TruSight Tumor 26 Reference Guide (15042911 v02)

TruSight Tumor 26 Reference Guide (15042911 v02)

  Chapter 2  Protocol

Protocol

Introduction

Tips and Techniques

TruSight Tumor 26 Workflow

Qualify DNA Extracted from FFPE Samples

Hybridize the Oligo Pool

Remove Unbound Oligos

Extend and Ligate Bound Oligos

Amplify Libraries

Check Library Preparation (Optional)

Clean Up Libraries

Check Libraries

Normalize Libraries

Pool Libraries

18

21

22

24

9

12

14

17

25

26

6

7

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TruSight Tumor 26 Reference Guide

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Introduction

This chapter describes the TruSight Tumor 26 protocol.

}

Review Best Practices before proceeding. See

Additional Resources on page 4

for information on how to access TruSight Tumor 26 Best Practices on the Illumina website.

} Follow the protocols in the order shown, using the specified volumes and incubation parameters.

}

As a troubleshooting aid, ACD1 (Amplicon Control DNA), and ACP1 (Amplicon

Control Oligo Pool) have been included in this kit. Using ACD1 instead of gDNA and

ACP1 instead of FPA and FPB in the TruSight Tumor 26 assay can help narrow down issues arising from gDNA sample prep or primer contamination. Libraries made with these controls cannot be sequenced alongside other TruSight Tumor 26 libraries as they require longer cycles.

Prepare for Pooling

If you plan to pool libraries, record information about your samples before beginning library prep. Different methods are available depending on the sequencing instrument you are using. See the TruSight Tumor 26 support page for more information.

Review the planning steps in the Pooling Guide when preparing libraries for Illumina sequencing systems that require balanced index combinations.

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Tips and Techniques

Unless a safe stopping point is specified in the protocol, proceed immediately to the next step.

Avoiding Cross-Contamination

} When adding or transferring samples, change tips between each sample.

} When adding adapters or primers, change tips between each row and each column.

}

Remove unused index adapter tubes from the working area.

Sealing the Plate

}

Always seal the 96-well plate before the following steps in the protocol:

}

Shaking steps

}

Vortexing steps

} Centrifuge steps

}

Thermal cycling steps

}

Apply the adhesive seal to cover the plate and seal with a rubber roller.

}

Microseal 'B' adhesive seals are effective at -40°C to 110°C, and suitable for skirted or semiskirted PCR plates. Use Microseal 'B' for shaking, centrifuging, and long-term storage.

}

Microseal 'A' adhesive film is effective for thermal cycling and easy to cut when using fewer than 96 wells.

Plate Transfers

}

When transferring volumes between plates, transfer the specified volume from each well of a plate to the corresponding well of the other plate.

Centrifugation

}

Centrifuge at any step in the procedure to consolidate liquid or beads in the bottom of the well, and to prevent sample loss.

}

To pellet beads, centrifuge at 280 × g for 1 minute.

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 (~2 minutes).

}

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.

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TruSight Tumor 26 Workflow

The following diagram illustrates the workflow using the TruSight Tumor 26 Kit. Safe stopping points are marked between steps.

Figure 1 TruSight Tumor 26 Workflow

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Qualify DNA Extracted from FFPE Samples

During this step, a qPCR reaction determines the amplification potential of FFPE-extracted gDNA samples. By comparing the amplification potential of FFPE DNA relative to that of the QCT non-FFPE reference gDNA, a ΔCq value is calculated for each sample. The ΔCq value is then used to predict sample performance in the TruSight Tumor 26 assay. The exact amount of FFPE DNA input varies according to the quality of the extracted DNA.

Consumables

} QCT (Quality Control Template)

} QCP (Quality Control Primer)

}

Genomic DNA

}

48 or 96-well plate

} Adhesive seal (dependent on qPCR machine)

} KAPA SYBR FAST qPCR Master Mix (2X) (Universal)

}

Nuclease-free water

}

PCR 8-tube strips (if using QCT for the first time)

Preparation

1 Prepare the following consumables.

Item

QCT

QCP

Genomic DNA

KAPA SYBR FAST qPCR Master Mix

(2X) (Universal)

Storage

-25°C to -15°C

-25°C to -15°C

-25°C to -15°C

-25°C to -15°C

Instructions

Thaw at room temperature for up to 30 minutes.

Thaw at room temperature for up to 30 minutes.

Thaw at room temperature for up to 30 minutes.

Thaw at room temperature for up to 30 minutes.

2 [Optional] Make 5 µl aliquots of QCT into different PCR tube strips.

3 Place thawed tubes on ice.

Procedure

1 Add 5 µl QCT to 495 µl nuclease-free water in a microcentrifuge tube.

2 Vortex to mix.

3 Add 1 µl QCP to 9 µl nuclease-free water in a microcentrifuge tube.

NOTE

Make a larger dilution if qualifying more than one genomic DNA sample.

4 Vortex to mix.

5 Add 1.5 µl extracted genomic DNA to 148.5 µl nuclease-free water in microcentrifuge tubes to make a 100-fold dilution.

6 Vortex to mix.

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10

7 Determine the plate layout of the qPCR reaction.

For 10 samples, use the following layout.

A

B

C

D

E

F

1

QCT

QCT

QCT

NTC*

NTC*

NTC*

2

Sample 1

Sample 1

Sample 1

Sample 2

Sample 2

Sample 2

3

Sample 3

Sample 3

Sample 3

Sample 4

Sample 4

Sample 4

4

Sample 5

Sample 5

Sample 5

Sample 6

Sample 6

Sample 6

5

Sample 7

Sample 7

Sample 7

Sample 8

Sample 8

Sample 8

NTC: No template control. Illumina recommends using nuclease-free water.

8 Prepare the SYBR master mix reaction as follows.

The master mix contains extra volume.

Consumable

Per well

5.0 µl

Per 48-well plate

275 µl

KAPA SYBR FAST qPCR Master Mix

(2X) (Universal)

Diluted QCP

Nuclease-free water

1.0 µl

2.0 µl

55 µl

110 µl

6

Sample 9

Sample 9

Sample 9

Sample 10

Sample 10

Sample 10

Per 96-well plate

550 µl

110 µl

220 µl

9 Gently invert to mix.

10 Set aside on ice away from light.

11 Add 8 µl of the master mix to each well.

12 According to your plate layout, add 2 µl of the QCT dilution, the sample dilutions, or nuclease-free water to each well of the plate.

13 Seal the plate using an appropriate seal for your qPCR machine.

14 Centrifuge at 250 × g for 1 minute.

15 Make sure that the seal is free of any liquid or dust.

16 Place on the qPCR machine, close the lid, and run the following program.

Procedure

Hot Start x40

Temperature

50°C

95°C

95°C

60°C

72°C

Time

2 minutes

10 minutes

30 seconds

30 seconds

30 seconds

17 Confirm that the instrument captures images after the 72°C step.

NOTE

Set the Cq threshold to a value that avoids inaccurate measurements due to background

(100 RFU on the Bio-Rad 396CFX System).

18 After the final step, the thermal cycler analyzes the quantified libraries. Make sure that amplification of the NTC occurs at least 10 cycles after QCT amplification.

19 Make sure that there is good amplification for the QCT and remove outliers from a triplicate group that are > 0.5 Cq different from the rest of the group.

NOTE

Four or more outliers per plate indicate technical errors.

20 Exclude replicates exhibiting abnormal amplification curves.

Document # 15042911 v02

For more information, see Illumina sequencing white paper, Generating Sequencing

Libraries Using DNA from FFPE Samples.

21 Subtract the average Cq for the QCT from the average Cq for each sample to yield the

ΔCq values for each sample.

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Hybridize the Oligo Pool

During this step, a custom pool containing upstream and downstream oligos specific to your targeted regions of interest is hybridized to your genomic DNA samples.

WARNING

This set of reagents contains formamide, an aliphatic amide that is a probable reproductive toxin. Personal injury can occur through inhalation, ingestion, skin contact, and eye contact. Wear protective equipment, including eye protection, gloves, and laboratory coat. Handle used reagents as chemical waste and discard in accordance with

the governmental safety standards for your region. For environmental, health, and safety information, see the SDS for this kit at support.illumina.com/sds.html

.

NOTE

Illumina does not support the use of gDNA samples giving a delta Cq value of > 4.

Consumables

} [Optional] ACD1 (Amplicon Control DNA1)

}

[Optional] ACP1 (Amplicon Control Oligo Pool 1)

}

FPA (TruSight Tumor Oligo Pool A)

} FPB (TruSight Tumor Oligo Pool B)

} OHS3 (Oligo Hybridization for Sequencing 3)

}

Genomic DNA

}

96-well skirted PCR plate

} [Optional] Adhesive aluminum foil seal (if a heat sealer is not available)

} Troughs

About Reagents

} Use ACPD1 and ACP1 to enable Illumina Technical Support troubleshooting. Include control samples in your assay periodically to establish baselines and monitor overall performance.

}

If you are using controls, use ACD1 instead of gDNA and ACP1 instead of FPA and

FPB.

} ACP1 is specific for Homo sapiens and does not work with DNA from other species.

Preparation

1 Prepare the following consumables.

Item

[Optional] ACD1

Storage

-25°C to -15°C

[Optional] ACP1

FPA

FPB

OHS3

-25°C to -15°C

-25°C to -15°C

-25°C to -15°C

-25°C to -15°C

Genomic DNA -25°C to -15°C

Instructions

Thaw at room temperature for up to 30 minutes.

Thaw at room temperature for up to 30 minutes.

Thaw at room temperature for up to 30 minutes.

Thaw at room temperature for up to 30 minutes.

Thaw at room temperature. If precipitate is observed, incubate at 37°C for 10 minutes and vortex for 1 minute. Repeat as needed until precipitate is no longer visible.

Thaw at room temperature for up to 30 minutes.

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2 Place thawed tubes on ice.

3 Preheat a 96-well heat block to 95°C.

4 Label a new 96-well PCR plate "HYP_Plate_ID".

Procedure

1 Dilute 10 µl of genomic DNA extracted from FFPE samples.

Use the following table to determine the fold dilution required for each calculated delta

Cq.

Delta Cq

Dilution

-2.5 to -1.5

16x

-1.5 to -0.5

8x

-0.5 to 0.5

4x

0.5 to 1.5

2x

1.5 to 4

No dilution

NOTE

If preparing libraries from the control DNA in parallel with FFPE DNA samples, dilute 5

µl of ACD1 with 45 µl of TE Buffer. Add 10 µl to each of 2 control wells for FPA and FPB, and/or 2 control wells for ACP1.

2 Add 10 µl diluted sample to wells on the left half of the plate, starting with column 1.

3 Add 10 µl diluted sample to wells on the right half of the plate, starting with column 7.

4 Add 5 µl FPA to each well containing sample on the left half of the plate.

5 Add 5 µl FPB to each well containing sample on the right half of the plate.

NOTE

If preparing libraries from ACD1, add 5 µl FPA to 1 control well and 5 µl FPB to the second control well. If preparing libraries using ACP1, add 5 µl ACP1 to 2 additional control wells of ACD1.

6 Add 35 µl OHS3 to each well containing sample. Gently pipette to mix.

NOTE

Make sure that there are no crystals or precipitate visible in the OHS3.

7 Seal with a heat sealer or foil seal.

8 Centrifuge at 1000 × g at 20°C for 1 minute.

9 Place on the heat block and incubate for 1 minute.

10 Reduce the heat block to 40°C, and incubate for 14–18 hours.

NOTE

Moving the plate from the 95°C heat block to another preheated block set to 40°C can adversely affect hybridization.

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Remove Unbound Oligos

This process removes unbound oligos from genomic DNA using a filter capable of size selection. Two wash steps using SW1 ensure complete removal of unbound oligos. A third wash step using UB1 removes residual SW1 and prepares samples for the extensionligation step.

Consumables

} ELM4 (Extension-Ligation Mix 4)

}

SW1 (Stringent Wash 1)

}

UB1 (Universal Buffer 1)

} Filter plate with lid (keep spare filter plates as general lab supplies)

} Adapter collar (reusable)

}

Midi plate

}

Troughs

WARNING

This set of reagents contains formamide, an aliphatic amide that is a probable reproductive toxin. Personal injury can occur through inhalation, ingestion, skin contact, and eye contact. Wear protective equipment, including eye protection, gloves, and laboratory coat. Handle used reagents as chemical waste and discard in accordance with

the governmental safety standards for your region. For environmental, health, and safety information, see the SDS for this kit at support.illumina.com/sds.html

.

WARNING

This set of reagents contains ß-mercaptoethanol. Perform the following procedure in a hood or well-ventilated area.

Preparation

1 Prepare the following consumables.

Item

ELM3

SW1

UB1

Storage

-25°C to -15°C

2°C to 8°C

2°C to 8°C

Instructions

Thaw at room temperature for ~20 minutes.

Set aside at room temperature.

Set aside at room temperature.

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2 Assemble the filter plate assembly unit (FPU) in the following order, from top to bottom:

Figure 2 Filter Plate Unit Assembly

A Lid

B Filter plate

C Adapter collar

D midi plate

3 Apply the FPU barcode plate sticker to the filter plate.

4 Prewash the FPU membrane, as follows.

NOTE

Prewash only the wells to be used in the current assay. If using a previously opened filter plate, use only unused wells. Do not reuse wells that have been used in a previous assay.

a Add 50 µl of SW1 to each well.

b Cover with the filter plate lid.

c Centrifuge at 2,400 × g at 20°C for 5 minutes.

5 Preheat the incubator to 37°C.

6 After the overnight incubation, confirm that the heat block has cooled to 40˚C.

NOTE

If the heat block fails to cool to 40˚C overnight, repeat library preparation.

Procedure

NOTE

Cover the FPU with the lid during each centrifugation step.

1 Centrifuge at 1000 × g at 20°C for 1 minute to collect condensation.

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2 Transfer the entire volume to the corresponding wells of the FPU.

3 Centrifuge at 2400 × g at 20°C for 5 minutes.

4 Wash the FPU, as follows.

a Add 50 µl of SW1 to each sample well.

b Centrifuge at 2,400 × g for 5 minutes.

5 Repeat the wash as described in the previous step.

6 Discard waste in a hazardous waste container, and then reassemble the FPU.

The same midi plate can be reused for the rest of the pre-amplification process.

7 Add 45 µl UB1 to each well containing sample.

8 Centrifuge at 2400 × g for 5 minutes.

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Extend and Ligate Bound Oligos

This process connects the hybridized upstream and downstream oligos. A DNA polymerase extends the upstream oligo through the targeted region, and is ligated to the 5’ end of the downstream oligo using a DNA ligase. The ligation step forms products containing the targeted regions of interest flanked by sequences required for amplification.

Consumables

} ELM3 (Extension-Ligation Mix 3)

} Adhesive aluminum foil seal

}

Troughs

Procedure

1 Add 45 µl ELM3 to each well containing sample.

The extension-ligation reaction takes place on the filter plate membrane.

If you use care to avoid cross-contamination, changing tips between columns is not required.

2 Seal with a foil seal, and then cover with the lid.

3 Incubate in the preheated 37°C incubator for 45 minutes.

4 During incubation, prepare the IAP (Indexed Amplification Plate) as described in the following section.

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Amplify Libraries

In this step, the extension-ligation products are amplified using PCR. This step adds index sequences for sample multiplexing (i5 and i7) and common adapters required for amplifying clusters on the flow cell.

Consumables

} PMM2 (PCR Master Mix 2)

} i5 index adapters (A501–A508)

} i7 index adapters (A701–A712)

} TDP1 (TruSeq DNA Polymerase 1)

} 0.05 N NaOH (freshly prepared from 10 N NaOH)

}

96-well skirted PCR plate

}

Microseal 'B' adhesive film

} Troughs

Preparation

1 Prepare the following consumables.

Item

PMM2

Index primers

(i5 and i7)

Storage

-25°C to -15°C

-25°C to -15°C

Instructions

Thaw at room temperature. Vortex to mix, and then briefly centrifuge.

Thaw at room temperature. Vortex each tube to mix, and then briefly centrifuge.

2 To prepare fresh 0.05 N NaOH, add 20 µl of 10 N NaOH to 3.98 ml sterile water.

3 Save the following program as PCR AMP on a thermal cycler with a heated lid.

}

Choose the preheat lid option and set to 100°C

}

95°C for 3 minutes

}

27 cycles of:

} 95°C for 30 seconds

}

62°C for 30 seconds

}

72°C for 60 seconds

}

72°C for 5 minutes

} Hold at 10°C

Procedure

1 Arrange the index primers in the TruSeq Index Plate Fixture, as follows.

} Index 1 (i7) adapters (A701–A712) in columns 1–12

} Index 2 (i5) adapters (A501–A508) in rows A–H

Collect all liquid in the bottoms of the tubes by holding them in place in the rack and tapping it against the bench.

NOTE

If fewer than 48 samples (96 reactions) are being prepared or alternate index combinations are being employed, the Index Plate Fixture is not needed. The indexes can then be added to the appropriate wells of the IAP plate manually.

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Figure 3 TruSeq Index Plate Fixture

A

Columns 1–12: Index 1 (i7) adapters (orange caps)

B

Rows A–H: Index 2 (i5) adapters (white caps)

C

IAP plate

2 Label a new 96-well PCR plate IAP (Indexed Amplification Plate) and place on the

TruSeq Index Plate Fixture.

3 Using a multichannel pipette, add 9 µl of each Index 1 (i7) adapter down each column.

Replace cap on each i7 adapter tube with a new orange cap.

4 Using a multichannel pipette, add 9 µl of each Index 2 (i5) adapter across each row.

Replace the cap on each i5 adapter tube with a new white cap.

NOTE

 After use, remove all index primer tubes from the working area.

5 Prepare the PMM2/TDP1 PCR master mix according to the number of reactions.

} For 96 reactions, add 60 µl of TDP1 to 2.58 ml of PMM2.

}

If preparing fewer reactions, use the following calculation:

Number of reactions × (0.625 µl TDP + 26.875 µl PMM2).

NOTE

Do not pipette volumes of < 5 µl of TDP1.

6 Invert 20 times to mix. Do not vortex.

7 When the 45 minute extension-ligation reaction is complete, remove FPU from the incubator. To ensure the reaction supernatant drains into the waste plate, remove foil seal and replace with the filter plate lid.

8 Centrifuge at 2400 × g for 2 minutes.

9 Add 25 µl of 0.05 N NaOH to each containing sample. Pipette to mix.

10 Incubate at room temperature for 5 minutes.

11 During incubation, transfer 22 µl master mix to each well of the IAP plate containing index adapters.

12 Transfer samples eluted from the FPU plate to the IAP plate, as follows.

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NOTE

Slightly tilt the FPU plate to ensure complete aspiration and to avoid air bubbles.

a Pipette to mix the contents in the first column of the FPU plate.

b Transfer 20 µl from the FPU plate to the corresponding column of the IAP plate.

c Gently pipette to mix.

d Transfer the remaining columns from the FPU plate to the IAP plate in a similar manner.

e After all the samples have been transferred, discard the waste collection midi plate of the FPU. Store the metal adapter collar.

13 Centrifuge 1000 × g at for 1 minute.

14 Transfer the IAP plate to the post-amplification area.

15 On the thermal cycler, set the reaction volume to 60 µl and the temperature ramp speed to maximum.

16 Place on the preprogrammed thermal cycler and run the PCR AMP program.

SAFE STOPPING POINT

If you are stopping, seal the plate and store at 2°C to 8°C for up to 2 days. Alternatively, leave on the thermal cycler overnight.

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Check Library Preparation (Optional)

1 After PCR, combine 5 µl of amplified product with 15 µl of DEPC/DI H20.

2 Run on a 4% TBE agarose gel along with 100 bp ladder to confirm the presence of the

300–330 bp library product. Alternatively, the products can be run on a Bioanalyzer.

If generating libraries with ACP1, expect the product to be present at 350–380 bp.

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Clean Up Libraries

This process uses AMPure XP beads to purify the PCR products from the other reaction components.

Consumables

} EBT (Elution Buffer with Tris)

}

AMPure XP beads

}

96-well midi plates

} Freshly prepared 80% ethanol (EtOH)

} Microseal 'B' adhesive film

}

Troughs

Preparation

1 Bring the AMPure XP beads to room temperature.

2 Prepare fresh 80% ethanol from absolute ethanol.

3 Label a new midi plate CLP_Plate_ID (Clean-up Plate).

4 Label a new 96-well PCR plate SGP (Storage Plate).

Procedure

1 Centrifuge the IAP plate at 1000 × g at 20°C for 1 minute to collect condensation.

2 Invert AMPure XP beads 10 times. Vortex vigorously and then invert again 10 times.

NOTE

Immediately proceed to the next step to avoid settling of the beads.

3 Add 55 µl of AMPure XP beads to each well of the CLP plate.

NOTE

The ACD1/ACP1 control can be processed using the same conditions as your CAT.

4 Transfer 55 µl PCR product from the IAP plate to the CLP plate.

5 Shake at 1800 rpm for 2 minutes.

6 Incubate at room temperature without shaking for 10 minutes.

7 Place on a magnetic stand and wait until the liquid is clear (~2 minutes). Keep the plate on the magnetic stand until step

11

.

8 Remove and discard the supernatant.

NOTE

Delays during this step can lead to bead clumping following removal of the supernatant.

Proceed immediately to next step when all supernatant is removed.

9 Wash 2 times, as follows.

a Add 200 µl freshly prepared 80% EtOH. Avoid disturbing the beads.

b Incubate on the magnetic stand for 30 seconds or until the supernatant appears clear.

c Remove and discard all supernatant from each well.

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10 Remove residual EtOH from each well.

11 Remove the CLP plate from the magnetic stand.

12 Air-dry the beads for 5 minutes.

13 Add 40 µl EBT to each well containing sample.

If you use care to avoid cross-contamination, changing tips is not required.

14 Shake at 1800 rpm for 5 minutes.

After shaking, if any samples are not resuspended, gently pipette or tap on the bench to mix, and then repeat this step.

15 Incubate at room temperature without shaking for 2 minutes.

16 Place the plate on the magnetic stand for 2 minutes.

17 Transfer 40 µl supernatant from the CLP plate to the SGP plate.

18 Centrifuge the SGP plate at 1000 × g for 1 minute.

NOTE

Store the SGP plate at room temperature during the library quantification and normalization steps. After library normalization, store the SGP plate at -25°C to -15°C until ready to quantify and normalize any remaining samples.

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Check Libraries

To achieve the highest data quality, it is important to create optimum cluster densities. This step requires accurate quantification of DNA libraries. Illumina recommends quantifying libraries generated from FFPE samples using the Agilent Technologies Bioanalyzer 2100.

Procedure

1 Load 1 µl of the resuspended library on an Agilent Technologies 2100 Bioanalyzer using the Agilent DNA-1000. 

Refer to the Agilent DNA Kit Guide (Part Number G2938-90014) and Agilent DNA

1000 Kit Quick Start Guide (Part Number G2938-90015) for complete instructions on using the Agilent Technologies 2100 Bioanalyzer.

2 Check the size and purity of the sample. The expected final product is a band at ~300–

330 bp, as in

Figure 4

.

If generating libraries using ACP1, the expected final product is present at ~350–380 bp.

Expect this concentration to correlate with the relative library intensities previously observed on the agarose gel. For more information, see

Check Library Preparation

(Optional) on page 21 .

Figure 4 Representative DNA Sample Prep Library Size Distribution, 300–330 bp

NOTE

If > 10% of the library product is present in the 150–250 bp range, Illumina recommends repeating the PCR Clean-Up procedure. Use 40 µl of product and 32 µl of AMPure XP beads for the repeat procedure.

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Normalize Libraries

This process normalizes the quantity of each library to ensure more equal library representation when libraries are pooled.

Consumables

} EBT (Elution Buffer with Tris)

} Microseal 'B' adhesive film

}

Midi plate

Procedure

1 From the Agilent Bioanalyzer run, determine the concentration for all samples. Add all concentrations (in terms of nM) for all peaks in the 300–330 bp range (or 350–380 bp for ACP1 libraries) and record the values.

The expected concentration range is 4–300 nM.

2 Label a new midi plate LNP_plate (Library Normalization Plate).

3 Dilute 4 µl of all samples > 20 nM to 4 nM with the EBT buffer.

For example, if a sample is 254 nM, add 4 µl of library to 250 µl of EBT buffer to give 4 nM.

4 Dilute at least 20 µl of any samples ≤ 20 nM to 4 nM.

SAFE STOPPING POINT

If you are stopping, seal the plate and store at -25°C to -15°C for up to 7 days.

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Pool Libraries

Pooling libraries combines equal volumes of normalized libraries in a single tube. After pooling, dilute and heat-denature the library pool before loading libraries for the sequencing run.

Consumables

} EBT (Elution Buffer with Tris)

}

Microcentrifuge tubes (screw-cap recommended)

}

PCR 8-tube strip

} Stock 1.0 N NaOH

Preparation

1 If the LNP plate was stored frozen, thaw at room temperature. When thawed, pipette to mix libraries.

2 Centrifuge at 1000 × g at 20°C for 1 minute to collect condensation.

3 Determine the libraries to be pooled.

Procedure

Sequence 4 tumor samples (8 libraries total) per run. If sequencing a different number of samples, adjust the following procedure accordingly.

NOTE

Two libraries represent each sample and are generated from the TruSight Tumor Oligo

Pools (FPA and FPB). The FPA and FPB libraries for each sample must be run together on the same flow cell to analyze the results for each sample.

NOTE

Control libraries generated from ACD1 with ACP1 must be pooled and run separately from those libraries prepared with FPA and FPB, because they require a run of 151 cycles.

1 Add 10 µl of 1N NaOH to 140 µl EBT buffer. Vortex to mix.

2 Transfer 5 µl of each 4 nM library from the LNP plate to its own tube in a PCR 8-tube strip.

NOTE

After use, the sealed LNP plate can be stored at -25°C to -15°C for up to 7 days.

3 Add 15 µl NaOH/EBT solution to each 5 µl of library.

4 Incubate for 5 minutes at room temperature.

5 Label a microcentrifuge tube PAL (Pooled Amplicon Library).

6 Add 10 µl of each library/NaOH/EBT solution to the PAL tube.

7 Pipette to mix.

8 Denature and dilute pooled libraries to the loading concentration for the instrument you are using. See the denature and dilute libraries guide for your instrument.

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