program as listed below.
   Thermal cycler program: 15 min at 30°C with open lid or lid heating OFF
2.  Prepare the Normalase 1 Master Mix, as listed in the table below. The mix can be prepared at room temperature and stored on ice until use if prepared in advance. Ensure that it is thoroughly mixed by moderate vortexing followed by a pulse spin to collect contents prior to use. For libraries with lower yields >6nM, or for a final pool of 2nM (instead of 4nM), use only half the specified volume of Reagent S2 with an equal volume of TE, reducing the concentration by two-fold. 
   

   Reagent	Per library (µL)	16 libraries (µL)	96 libraries (µL)
   Buffer S1	4.3	68.8	412.8
   Reagent S2	0.2	3.2	19.2
   Enzyme S3	0.5	8.0	48
   Total volume	5	80	480

   Important: The Normalase 1 Master Mix should be built for a minimum of 10 reactions to ensure pipetting accuracy. 
3. With a calibrated P10 pipette, add 5µL of Normalase 1 Master Mix into each 20µL library eluate at room temperature, then thoroughly mix by moderate vortexing for 5 seconds.
4. Spin down the sample tube in a microcentrifuge. Place in the thermal cycler and run the program described in step 1.

Perform equal volume library pooling

Sufficient Normalase II reagents are supplied so that this step can be repeated to enable various repooling combinations, as only 5µL of post-Normalase I library (out of a 25µL volume) is used for pooling. Also note that stability of normalized pools (after Normalase II) is limited, with a storage time of four weeks, since the resulting normalized pools contain single-stranded DNA. Therefore, if re-sequencing is required after four weeks, re-pool the Normalase I libraries and repeat Normalase II and inactivation.

Note: If pooling <5 libraries, see the Appendix A: Sequencing recommendations for low-plex pooling recommendations.

Note: If pooling 5µL per sample does not generate a normalized pool of sufficient volume for instrument loading, see the Appendix A: Sequencing recommendations for high sample volume pooling recommendations.

Important: Consider your desired number of reads for each sample and pool only those samples together that have the same required depth. For example, samples each requiring 50,000 reads can be pooled together, whereas samples requiring 1 million reads should be combined in a separate pool. Thus, you can adjust your ratio of pools when loading the instrument to achieve the desired sequence depth for each pool.

1. Following the Normalase I incubation, generate a library pool (or pools) by placing 5µL of each individual library into a single, 0.2mL PCR tube if pooling 30 libraries or less (achieves up to a final volume of 186µL). Alternatively, use a 1.5mL microcentrifuge tube, particularly when pooling greater than 30 libraries, as the volume will exceed the PCR tube maximum volume.
   Tip: To ensure even pooling, use a calibrated P10 pipette.
2. Thoroughly mix samples, spin the library pools in a microcentrifuge, and proceed to Perform the Normalase II reaction.

Perform the Normalase II reaction

1. Preset a thermal cycler program as shown below. Alternatively, if using a 1.5mL microcentrifuge tube, set a heat block at 37°C.
   

   Thermal cycle program	Heat block (1.5mL microcentrifuge tube)
   15 min at 37°C with open lid or lid heating OFF	15 min at 37°C

2. Pre-mix Normalase II Master Mix (listed in the table below). The master mix can be stored on ice until use and then added to pools at room temperature.
   

   Reagent	Per Library (µL)	24 libraries (µL)	96 libraries (µL)
   Buffer N1	0.96	23.04	92.16
   Enzyme N2	0.04	0.96	3.84
   Total volume	1	24	96

   * It is recommended to prepare Normalase II Master Mix for a minimum of 16 samples, even if you are processing less than 16 samples, to avoid pipetting extremely low volumes. For best results, use a calibrated, P2 pipette to add Enzyme N2. Although sufficient reagents are supplied for up to 5 repeated Normalase II reactions per sample, repeatedly processing a lower number of samples will result in significant loss of Normalase II reagents.

3. Add 1µL of Normalase II Master Mix, multiplied by the total number of libraries in each prepared pool.
4. Mix well by vortexing for 5 seconds, and spin down the library pools in a microcentrifuge.
5. Place the library pools in the thermal cycler and run the program or place the 1.5 mL microcentrifuge tubes into the 37°C heat block.

Perform Normalase inactivation

1. Following the Normalase II reaction, preset a thermal cycler program as shown below. Alternatively, if using a 1.5mL microcentrifuge tube, set a heat block at 95°C.
   

   Thermal cycler program	Heat block (1.5 mL microcentrifuge tube)
   Hold at 95°C	2 min at 95°C
   2 min at 95°C with lid kept at 95°C
   Hold at 4°C

2. Add 0.2µL of Reagent X1, multiplied by the total number of libraries in each prepared pool. See examples below:
   

   Reagent	Per Library	24-plex pool	96-plex pool
   Reagent X1	0.2µL	4.8µL	19.2µL

3. Place the library pools in the thermal cycler and advance the program, or place the 1.5mL microcentrifuge tubes into the heat block. If using a 1.5mL microcentrifuge tube, set a heat block at 95°C to incubate your library pools, being careful not to incubate the samples longer than 2 minutes.
4. Your final multiplexed library pools are now equimolar. Proceed to qPCR quantification of your Normalase pool and sequencing. It is not necessary to perform an additional purification step.

Perform calibration of Normalase pools

For better sequencing results, perform a qPCR quantification on your final Normalase pool(s). Final library pools are dsDNA and cannot be quantified by dsDNA-based fluorometric methods or electrophoretic fragment analysis. If you do not have a qPCR assay, use a commercially available kit by calibrating your qPCR results and sequencer loading concentrations for optimal clustering before proceeding (KAPA Library Quantification Kit, Cat. No. KK4828).

Calibration of Normalase output to your qPCR assay, sequencer loading procedure, and clustering output is required during testing due to variation across different qPCR assays and laboratory practices. Use a qPCR assay that predicts an optimal number of reads on your sequencing instrument, as this is important to achieve Normalase calibration. Across other Illumina platforms, library types, and insert sizes, optimization of loading concentration may be required to achieve the optimal number of reads supported by the sequencing flow cell of choice. If you have chosen the 6nM to 2nM option but require a higher pool concentration for your sequencer, perform a 2.0X SPRI cleanup to concentrate pools and then proceed to qPCR quantification and loading.

Once you have calibrated your sequencer loading procedure to the Normalase output and have established that your samples meet the minimum threshold for Normalase, qPCR of the final pool is optional but still recommended. For example, a Normalase workflow error may have occurred that would lead to unexpected results.

APPENDIX A: SEQUENCING RECOMMENDATIONS

Spotlight Amplicon Panel libraries may be sequenced using paired-end sequencing on Illumina instruments. We strongly recommend using 2 x 150 paired-end reads. The depth of coverage required will depend on the application (refer to the Overview section). Be sure to use 10 sequencing cycles for each index read.

See Appendix E for panel-specific information on panel content and size, as well as multiplexed sequencing recommendations.

Due to the complexity of the libraries, no PhiX spike-in is required on the MiSeq or MiniSeq (Illumina) instruments. The NextSeq® 550 (Illumina) may be sensitive to low complexity and PhiX or another suitable high-complexity library spike-in may be required. Contact Illumina technical support for further information regarding sequencing instrument compatibility with low-complexity sequences.

APPENDIX B: SEQUENCING DATA ANALYSIS

All Spotlight panels include variant calling using Fluxion's ERASE-Seq bioinformatics. 

Features

• Simple Windows application
• Easy-to-run workflow 

Hardware minimum requirements

• Windows 10 or 11
• Internet connection

Analysis overview

1. Generate FASTQ files for each sample. Follow naming conventions in ERASE-Seq software instructions
2. Upload files to AWS cloud server using ERASE-Seq software
3. User will receive email when reports are ready for download
4. Receive reports in VCF and Excel format

APPENDIX C: INDEXED ADAPTER SEQUENCES

The full-length adapter sequences are below, where the underlined text indicates the location of the index sequences which are 10 bp for UDI. These sequences represent the adapter sequences following completion of the indexing PCR step.

Index 1 (i7) adapters:

Index 2 (i5) adapters:

Refer to the accompanying Index Sequences Master List for index sequences for preparing your Illumina sequencing sample sheet on the instrument of your choice.

APPENDIX D: PRIMER SEQUENCES

For reference, the primer sequences are below. These primers include full-length Illumina adapter and index

sequences.

i7 primer: Replace 10 UDI X’s with the REVERSE COMPLEMENT of the specified i7 index sequence in the Index Master List:

i5 primer: Replace 10 UDI Y's with the specified Forward Strand Workflow i5 index sequence in the Index Master List: