LCM Pilot Zymo Trio WGBS Library Prep Round 3

LCM Zymo Trio WGBS Library Prep

Protocol link here

Samples: 12/15/24 Extraction of LCM Pilot Samples

Extracting LCM samples to use for WGBS. These are from the same slides as the RNA ones, but different section.

frag sample_id concentration (ng/uL) DIN
A 1 4.35 3.4
A 3 4.08 3.4
B 11 3.41 3
B 12 3.61 3
D 24 3.63 3.1
D 25 3 1.8
  • Eluted DNA in 20 uL Tris-HCl, used 2 uL for Qubit and 1 uL for gDNA tapestation

Full results can be found here (did not QC RNA)

Library Prep

  • 17 uL (brought up to 20 uL with 3uL Tris) processed using EZ Methylation Gold Kit for Bisulfite Conversion, cleaned according to protocol and eluted in 19 uL DNA elution buffer
  • Did 20 PCR cycles.

QC

Run to visualize libraries. Here’s an example of what the library should look like on a Tapestation:

“If adapter dimers are present, they will form an approximately 130-180 bp band. Yields will vary depending on the total quantity and quality of sample input DNA.

Results

YAY!!! Libraries! They have more dimer than the re-amped ones. Probably stopping the PCR and cleaning and then re-amping helps a lot with dimers. But that would use a Lot of master mix and primer for all the samples.

Full results can be found here

Second Bead cleanup to remove Dimers and fragments > 1000bp:

  • Double sided size selection protocol based on Zymo Magbead Select-a-Size clean and concentrate protocol.
  1. Add 32 uL DNA elution buffer (EB) to bring volume to 50 uL
  2. Bind larger fragments (> 1100 bp) to beads to remove from sample:
    1. 0.48X ratio: Add 24 uL beads
      1. total volume = 74 uL
    2. Mix thoroughly by pipetting until homogenous and incubate for 10 minutes at room temperature
    3. Place the tube on a magnetic stand for 3 minutes, or until the supernatant is clear.
    4. Move 74 uL of supernatant to new tube KEEP SUPERNATANT. Discard the beads.
  3. Add second volume of beads to supernatant tube to clean up all the fragments < 1000 bp
    1. 1.32X ratio = 1.32 * 50 uL = 66 uL
    2. Add 66 uL beads to the supernatant that was transferred into the new tube
    3. Mix thoroughly by pipetting until homogenous and incubate for 10 minutes at room temperature
    4. Place the tube on a magnetic stand for 3 minutes, or until the supernatant is clear.
    5. Carefully remove the supernatant without disturbing the magnetized bead pellet.
    6. Without removing from the magnetic stand, add 200 μL of DNA Wash Buffer to the tube, incubate for at least 30 seconds, and then remove the supernatant completely without disturbing the magnetized bead pellet. Repeat this wash step for two washes total.
    7. Remove the tube from the magnetic stand and centrifuge very briefly. Then return the tube to the magnetic stand, wait for the beads to pellet, and remove any residual DNA Wash Buffer with a 10 μL pipette tip.
    8. Dry on magnetic stand for 2-3 minutes with cap open, do not over-dry beads
    9. Elute in 50 uL DNA EB. Fully resuspend beads and incubate for 5 minutes at room temperature.
    10. Place the tube back on the magnetic stand for 2 minutes or until the supernatant is clear.
    11. Transfer the indicated volume of eluate to a new tube. Discard the beads.
  4. Deplete small fragments (< 200 bp) with another volume of beads
    1. 0.8X ratio = 0.8 * 50 uL = 40 uL beads
    2. Mix thoroughly by pipetting until homogenous and incubate for 10 minutes at room temperature
    3. Place the tube on a magnetic stand for 3 minutes, or until the supernatant is clear.
    4. Carefully remove the supernatant without disturbing the magnetized bead pellet.
    5. Without removing from the magnetic stand, add 200 μL of DNA Wash Buffer to the tube, incubate for at least 30 seconds, and then remove the supernatant completely without disturbing the magnetized bead pellet. Repeat this wash step for two washes total.
    6. Remove the tube from the magnetic stand and centrifuge very briefly. Then return the tube to the magnetic stand, wait for the beads to pellet, and remove any residual DNA Wash Buffer with a 10 μL pipette tip.
    7. Dry on magnetic stand for 2-3 minutes with cap open, do not over-dry beads
    8. Elute in 33 uL DNA EB. Fully resuspend beads and incubate for 5 minutes at room temperature.
    9. Place the tube back on the magnetic stand for 2 minutes or until the supernatant is clear.
    10. Transfer 32 uL of eluate to a new tube. Discard the beads.
  • 1 uL used for D5000 Tapestation
  • 31 left over to be re-cleaned up

Results

YAY! these look good but need a final clean-up to remove dimers.

Full results can be found here

Third bead cleanup to remove Dimers

  • Right sided size selection protocol based on Zymo Magbead Select-a-Size clean and concentrate protocol.
  1. Add 19 uL DNA elution buffer (EB) to bring volume to 50 uL
  2. Deplete small fragments (< 200 bp) with another volume of beads
    1. 0.8X ratio = 0.8 * 50 uL = 40 uL beads
    2. Mix thoroughly by pipetting until homogenous and incubate for 10 minutes at room temperature
    3. Place the tube on a magnetic stand for 3 minutes, or until the supernatant is clear.
    4. Carefully remove the supernatant without disturbing the magnetized bead pellet.
    5. Without removing from the magnetic stand, add 200 μL of DNA Wash Buffer to the tube, incubate for at least 30 seconds, and then remove the supernatant completely without disturbing the magnetized bead pellet. Repeat this wash step for two washes total.
    6. Remove the tube from the magnetic stand and centrifuge very briefly. Then return the tube to the magnetic stand, wait for the beads to pellet, and remove any residual DNA Wash Buffer with a 10 μL pipette tip.
    7. Dry on magnetic stand for 2-3 minutes with cap open, do not over-dry beads
    8. Elute in 33 uL DNA EB. Fully resuspend beads and incubate for 5 minutes at room temperature.
    9. Place the tube back on the magnetic stand for 2 minutes or until the supernatant is clear.
    10. Transfer 32 uL of eluate to a new tube. Discard the beads.
  • 1 uL used for D5000 Tapestation
  • 30 uL final volume for Sequencing, move to LoBind tube.

Results

Primer dimer is still >5% of the total library. I need to clean one more time, and with a lower ratio than 0.8X.

Full results can be found here and here (scaled)

Final, 0.7X Bead cleanup to remove Dimers

  • Right sided size selection protocol based on Zymo Magbead Select-a-Size clean and concentrate protocol.
  1. Add 19 uL DNA elution buffer (EB) to bring volume to 50 uL
  2. Deplete small fragments (< 200 bp) with another volume of beads
    1. 0.7X ratio = 0.7 * 50 uL = 35 uL beads
    2. Mix thoroughly by pipetting until homogenous and incubate for 10 minutes at room temperature
    3. Place the tube on a magnetic stand for 3 minutes, or until the supernatant is clear.
    4. Carefully remove the supernatant without disturbing the magnetized bead pellet.
    5. Without removing from the magnetic stand, add 200 μL of DNA Wash Buffer to the tube, incubate for at least 30 seconds, and then remove the supernatant completely without disturbing the magnetized bead pellet. Repeat this wash step for two washes total.
    6. Remove the tube from the magnetic stand and centrifuge very briefly. Then return the tube to the magnetic stand, wait for the beads to pellet, and remove any residual DNA Wash Buffer with a 10 μL pipette tip.
    7. Dry on magnetic stand for 2-3 minutes with cap open, do not over-dry beads
    8. Elute in 32 uL DNA EB. Fully resuspend beads and incubate for 5 minutes at room temperature.
    9. Place the tube back on the magnetic stand for 2 minutes or until the supernatant is clear.
    10. Transfer 31 uL of eluate to a new tube. Discard the beads.
  • 1 uL used for D5000 Tapestation
  • 30 uL final volume for Sequencing, move to LoBind tube.

Results

Done. Sent for sequencing (along with the 12/6 and 12/12 libraries) on 12/17/24.

Full results (these are the first 6 lanes, the other 4 libraries are the 12/6 and 12/12 libraries) can be found here and here (scaled)

Written on December 15, 2024