laboratory

Nexterra Library Prep

• 1. Tagment Genomic DNA
  • Preparation
  • Procedure
• 2. Post Tagmentation Cleanup
  • Preparation
  • Procedure
• 3. Amplify Tagmented DNA
  • Preparation
  • Procedure
• 4. Clean Up Libraries
  • Preparation
  • Procedure
• 5.Pool Libraries
  • For DNA Inputs of < 100 ng

1. Tagment Genomic DNA

Preparation

1. Prepare the following consumables:

Item	Storage	Instructions
BLT	2°C to 8° C	Bring to room temperature. Vortex to mix. Do not centrifuge before pipetting.
TB1	-25°C to ‑15°C	Bring to room temperature. Vortex to mix.

1. Save the following TAG program on the thermal cycler:

• Choose the preheat lid option and set to 100°C
• Set the reaction volume to 50 µl
• 55°C for 15 minutes
• Hold at 10°C

Procedure

1. Add 2–30 µl DNA to each well of a 96-well PCR plate so that the total input amount is 100–500 ng.
2. If DNA volume < 30 µl, add nuclease-free water to the DNA samples to bring the total volume to 30 µl.
3. Vortex BLT vigorously for 10 seconds to resuspend. Repeat as necessary.
4. Combine the following volumes to prepare the tagmentation master mix. Multiply each volume by the number of samples being processed.

• BLT (11 µl)
• TB1 (11 µl)

1. Vortex the tagmentation master mix thoroughly to resuspend.
2. Divide the tagmentation master mix volume equally into an 8-tube strip.
3. Using a 200 µl multichannel pipette, transfer 20 μl tagmentation master mix to each well of the plate containing a sample. Use fresh tips for each sample column.
4. Discard the 8-tube strip after the tagmentation master mix has been dispensed.
5. Pipette each sample 10 times to resuspend. Use fresh tips for each sample column.
6. Seal the plate with Microseal ‘B’, place on the preprogrammed thermal cycler, and run the TAG program.

2. Post Tagmentation Cleanup

Preparation

1. Prepare the following consumables:

Item	Storage	Instructions
TSB	15°C to 30°C	If precipitates are observed, heat at 37°C for 10 minutes, and then vortex until precipitates are dissolved. Use at room temperature.
TWB	room temperature	Vortex thoroughly to mix.

1. Save the following PTC program on the thermal cycler:

• Choose the preheat lid option and set to 100°C
• Set the reaction volume to 50 µl
• 55°C for 15 minutes
• Hold at 10°C

Procedure

1. Add 10 µl TSB to the tagmentation reaction.
2. Slowly pipette each well 10 times to resuspend the beads.
3. Seal the plate with Microseal ‘B’, place on the preprogrammed thermal cycler, and run the PTC program.
4. Place the plate on the magnetic stand and wait until liquid is clear (~3 minutes).
5. Using a multichannel pipette, remove and discard supernatant.
6. Wash two times as follows:
   1. Remove the sample plate from the magnetic stand and use a deliberately slow pipetting technique to add 100 µl TWB directly onto the beads. A deliberately slow pipetting technique minimizes the potential of TWB foaming to avoid incorrect volume aspiration and incomplete mixing.
   2. Pipette slowly until beads are fully resuspended.
   3. Place the plate on the magnetic stand and wait until the liquid is clear (~3 minutes).
   4. Using a multichannel pipette, remove and discard supernatant.
7. Remove the plate from the magnetic stand and use a deliberately slow pipetting technique to add 100 µl TWB directly onto the beads.
8. Pipette each well slowly to resuspend the beads.
9. Seal the plate and place on the magnetic stand until the liquid is clear (~3 minutes). Keep on the magnetic stand until step 4 of the Procedure section in Amplify Tagmented DNA.

3. Amplify Tagmented DNA

Preparation

1. Prepare the following consumables:

Item	Storage	Instructions
EPM	‑25°C to‑15°C	Thaw on ice. Invert to mix, then briefly centrifuge.
Index Adapters	‑25°C to‑15°C	Thaw at room temperature. Vortex to mix, then centrifuge briefly. Spin briefly before use.

1. Save the following BLT PCR program on a thermal cycler using the appropriate number of PCR cycles, indicated in the table below.

• Choose the preheat lid option and set to 100°C
• 68°C for 3 minutes
• 98°C for 3 minutes
• X cycles of:
  • 98°C for 45 seconds
  • 62°C for 30 seconds
  • 68°C for 2 minutes
• 68°C for 1 minutes
• Hold at 10°C

Total DNA Input (ng)	Number of PCR Cycles (X)
1–9	12
10–24	8
25–49	6
50–99	5
100–500	5

Procedure

1. Combine the following volumes to prepare the PCR master mix. Multiply each volume by the number of samples being processed.

• EPM (22 µl)
• Nuclease-free water (22 µl)

1. Vortex, and then centrifuge the PCR master mix at 280 × g for 10 seconds.
2. With the plate on the magnetic stand, use a 200 µl multichannel pipette to remove and discard supernatant.
3. Remove from the magnet.
4. Immediately add 40 µl PCR master mix directly onto the beads in each sample well.
5. Immediately pipette to mix until the beads are fully resuspended. Alternatively, seal the plate and use a plate shaker at 1600 rpm for 1 minute.
6. Seal the sample plate and centrifuge at 280 × g for 3 seconds.
7. Add the appropriate index adapters to each sample.

Index Kit Type	Kit Configuration	Volume of Index Adapter per Sample
24 plex (dual index)	Individual tubes	5 µl i7 adapter, 5 µl i5 adapter
96 plex (dual index)	96-well plate	10 µl pre-paired i7 and i5 index adapters

1. Using a pipette set to 40 µl, pipette 10 times to mix. Alternatively, seal the plate and use a plate shaker at 1600 rpm for 1 minute.
2. Seal the plate with Microseal ‘B’, and then centrifuge at 280 × g for 30 seconds.
3. Place on the thermal cycler and run the BLT PCR program.

SAFE STOPPING POINT

If you are stopping, store at 2°C to 8°C for up to 3 days.

4. Clean Up Libraries

Preparation

1. Prepare the following consumables:

Item	Storage	Instructions		PB	2°C to 8°C	Let stand at room temperature
for 30 minutes. Vortex and invert to mix.		RSB	-25°C to -15°C	Thaw and
bring to room temperature. Vortex to mix.

1. Prepare fresh 80% EtOH from absolute ethanol.

Procedure

1. Centrifuge at 280 × g for 1 minute to collect contents at the bottom of the well.
2. Place the plate on the magnetic stand and wait until the liquid is clear (~5 minutes).
3. Transfer 45 µl supernatant from each well of the PCR plate to the corresponding well of a new midi plate.
4. Vortex and invert PB multiple times to resuspend.
5. For standard DNA input, perform the following steps.
   1. Add 40 µl nuclease-free water to each well containing supernatant.
   2. Add 45 µl PB to each well containing supernatant.
   3. Pipette each well 10 times to mix. Alternatively, seal the plate and use a plate shaker at 1600 rpm for 1 minute.
   4. Seal the plate and incubate at room temperature for 5 minutes.
   5. Place on the magnetic stand and wait until the liquid is clear (~5 minutes).
   6. During incubation, thoroughly vortex the PB (undiluted stock tube), and then add 15 µl to each well of a new midi plate.
   7. Transfer 125 µl supernatant from each well of the first plate into the corresponding well of the second plate (containing 15 µl undiluted PB).
   8. Pipette each well in the second plate 10 times to mix. Alternatively, seal the plate and use a plate shaker at 1600 rpm for 1 minute.
   9. Discard the first plate.
6. For small PCR amplicon input, perform the following steps.
   1. Add 81 µl PB to each midi plate well containing supernatant.
   2. Pipette each well 10 times to mix. Alternatively, seal the plate and use a plate shaker at 1600 rpm for 1 minute.
7. Incubate the sealed midi plate at room temperature for 5 minutes.
8. Place on the magnetic stand and wait until the liquid is clear (~5 minutes).
9. Without disturbing the beads, remove and discard supernatant.
10. Wash two times as follows.
    1. With the plate on the magnetic stand, add 200 µl fresh 80% EtOH without mixing.
    2. Incubate for 30 seconds.
    3. Without disturbing the beads, remove and discard supernatant.
11. Use a 20 µl pipette to remove and discard residual EtOH.
12. Air-dry on the magnetic stand for 5 minutes.
13. Remove from the magnetic stand.
14. Add 32 µl RSB to the beads.
15. Pipette to resuspend.
16. Incubate at room temperature for 2 minutes.
17. Place the plate on the magnetic stand and wait until the liquid is clear (~2 minutes).
18. Transfer 30 µl supernatant to a new 96-well PCR plate.

SAFE STOPPING POINT

If you are stopping, seal the plate with Microseal ‘B’ adhesive or Microseal ‘F’ foil seal, and store at ‑25°C to ‑15°C for up to 30 days.

5.Pool Libraries

When the DNA input is 100-500 ng, quantifying and normalizing individual libraries generated in the same experiment is not necessary. However, the final yield of libraries generated in separate experiments can vary slightly. To achieve optimal cluster density, pool equal library volumes and quantify the pool before sequencing.

For DNA Inputs of < 100 ng

1. Quantify each library individually using Qubit or PicoGreen.

Check Library Quality (Optional)

1. Run 1 µl library or pooled libraries on one of the following instruments:

• Add 1 µl RSB to the library to achieve the 2 µl volume required for Fragment Analyzer. The following figures show typical library size profiles with an average fragment size of 600 bp when analyzed with a size range of 150–1500 bp. Dilute Libraries to the Starting Concentration

Table 1: Recommended Read Length on Illumina Systems

Sequencing System	Read Length
NovaSeq 6000	2 x 151

Assumes use of the 200 cycle kit.

1. Calculate the molarity value of the library or pooled libraries using the following formula.

• For libraries qualified on a Bioanalyzer, use the average size obtained for the library.
• For all other qualification methods, use 600 bp as the average library size.

1. Using the molarity value, calculate the volumes of RSB and library needed to dilute libraries to the starting concentration for your system.

Sequencing System	Starting Concentration (nM)	Final Loading Concentration (pM)
NovaSeq 6000	2	See document # 1000000019358 (NovaSeq 6000 System Guide)

1. Dilute libraries using RSB:

• Libraries quantified as a multiplexed library pool —Dilute the pool to the starting concentration for your system.
• Libraries quantified individually —Dilute each library to the starting concentration for your system. Add 10 µl each diluted library to a tube to create a multiplexed library pool.