WGBS library prep (Zymo Pico Methyl-Seq)

DNA methylation
Porites astreoides
Bisulfite-convert 10 ng of coral genomic DNA and build indexed whole-genome bisulfite sequencing libraries with the Zymo Pico Methyl-Seq kit, with QC and a re-amplification rescue.
Author

Kevin H. Wong

Published

October 5, 2026

Version 1.0 · last updated 2026-10-05 · Used in 11 notebook entries ↓

At a glance

Input 10 ng genomic DNA (1 ng µL⁻¹ in 10 mM Tris-HCl)
Output Dual-indexed WGBS library, ~15 µL, QC’d by Qubit and D5000 TapeStation
Kit Zymo Pico Methyl-Seq Library Prep Kit + KAPA Pure Beads
Time 1.5 days: dilution + conversion in the afternoon (overnight stop), then cleanup through QC the next day
Tested on Porites astreoides adults and larvae

This is an adapted workflow, not a replacement for the kit manual. Follow the Zymo Pico Methyl-Seq manual for the full reagent details. The changes from the manual are 10 ng input (instead of 1 ng) with fewer amplification cycles, and a diluted polymerase so you never pipette 0.3 µL.

flowchart LR
  A[10 ng DNA] --> B[Bisulfite<br/>conversion]
  B --> C[Column<br/>cleanup]
  C --> D[PrepAmp<br/>random priming]
  D --> E[DCC]
  E --> F[Amp 1<br/>8 cycles]
  F --> G[DCC]
  G --> H[Amp 2 + UDI index<br/>10 cycles]
  H --> I[1× bead<br/>cleanup]
  I --> J[(Qubit +<br/>TapeStation)]

Materials

  • Zymo Pico Methyl-Seq Library Prep Kit; KAPA Pure Beads (Roche)
  • Unique dual index (UDI) i5/i7 primers, 10 µM. Plan the pairs before you start (see Before you start)
  • 10 mM Tris-HCl pH 8.0; 100% ethanol (fresh 80% each day); ultrapure water
  • Thermocycler, thermomixer (56 °C), microcentrifuge (16,000 × g), magnetic plate for PCR strips, tube rotator/shaker
  • PCR strip tubes, 1.5 mL tubes, filter tips
  • Qubit dsDNA BR assay; Agilent TapeStation with D5000 ScreenTape and reagents

Before you start

  • Dilute DNA to 1 ng µL⁻¹ in 10 mM Tris-HCl. Calculate all dilutions ahead of time.
  • Plan indexes: every library gets a unique i5 and a unique i7. Don’t reuse an index number within a pool, even in a different pairing (this limits index hopping). Putnam Lab UDI list: UDI index primer pairs.
  • Thermocycler programs to save: BS conversion, PrepAmp priming, Amp 1 and Amp 2 (cycle numbers below).
  • Work out n = number of samples + ~5% for every master mix.
  • While each step runs, prepare the master mixes for the next step on ice.

Procedure

Day 1: Bisulfite conversion

  1. In PCR strips: 10 µL DNA (10 ng) + 10 µL 10 mM Tris-HCl = 20 µL.
  2. Add 130 µL Lightning Conversion Reagent (150 µL total). Vortex and spin down.
  3. Run: 98 °C 8 min → 54 °C 60 min → 4 °C hold (~1.5 h).
Tip⏸ Safe stopping point

Converted DNA is stable at 4 °C for up to 20 h. Stop here overnight.

Day 2

Post-conversion column cleanup

  1. Add 600 µL M-Binding Buffer to a spin column, then the whole 150 µL conversion. Cap, invert 5–10×, spin 30 s. Discard flow-through into the kit’s hazardous-waste container.
  2. 100 µL M-Wash, spin 30 s.
  3. 200 µL L-Desulphonation Buffer, stand 15 min at room temp, spin 30 s. Meanwhile warm the DNA Elution Buffer to 56 °C.
  4. 200 µL M-Wash, spin 30 s. Repeat, spinning 2 min the second time.
  5. Move the column to a 1.5 mL tube. Add 8 µL warm Elution Buffer to the membrane, wait 5 min, spin 30 s. Keep on ice and check all tubes have the same volume.

PrepAmp (random priming)

  1. Master mixes (× n), on ice:

    Mix Per sample
    PMM (priming) 2 µL 5× PrepAmp Buffer + 1 µL 40 µM PrepAmp Primer
    PAMM (PrepAmp) 1 µL 5× PrepAmp Buffer + 3.75 µL PrepAmp Pre-mix + 0.3 µL PrepAmp Polymerase
    Diluted polymerase 0.3 µL PrepAmp Polymerase + 0.2 µL Elution Buffer (so you add 0.5 µL)

  2. 7 µL converted DNA + 3 µL PMM in new strips. Vortex, spin, and start the priming program:

    Step Temp / time
    Denature 98 °C 2 min → 8 °C 1 min → 8 °C hold
    ① At the hold Take tubes out, vortex/spin, add 5.05 µL PAMM, vortex/spin, return and continue
    Priming 8 °C 4 min, then 16, 22, 28, 36, 36.5 °C for 1 min each (3% ramp), 37 °C 8 min
    Repeat Denature → hold again
    ② At the hold Add 0.5 µL diluted polymerase, vortex/spin, continue the priming steps

DNA Clean & Concentrator (DCC) 1

  1. Add the whole priming reaction (~15.5 µL) to 108.85 µL DNA Binding Buffer (7:1), vortex, load a spin column, spin 30 s.
  2. 200 µL DNA Wash, spin 30 s; repeat with a 2 min spin.
  3. Elute in 12 µL warm Elution Buffer (5 min, spin 30 s).

Amplification 1

  1. Per sample: 12.5 µL 2× Library Amp Mix + 1 µL 10 µM Library Amp Primers (13.5 µL) + 11.5 µL DCC eluate.
  2. Run Amp 1 with 8 cycles for 10 ng input (10 cycles for 1 ng) (~30 min).

DCC 2

  1. 25 µL reaction + 175 µL DNA Binding Buffer (7:1). Bind, wash 2× as above, elute in 12.5 µL warm Elution Buffer.

Amplification 2 (indexing)

  1. Per sample: 12 µL DNA + 14 µL 2× Library Amp Mix + 1 µL i5 + 1 µL i7 (10 µM). Tick each addition off on a sheet as you go.
  2. Run Amp 2 with 10 cycles for 10 ng input (12 for 1 ng) (~30 min). Take the KAPA beads out of the fridge now.

1× bead cleanup

  1. Beads at room temp (≥ 30 min), swirled not vortexed. Make fresh 80% ethanol.
  2. Add 26 µL beads (1×) to each library. Pipette-mix ≥ 10× and rotate 15 min at 200 rpm.
  3. On the magnet, wait until clear, then remove the supernatant.
  4. Wash 2× with 200 µL 80% ethanol without disturbing the beads. Remove all residual liquid with a P20 and air-dry ~1 min.
  5. Off the magnet, resuspend in 15 µL Elution Buffer, shake 5 min at 200 rpm, return to the magnet and transfer the clear eluate to labeled final tubes (sample, date, “lib”, initials).
Tip⏸ Safe stopping point

Libraries can go to −20 °C here if you’re not doing QC the same day.

QC and expected results

Check Method Good result
Concentration Qubit dsDNA BR, read twice and average > 5 ng µL⁻¹ (re-amplify below that, see below)
Size D5000 TapeStation Smooth peak in the same range as the other libraries; no library with a very different size profile

In the Thermal Transplant preps, libraries came out at about 3–22 ng µL⁻¹. An example of a good trace that was sequenced:

TapeStation D5000 trace of a good WGBS library

Rescue: re-amplify low-yield libraries (< 5 ng µL⁻¹)

Repeat Amp 2 + bead cleanup with 4 cycles: 12 µL library + 13 µL 2× Library Amp Mix + 1 µL combined i5/i7 (the same indexes as before), then a 1× bead cleanup. In 2021 this brought most low libraries to 7–27 ng µL⁻¹. A few libraries that read “too low” stayed too low and need to be re-prepped from DNA.

Troubleshooting

Problem Likely cause Fix
Library fails (Qubit “too low”) Unequal elution after conversion, or sample loss in the cleanups Check volumes after step 8; elute slowly onto the membrane center
Less than 15.5 µL after priming Evaporation (priming program has a low lid temperature) Expected; transfer everything into the binding buffer
0.3 µL polymerase doesn’t leave the tip Volume too small Use the diluted polymerase (0.5 µL)
Library yield < 5 ng µL⁻¹ Low input or efficiency Re-amplify with 4 cycles (above)

References

Notebook entries using this protocol

Changelog

Date Version Change
2026-10-05 1.0 Rewritten from the 2021 Thermal Transplant protocol and library-prep trials; re-amplification rescue added
2021-05 – Re-amplification of low libraries (4 cycles)
2021-04 – 10 ng input with reduced Amp 1/Amp 2 cycles; eight library-prep batches
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