Bisulfite amplicon sequencing library preparation (using TruSeq adapters)

Reagents

• Bisulfite‑treated DNA (≥1 ng/µL recommended; A260/280 ~1.8)

• U‑tolerant EpiMark Hot Start Taq for PCR-1

• LongAmp Taq for PCR‑2 (you may reuse the U‑tolerant enzyme if preferred)

• dNTP mix (10 mM each) – if not included in Master Mix

• Primers (HPLC‑purified/desalted):

  • PEP_BS_01_L and PEP_BS_01_R (10 µM working stocks)

  • Indexing primers (TruSeq HT style):

    • P5/i5 primer: AATGATACGGCGACCACCGAGATCTACAC + [i5 index] + ACACTCTTTCCCTACACGACGCTCTTCCGATCT
    • P7/i7 primer: CAAGCAGAAGACGGCATACGAGAT + [i7 index] + GTGACTGGAGTTCAGACGTGTGCTCTTCCGATCT

• AMPure XP (or SPRIselect) beads + 80% ethanol (fresh)

• Elution buffer (10 mM Tris‑HCl, pH 8.5) or nuclease‑free water

• Qubit dsDNA HS kit (or equivalent)

• Agarose + Gel stain (for quick size check) or Bioanalyzer/TapeStation (preferred)

• PhiX control (10 nM or 4 nM stock from Illumina; denature per kit)

• MiSeq Reagent Kit (v2 2×250 or v3 2×300)

• 0.2 N NaOH and HT1 (hybridization buffer) for denaturation/dilution (per Illumina kit)

Equipment

• Thermal cycler with heated lid
• Magnetic stand for 0.2/0.5 mL tubes or 96‑well plates
• Microcentrifuge, pipettes, filtered tips
• Blue‑light gel system or capillary electrophoresis (FA/BA)
• MiSeq instrument

Primer preparation

1. Reconstitute lyophilized primers in nuclease‑free water or 10 mM Tris‑HCl pH 8.5 to 100 µM stocks.
2. Prepare 10 µM working stocks (store at −20 °C).

PCR‑1: Amplify bisulfite DNA with tailed primers

Goal: Generate ~558‑bp product containing your target + Read‑primer overhangs.

Reaction (25 µL per sample)

• 5 µL 5× EpiMark buffer
• 0.5 µL 10 µM dNTP mix (200 µM each)
• 0.5 µL 10 µM Primer 1 (0.2 µM each TruSeq‑tailed primer)
• 0.5 µL 10 µM Primer 2
• 0.125 µL EpiMark Hot Start Taq (~0.625 U / 25 µL)
• 1-4 µL (1–10 ng) bisulfite DNA
• Optional 3–5% DMSO for very AT‑rich regions
• Make up to to 25 µL with ultra pure water

Cycling (typical, optimize once):

• 95 °C 5 min
• 40 cycles of 95 °C 30 s / 58 °C 30 s / 68 °C 30 s (≈30 s per ~600 bp)
• 68 °C 3 min, 4 °C hold

Notes: • Anneal temp should be set by the locus‑specific Tm (do not use overhangs). 55–60 °C works well; start at 58 °C. • Use a no‑template control (NTC) to monitor primer dimers.

Cleanup (1.0× SPRI):

1. Vortex beads, add 25 µL beads to 25 µL PCR. Mix 10×.
2. Incubate 5–10 min at RT; place on magnet 5 min; remove supernatant.
3. Wash twice with 200 µL 80% EtOH, 30 s each.
4. Air‑dry beads 2–5 min (do not over‑dry).
5. Elute in 22–30 µL EB; transfer eluate.

QC:

• Qubit concentration ≥ 1–5 ng/µL is usually adequate.
• Size: single band around ~558 bp on a 2% agarose gel (or a clean FA/BA trace).
• If you see a strong ~150–200 bp dimer: repeat cleanup at 0.8× beads (selects ≥~300 bp).

PCR‑2: Indexing PCR (add P5/P7 and i5/i7)

Goal: Add the remaining adapter + indexes to yield a ~627‑bp library.

Reaction (50 µL per sample)

• 1× LongAmp buffer
• dNTPs 200 µM each
• i5/P5 indexing primer 0.5 µM
• i7/P7 indexing primer 0.5 µM
• 5 µL PCR‑1 product (unquantified is fine)
• LongAmp Taq per NEB’s unit guidance (typ. ~1–2 U / 50 µL)

Cycling (starting point):

• 95 °C 30 s
• 8 cycles of 95 °C 10 s / 60 °C 20 s / 65 °C 30 s
• 65 °C 2 min, 4 °C hold

Handling tips to minimize index‑primer dimers (since LongAmp is not hot‑start):

• Assemble on ice, add enzyme last, and load into a pre‑heated (95 °C) cycler if possible.
• Keep cycles at 6–8; don’t exceed 10 unless needed.
• If you ever see a dimer peak (~150–200 bp), do a 0.7–0.8× SPRI cleanup.

Cleanup (0.8× SPRI recommended):

• Add 40 µL beads to 50 µL PCR; proceed as in previous cleanup step, elute in 25–30 µL EB.

QC:

• Qubit libraries typically 5–30 ng/µL.
• Fragment length: single peak ~620–635 bp on FA/BA; minimal dimer (<5% area at ~150–200 bp).

Normalize, pool, and calculate molarity

Convert ng/µL → nM

$$ \text{nM} = \frac{\text{ng/µL} \times 10^6}{660 \times \text{bp}} $$

For 627 bp libraries: nM ≈ (ng/µL × 10^6) / (660 × 627) = ng/µL × 2.4165

Example: 10 ng/µL → ~24.2 nM.

1. Normalize each library to a common molarity (e.g., 4 nM).
2. Pool equal molar amounts of each sample.