RAD sequencing days 1&2

Yesterday was day one of my first ddRAD/EpiRAD run, which also serves as a training session. The first step was digestion of the DNA (double digest) with restriction enzymes. I chose the combination PstI/MspI for the ddRAD libraries and PstI/HpaII for the EpiRAD libraries. The cut sites for the MspI and HpaII are identical, except that HpaII is methylation sensitive. The following protocol was adapted by the protocol given to me by Kevin Epperly, the ddRAD guru.

Day 1: double digest

Here the two restriction enzymes shear the DNA at specific restriction sites so barcoded adapters can be attached, followed by indexes once fragments of the correct size have been selected.

1. DNA was diluted in advance with Qiagen AE buffer to reach a volume of 43 uL at the desired concentration. The targeted amount of DNA was 500 ng but some samples had higher and lower quantities (suggested min is 200 ng, max 750 ng). Samples were arranged in a 96 well plate in advance.

2. Four samples (#101-104) were selected as pre- and post-digestion controls to see if the digestion worked. Before starting the digest, 1 ul oL of each of these samples was set aside in labeled PCR tubes with 4 uL loading dye.

3. A master mix of the following was made:

ddRAD:

5uL CutSmart buffer x n

1uL PstI x n

1uL MspI x n

EpiRAD:

5uL CutSmart buffer x n

1uL PstI x n

1uL HpaII x n

n = number of samples plus slop. (ex. 48 samples plus 3 for slop would be 255ul CutSmart buffer + 51uL Sbf1 + 51uL Msp1 for a total master mix of 357uL) We used a spreadsheet to calculate the master mix.

4. 7uL of the master mix was added to the 43uL of DNA.

5. Samples were placed in thermocycler using the protocol “double digest” in Kevin’s folder: 37° C for 5 hours, hold

at 4° C

 

Day 2

Check pre- and post- digest samples to make sure digest was successful

1. Kevin made up a gel for us to run the pre- and post- digest controls

2. A post-digest 1 uL sample was taken from samples 101-104 and put into PCR tubes with 4 uL loading dye

3. Samples were loaded onto the gel, pre- and post- digest together, with a space in between each set. A ladder (1 uL) was placed in the first well.

4. The gel was run for 30-45 min at 121 volts.

The gel image below shows that the digest was successful: pre-digest samples are comprised of high molecular weight DNA, while post-digest samples show smearing indicative of the shearing of the enzymes (note: 101 pre-digest is very faint probably due to a pipetting error).

IMG_20160802_102032133

 

Bead cleanup following double digest

20 min before starting

1. Beads take out to warm to room temperature, kept in dark

2. Fresh 75% EtOH prepared (37.5 mL EtOH, filled up to 50 mL with MilliQ water in 50 ml conical tube)

When ready to start

1. Beads mixed thoroughly and poured into trough for multi-channel pipettor

2. Ratio is 1:1.5 sample:beads; sample volume was 50 uL, so used 75 uL beads

3. Used 12-channel pipettor and brand new box of tips (helps keep place)

4. Bead solution is very viscous; pipet slowly and change tips halfway through

IMG_20160802_090616431_HDR

5. 75 uL beads pipetted into special round-bottom plate for use with magnet

6. Pippettor set to 55 uL and samples pipetted from plate to round bottom bead plate

7. Sample and beads mixed by pipetting up and down several times

8. Put plate in dark for 5 min

9. PLate set on magnetic stand; beads should adhere to sides of wells and solution should clear

IMG_20160802_124154721_HDR

10. Solution removed being careful not to disturb beads (pipettor set to 135 uL)

11. 75% ethanol poured into trough

12. 195 uL ethanol added to wells

13. Samples should sit in ethanol for at least 30 sec

14. Pipettor turned up to 204 uL and ethanol removed slowly

15. Steps 12-14 repeated for second wash

16. At this point it was critical to make sure all ethanol gone, but beads should not dry out completely – watch them

17. When beads looked dry enough, started to add MilliQ water, 40 uL per well; add quickly, mix later

18. Come back w/ second set of tips to mix samples; may need to do third time. Avoid bead clumps

19. Let sit for at least 5 min off stand; DNA is now off beads in water

20. Label a new 96 well plate

21. Put plate back on magnetic stand and let beads adhere to sides

21. Transfer samples to new plate (40 uL), being careful to avoid beads

 

Quantify DNA w/ Qubit 

1. Important that all samples are quantified so that they can be pooled together in equal amounts. Adjustments may need to be made.

2. Pools should be comprised of samples with similar DNA conc.

3. Dilution may be necessary; I had to dilute 2 pools worth of samples down to 15 ng/uL.

4. Ideal concentration is 10 – 15 ng/uL. 20 is too much

5. Use 2 uL of sample with Qubit HS assay; may need BR assay for some samples if too high

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