Apr 03, 2026

Whole Genome Sequencing Protocol using Illumina NextSeq 1000 Platform

DOI
https://dx.doi.org/10.17504/protocols.io.ewov1b36pgr2/v1
  • Ma. Carmel Javier1,
  • Jasmine Velo1,
  • Nadelene Faith Cartago1,
  • Victor Marco Emmanuel Ferriols1
  • 1Philippine Genome Center Visayas
Jasmine Velo
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DOI: https://dx.doi.org/10.17504/protocols.io.ewov1b36pgr2/v1
Protocol Citation: Ma. Carmel Javier, Jasmine Velo, Nadelene Faith Cartago, Victor Marco Emmanuel Ferriols 2026. Whole Genome Sequencing Protocol using Illumina NextSeq 1000 Platform. protocols.io https://dx.doi.org/10.17504/protocols.io.ewov1b36pgr2/v1
License: This is an open access  protocol  distributed under the terms of the  Creative Commons Attribution License,  which permits unrestricted use, distribution, and reproduction in any medium, provided the original author and source are credited
Protocol status: Working
We use this protocol and it's working
Created: March 05, 2026
Last Modified: April 03, 2026
Protocol  Integer ID: 255669
Keywords: whole genome sequencing protocol, illumina nextseq 1000 platform, illumina nextseq 1000 system, whole genome sequencing, workflow for whole genome sequencing, sequencing protocol, illumina dna prep kit, using illumina nextseq, genomic dna library preparation, sequencing, illumina nextseq, whole genome, genome, nextseq, using illumina, wg, dna, illumina
Abstract
This protocol describes the workflow for whole genome sequencing (WGS) using the Illumina NextSeq 1000 System. The procedure includes genomic DNA library preparation using the Illumina DNA Prep Kit, followed by high-throughput sequencing to generate paired-end reads.
DNA INPUT RECOMMENDATIONS:
The Illumina DNA Prep protocol is compatible with DNA inputs of 1–500 ng or higher. For human DNA samples and other large complex genomes, the recommended minimum DNA input is 100–500 ng. For small genomes, such as microbial, the DNA input amount can be reduced to as low as 1 ng. If reducing the DNA input amount, modify the PCR cycling conditions accordingly.
STEP 1: TAGMENT AMPLICONS
REAGENTS NEEDED:
● EBLTS HT (Enrichment BLT HT)
● TB1 HT (Tagmentation Buffer 1 HT)
● Nuclease-free water
About the Reagents
  • Store EBLTS HT upright at temperatures above 2°C. Make sure beads are always submerged in the buffer.
  • If beads are adhered to the side or top of the 96-well plate, centrifuge at 500 × g for 1 minute, and then pipette to resuspend.
ABC
Reagent Storage Instructions
EBLTS HT 2°C to 8°C Thaw at room temperature. Vortex thoroughly before use
TB1 HT -25°C to -15°C Thaw at room temperature. Vortex thoroughly before use
*Resuspend EBLTS HT by vortexing. Maintain at 4℃.

Label a new 96-well PCR plate as TAG1, Run #, and Date. Prepare Tagmentation Master Mix by combining the following. Multiply each volume by the number of samples.
TB1 HT (Tagmentation Buffer 1 HT) 12 ul
EBLTS HT (Enrichment BLT HT) 4 ul
NFW 20 ul

  • Add 30 ul master mix to each well of TAG1 plate.
NOTE: Do reverse pipetting using a single-channel pipette.
  • Make sure that the pipette tips will not touch the sides of the well. Discard and change pipette tips if touched.
  • After adding master mix, add 20 ul of PCR product to each corresponding well in TAG1 plate.
  • Seal and shake the TAG1 plate at 1600 rpm for 1 minute.
  • Centrifuge at 1000 x g for 1 minute if the contents of the well are found sticking to the walls or the seal of the TAG1 plate.
  • Place on a thermal cycler and run the following program:
  • Choose the preheat lid option
55℃ for 5 minutes
Hold at 10℃
  • Set the reaction volume to 50 μl.

STEP 2: POST TAGMENTATION AND CLEAN UP
CONSUMABLES
● ST2 HT (Stop Tagment Buffer 2 HT)
● TWB HT (Tagmentation Wash Buffer HT)
● Microseal B adhesive seal

REAGENTS
● Dispense ST2 HT and TWB HT slowly to minimize foaming.
● Dispense TWB HT directly onto beads
ABC
Reagent Storage Instructions
ST2 HT Room temperature Vortex before use.
TWB HT 2°C to 8°C Vortex before use.

PROTOCOL
  • Centrifuge the TAG1 plate at 500 x g for 1 minute.
  • Dispense 10ul of ST2 HT (Stop Tagment Buffer 2 HT) to each well of PCR 8-strip tube. Multiply each well by the number of samples per row. Add 5 ul in each tube/row for pipetting error.
  • Carefully remove the seal of TAG1 plate and add 10 μl of ST2 HT (Stop Tagment Buffer 2 HT) from the PCR 8-strip tube to each well of the TAG1 plate.
CRITICAL: Immediately add ST2 HT after PCR to stop the tagmentation.
  • Seal the plate and shake at 1600rpm for 1 minute
  • Incubate the plate at room temperature for 5 minutes. Note: Can start the timer while shaking.
  • Centrifuge at 500 x g for 1 minute.
  • Remove the seal carefully and place the plate on a magnetic stand.
  • Wait for 3 minutes or until the liquid is clear.
  • Inspect for bubbles on the seal. If present, centrifuge at 500 x g for 1 minute, and then place on the magnetic stand (~ 3 minutes)
  • Remove and discard all supernatants from the wells. Use a reservoir for waste.
  • Wash the beads as follows:
  • Remove the plate from the magnetic stand.
  • Add 100μl of TWB HT (Tagmentation Wash Buffer HT) to each well. Note: Place the TWB HT on the pipette reservoir.
  • Seal and shake at 1600 rpm for 1 minute
  • Centrifuge at 500 x g for 1 minute.
  • Remove the seal and place it on the magnetic stand and wait for 3 minutes or until the liquid is clear.
  • For the first wash only, remove and discard all supernatants from each well.
  • Repeat Wash beads
CRITICAL: Do not remove the plate from the magnetic stand. Do not discard the supernatant after the second wash to prevent the beads from over-drying.
step 3: AMPLIFY TAGMENTED AMPLICONS
CONSUMABLES AND REAGENTS
● EPM HT (Enhanced PCR Mix HT)
● Index adapters (IDT for Illumina-PCR Indexes Set 1, 2, 3, 4)
● Nuclease-free water
● 15 ml tubes
● 96 well PCR Plates

Reagent Storage Instructions
EPM HT -25°C to -15°C Invert to mix. Keep on ice until use.
Index adapters -25°C to -15°C Thaw at room temperature. Vortex to mix, and then centrifuge at 1000 × g for 1 minute.
PROTOCOL
  • Prepare PCR Master Mix by combining thefollowing:
AB
EPM HT (Enhanced PCR Mix HT) 24 ul
NFW 24 ul
  • Vortex PCR Master Mix
  • Remove and discard the supernatant (TWB HT) from the TAG1 plate.
CRITICAL: Do not remove supernatant (TWB HT) from all samples all at once to avoid over-drying of beads. Remove supernatant from the first 2 columns first.
  • Use a 10 μl or 20 μl pipette to remove any remaining TWB HT.
  • Add 40 μl of PCR Master Mix to each well.
NOTE: Move columns away from the magnetic stand before adding master mix. Add master mix directly to the beads.
  • Add 10 μl index adapters to each well.
NOTE: One index well per sample well. Index wells are sometimes empty. Check for volume before dispensing. Record index added to each sample well.
  • Seal and shake the plate at 1600 rpm for 1 minute and centrifuge the plate at 500 x g for 1 minute. CRITICAL: Inspect the tubes to make sure the beads are fully re-suspended and are not found at the bottom of the tubes. To resuspend, set the pipette to 35ul with plunger down, and then slowly pipette to mix.
  • Place on a thermal cycler and run the following PCR program:
  • Choose the preheat lid option and set the temperature to 100°C.
● 72°C for 3 minutes
● 98°C for 3 minutes
● 7 cycles of:
○ 98°C for 20 seconds
○ 60°C for 30 seconds
○ 72°C for 1 minute
● 72°C for 3 minutes
● Hold at 10°C
● Set reaction volume to 50 ul

STEP 4: POOL AND CLEAN UP LIBRARIES
CONSUMABLES AND REAGENTS
● ITB (Illumina Tune Beads)
● RSB HT (Resuspension Buffer HT)
● Freshly prepared 80% ethanol (EtOH)
● 1.5 ml tube (2 per 96-well sample plate)
● PCR 8-tube strip

Reagent Storage Instructions
ITB Room temperature Vortex thoroughly to mix.
RSB HT 2°C to 8°C Let stand for 30 minutes to bring to room temperature. Vortex and invert to mix.
PROTOCOL
  • Centrifuge the TAG1 plate at 500 x g for 1 minute.
  • Carefully remove the seal and place the plate on a magnetic stand and wait for 3 minutes or until the liquid is clear.
Pool libraries as follows:
  • Transfer 5 μl of library from each well of the plate to PCR 8-tube strip. This results in a total volume of 60 ul per row.
  • CRITICAL: Discard and change tips after each column of samples.
  • Transfer 55 μl of the pooled library from each well of the PCR 8-tube strip into a 1.5 ml microcentrifuge tube (Label as ITB tube). The total volume is 440 μl of pooled libraries (from 96 samples). If processing 3072 samples, these steps result in 32 Pooled ITB tubes.
  • Vortex the Pooled ITB tubes to mix, and spin down.
  • Vortex ITB (Illumina Tube Beads) to resuspend.
  • Add ITB using the resulting volume of Pooled ITB tube volume multiplied by 0.9X. (For example, for 96 samples: 440 μl ITB tube volume x 0.9 = add 396 μl). For NTC: 20 ul x 0.9x = 18 ul).
  • Vortex to mix.
  • Incubate the tube at room temperature for 5 minutes.
  • Centrifuge briefly.
  • Place the tube on the magnetic stand and wait for 5 minutes until the liquid is clear.
  • Remove and discard all supernatants.
  • Wash the beads as follows:
- Keep on the magnetic stand and add 1000 μl of freshly prepared 80% ethanol to each tube. Note: 80% ethanol can be less than 1000 μl long as the beads are submerged.
- Wait for 30 seconds.
- Remove and discard all supernatants.
- Wash beads a second time.
- Use a 20 ul pipette to remove all residual ethanol.
- Air-dry the beads for at least 2 minutes. NOTE: Do not over-dry the beads.
- Remove the tube from the magnetic rack. Add 55μl of RSB HT (Re-suspension Buffer). NOTE: 55 μl is good for 96 samples
- Vortex to mix and centrifuge briefly.
- Incubate at room temperature for 2 minutes.
- Place the tube on the magnetic stand and wait for 2 minutes until the liquid is clear.
- Transfer 50 μl of the supernatant from each Pooled ITB tube to a new low-bind microcentrifuge tube. Note: Leave at least 2 ul to avoid beads.
  • REPEAT steps f – t using a 1:1 ITB and sample ratio for second ITB clean up.
  • Label pooled library tube.
STEP 5: QUANTIFY AND NORMALIZE LIBRARIES
  • Analyze 2 µl library pool, NTC and PC using a Qubit dsDNA HS Assay kit. If libraries are outside the standard range, dilute to 1:10 concentration, and analyze again.
  • Using Agilent Bioanalyzer, estimate the size of the library using ~300bp as the average library size.
  • Calculate the molarity value using the following formula:



STEP 6: DILUTE LIBRARIES
NOTE: Use the excel sheet Library Dilution format for easy calculation. Always double check values before proceeding.
  • NextSeq 1000
  1. Dilute 5ul of the library to 4nM using the following formula:



  • Qubit 2nd Reading: Analyze 2 µl of 4nM library pool Qubit dsDNA HS Assay kit to obtain the exact concentration. If libraries are outside the standard range, dilute to 1:10 concentration, and analyze again.Calculate molarity using the library concentration formula above (Step 8: C)
  • Dilute to 2nM using the following formula. Note: Use the exact 2nd qubit concentration of 4nM Library.


  • Dilute to final loading concentration of 1nM/1000pM using the following formula:


STEP 7: LOAD LIBRARY
NextSeq 1000
a. Load 20ul of diluted library with final concentration of 1000pM.
b. Start Run.