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Bisulfite amplicon sequencing library preparation (using TruSeq adapters)

A **complete SOP** for generating **MiSeq‑ready bisulfite amplicon libraries** with **TruSeq‑tailed primers**. It assumes you already have **bisulfite‑treated genomic DNA** prepared and QC’d. EpiMark Hot Start Taq is formulated for **bisulfite‑converted, U‑containing, AT‑rich templates**. Family‑A Taq does **not stall on uracil**, and the **hot‑start** formulation suppresses primer‑dimers and off‑targets while you set up the reaction—important for long 5′‑tailed primers. **LongAmp Taq** (Taq + a proofreading partner, in an optimized buffer) is perfectly suitable for adding the Illumina adapters and indexes.

sample_prep
Version History
Version 1 Current
Effective: 2025-09-03

First version.

Procedure Steps (Version 1)

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.