SPRI Bead-based DNA Cleanup

Adapted for CCG use from NEBNext Ultra II DNA Library and QuantaBio manufacturer protocols.

Materials/Reagents

  • SparQ PureMag SPRI beads (aliquot, at room temperature)
  • Magnetic bead separation rack (suitable for tube/plate size)
  • 100% ethanol (EtOH), molecular-grade (~350 µL / sample)
  • Water, molecular-grade (and 0.1X TE buffer, if eluting/storing in TE)
  • Falcon tube or similar container for ethanol wash (minimum volume ~400 µL / sample)
  • Plastic trough (for ethanol wash, if using a multi-channel pipette)
  • PCR tubes (if larger tubes are used, ensure wash volume is sufficient to cover pelletized beads)

Selecting your Bead Ratio

Lower bead:sample ratios select for longer DNA fragments. While this can be useful, it can also cause total loss of some samples especially when working with archival DNA. Consult CCG staff before selecting a bead:sample ratio.

More on Selecting your Bead Ratio

Selecting the appropriate bead clean ratio is essential for your project. Below is an image of a ladder cleaned with different ratios. Notice how different ratios affect the retension of smaller fragments.

SPRI Bead Clean Ratio Gel

Bead ratios also affect your DNA yield. Below is a TapeStation overlay from various bead clean ratios. Notice how smaller ratios reduce the total yield even while retaining larger fragments.

SPRI Bead Clean Ratio TapeStation

This image comes from the Beckman Coulter SPRIselect Bead Interactive Protocol for Size Selection. This a helpful resource, but please consult with CCG Staff before approaching bead cleans for the first time.

Protocol

  1. Prepare a fresh aliquot of 80% EtOH (400 µL / sample) using molecular-grade water. If using a multi-channel pipette, dispense into a plastic trough immediately prior to wash steps.
  2. Bring beads to room temp. and vortex thoroughly. Re-vortex periodically if settling is present.
  3. Spin down samples and add bead solution. Mix thoroughly by pipetting, or vortex and briefly spin.

It’s important at this step to pipette measure your sample and calculate your bead volume according to your sample volume. Evaporation and pipetting error are common, and may affect your desired bead ratio.

  1. Incubate at least 5 min at room temperature (off magnet).
  2. Place sample tube(s) on magnet to pelletize beads and bound DNA. Keep on magnet until elution.
  3. Incubate at room temperature until beads are pelletized and supernatant is clear (~5 min).
  4. Remove and discard supernatant, taking care not to disturb the beads and bound DNA.
  5. Add 200 μL 80% ethanol wash. Incubate 30 sec. at room temperature. Remove and discard wash.
  6. Repeat wash step once for a total of two washes. Remove all visible liquid. If necessary, briefly spin down, return to magnet, and remove remaining ethanol with a small (e.g., p10) pipette.

  7. Air-dry beads for 1-2 minutes at room temperature, on the magnet with lid open.

    Do not over-dry the beads! Beads are sufficiently dry when pellets are dark and glossy, but all visible liquid has evaporated. If pellets change to light brown or start to crack, they are too dry.

  8. Remove sample tubes from magnet and resuspend beads in molecular-grade water or TE buffer. Pipette thoroughly to mix, or vortex and spin.

    Some buffer is often lost in the bead pellet during elution and transfer. Consider eluting in a slightly larger volume (appx. +2 µL) than what will be transferred and retained in Step 13 (and/or measuring the final volume by pipetting). Example: elute in 12 µL of buffer, but transfer only 10 µL.

  9. Incubate at least 2 minutes at room temperature (off magnet) to allow DNA to elute from beads.
  10. Return samples to magnet and incubate at room temperature until solution is clear (~5 min).
  11. Transfer samples to new tubes and discard used tubes with beads. Samples can be stored at -20 °C.

GHS Flammable Properly dispose of ethanol in the designated ethanol waste containers. Ethanol is a flammable chemical and cannot be disposed of in the trash or down the sink.