8/3/2016
The morning was spent re-Qubiting some samples that were out of range of the HS assay. Kevin also spent some time checking a few samples of mine and Greg’s since our readings for the control (standard #2) were higher than they should have been. It appears that we were making some pipetting errors. We ended up correcting our sample concentrations based on what the control reading should have been (i.e., determining what proportion of the “true” value our control values came out to, and using that as a correction factor). It is important to have a good idea of each sample’s concentration so you can pool them together in roughly equal proportions. My samples were completely reordered in a new 96 well plate in order to create pools of samples with similar concentrations. The spreadsheet can be used to help with reordering. Note that this must be done very carefully; I accidentally pipetted sample 1B to sample 3C of the original plate. Thus samples 102 and 120 were compromised and removed from further analysis. The next step was the ligation. Before I forget, here are the keys for the barcodes and indices:
| Index# | Bases |
| 1 | ATCACG |
| 2 | CGATGT |
| 3 | TTAGGC |
| 4 | TGACCA |
| 5 | ACAGTG |
| 6 | GCCAAT |
| 7 | CAGATC |
| 8 | ACTTGA |
| 9 | GATCAG |
| 10 | TAGCTT |
| 11 | GGCTAC |
| 12 | CTTGTA |
| Barcode# | Bases |
| 1 | TGCAT |
| 2 | AAGGA |
| 3 | GCATG |
| 4 | AACCA |
| 5 | CAACC |
| 6 | TCGAT |
| 7 | CGATC |
| 8 | GGTTG |
Ligation
Using a spreadsheet from Kevin, we calculated master mix volumes for the ligation. In this step, barcodes (aka P1, adapters) are ligated on to the DNA fragments, as well as P2 adapters for Illumina sequencing. There are 8 different barcodes used to label the 8 rows of a 96 well plate. If you end up with a wide range of sample concentrations, it is ideal to use two sets of master mixes so you do not use up too much reagent unnecessarily. This is what I ended up doing.
- Use spreadsheet to calculate master mix volumes
- Start up thermocycler – choose Kevin -> Ligation. Start program, then pause until ready
- Get out reagents and put on ice
- T4 Ligase
- T4 buffer
- Barcodes (8)
- P2 adapter
- Annealing buffer
- Obtain chilled plate/tube holder and two strip tubes (would only need one for a single master mix, but I made two)
- Label strip tubes (mostly for correct orientation (1-8)
- Add annealing buffer to each tube
- Add different barcode to each tube
- Mix P2 master mix in 1.5 mL tube (2 separate ones in my case)
- Mix T4 master mix in 1.5 mL tube (2””)
- Reagents are kept on ice at all times; keep in mind they will be viscous when pipetting
- Add P2 and T4 mixes together and mix well
- Divide combined volume by 8.2 to figure out quantity to add to each tube of barcodes
- Add T4+P2 to the adapters, mix well by pipetting, spin down, and vortex
- Now add 7 uL master mix to samples in plate using 8-channel pipettor, mix by pipetting up and down several times
- Spin down and vortex plate
- Put in thermal cycler and press start on ligation protocol (takes ~1.5 hrs)
Bead cleanup
Materials needed: magnet rack, 1.5 mL tubes, 75% EtOH, milliq water, EB buffer, pipets, tips, sterile toothpicks
1. Label 3 sets of 12 1.5 mL tubes (labeled with pools 1-12). One of these sets will be saved – put initials on them in addition to numbers.
2. Pipet 75 uL beads into each tube of one of the sets of tubes.
3. Set pipet to 45 uL to make sure you withdraw full 40 uL sample
4. Now the barcoded samples are combined into 12 pools. Pipet 40 uL sample from each column of the 96-well plate into each of the 12 tubes (only need to change tips for each new column)
5. Add 380 uL bead solution to each tube and mix w/ pipet
6. Let sit for 5 min in dark
7. Place tubes on magnet rack & wait 5-10 min for beads to adhere and solution to clear
8. Remove supernatant (set pipet to ~420 uL), being careful to avoid beads
9. Add 800 uL 75% EtOH & let sit for 30 sec
10. Remove supernatant
11. Repeat 9-10
12. Wick any excess EtOH from side or bottom of tube with a sterile toothpick
13. When beads seem dry enough (be careful not to overdry) add 50 uL water, pipeting against beads to dislodge, and pipetting repeatedly up and down to mix well
14. Let tubes sit for 5 min in dark, out of magnetic rack
15. Put tubes back on magnetic rack and wait 5-10 min for solution to clear
16. Pipet off 50 uL sample and put into new tubes (note: we stopped here for the day, so put the sample in new, empty tube. But normally, if there is time, proceed directly to second wash and put sample into new tube with 75 uL fresh bead solution).
Second bead cleanup (day 4, 8/4/16)
17. If continuing directly from first cleanup, put 50 uL sample into new tube with 75 uL bead solution. If sample is already in new tube, add 75 uL bead solution.
18. Mix well w/ pipet and let sit for 5 min in dark
18. Place tubes on magnet rack & wait 5-10 min for beads to adhere and solution to clear
19. Remove supernatant (set pipet to ~140 uL), being careful to avoid beads
20. Add 195 uL 75% EtOH & let sit for 30 sec
21. Remove supernatant
22. Repeat 9-10
23. Wick any excess EtOH from side or bottom of tube with a sterile toothpick
24. When beads seem dry enough (be careful not to overdry) add 30 uL EB buffer, pipeting against beads to dislodge, and pipetting repeatedly up and down to mix well
25. Let tubes sit for 5 min in dark, out of magnetic rack
26. Put tubes back on magnetic rack and wait 5-10 min for solution to clear
27. Pipet off 30 uL sample and put into new tubes
Size selection with Pippin Prep
1. Turn on machine if it is not already on (rear power button)
2. Select “ddRAD-SEQ” protocol; there are several protocols, choose the one exactly as written
3. Go to protocol editor. Start and End should ready 415, 515 base pairs (a mean size of 465 bp will be selected)
4. Must calibrate the machine if it has not already been used that day or previous day and left on
5. Use calibration “fixture” kept in blue sock
6. Place over blue light spots, ensuring that dark spots/ wrinkles on fixture are not placed over light spots
7. Hit calibrate button
8. Close machine lid
9. Hit calibrate and make sure it is successful (repeat if not)
10. Name each pool, e.g. “Pool_1_JD”. Each cassette has 5 lanes and can do 5 pools.
11. Open a new cassette package
12. Tap bubbles out holding the cassette in the up position
13. Inspect the gels to make sure they are not broken or cracked, no gaps
14. Place the cassette on the chamber and push it all the way to the left (there is some wiggle room, want to always keep it on left side)
15. Peel back stickers and set them aside in case some of the cassette lanes will be used at another time
16. Fill up any wells that are low on buffer – should be able to see rounded meniscus, almost to top. Must fill both ends of cassette & ensure wells are covered.
17. For the elution well, you want identical volumes. Remove existing buffer carefully with pipet (go directly down to bottom) and replace with exactly 40 uL new buffer. Go all the way to bottom and come up slowly, being careful to not introduce air bubbles.
18. Take care avoiding contamination with nearby lanes.
19. Put new tape strip over elution wells, with white tab facing inside.
20. Close lid. Lid will resist slightly because probes are being inserted.
21. Hit test button. If it fails, open and close lid again, and retry (note: any used lanes will fail)
22. Now to first five samples that will be run (in 1.5 mL tubes), add 10 uL of R2 mix (standards). Note: spin down and vortex R2 before using.
23. R2 is viscous, must go slow and pipet up and down to mix.
24. Vortex tubes lightly and let sit for 2 min.
25. To load Pippin cassette, remove 40 uL buffer from wells and replace with 40 uL sample (just like loading any other gel)
26. Check buffer again
27. Press start – will ask if you calibrated – click yes
28. Will run for ~30 min. Should see standard peaks at 50 and 150 bp
29. Will stop on its own if all lanes used, but will continue to run if not all lanes used, so must stop manually in that case
30. When it is close to being done, can start prepping next cassette
31. When done, open door, carefully remove tape over elution wells, and withdraw 40 uL sample into new labeled 1.5 mL tube.
32. Can start prepping Qubit tubes while waiting for runs to finish
Qubit post-Pippin
1. Label 12 Qubit tubes for samples plus one control and two standard tubes
2. Proceed with Qubit HS protocol.
3. Use 2 uL S2 for control; should read ~10 ng/uL
| Pool | Qubit DNA ~40μl Post-Pippin (ng/μL) |
| 1 | 0.264 |
| 2 | 0.39 |
| 3 | 0.411 |
| 4 | 0.411 |
| 5 | 0.35 |
| 6 | 0.321 |
| 7 | 0.372 |
| 8 | 0.302 |
| 9 | 0.42 |
| 10 | 0.358 |
| 11 | 0.236 |
| 12 | 0.201 |
control = 9.58 ng/uL
PCR – Illumina Indexes
Here a unique index is added to each pool, so they can later be combined, samples are amplified through PCR increasing the total quantity of DNA.
1. Use the spreadsheet to calculate how much DNA and water to combine
to get 34.5 ul at the targeted concentration. Ideal target of total DNA is 10 ng,
but this can be lower (ideally no lower than 5 ng/uL) if you do not have enough DNA after size selection.
2. Take reagents out of fridge or freezer and put on ice bucket.
3. Keep the 12 unique PCR primers/adapters in order.
4. Label new set of 12 PCR strip tubes and place in cold plate
5. Add DNA sample plus any water needed to get ~10 ng total DNA in 34.5 uL total volume (use spreadsheet to calculate)
6. In a 1.5 mL tube make a master mix of 2 ul PCR Primer 1, 10 ul 5X Phusion HF Buffer, 1ul
DNTP and 0.5 HF Phusion Taq per sample plus slop.
7. Add 13.5 uL master mix to each tube.
8. Add 2 uL of the appropriate PCR Primer 2 to each tube (each one is unique). Total reaction volume is now 50 uL.
9. Spin down and vortex tubes
4. Run samples on thermocycler using the “Phusion” protocol (98°C for 30 sec,
98°C for 10 sec, 62°C for 30 sec, 72° for 30 sec, repeat step 2-4 10 more
times, 72°C for 10 min, hold at 4°C).
Post PCR Bead Cleanup
before starting
1. Label 2 sets of 1.5 mL tubes, one for the cleanup
and one for the final elution
2. Make fresh 75% EtOH
When ready to start
3. Mix beads thoroughly
4. Add 75 uL Serapure per tube to first set of 1.5 mL tubes
5. Add samples (50uL) to 1.5 ml tubes of Serapure; mix by pipetting
6. Incubate at room temp for 5 min in the dark
7. Place tubes on magnet stand for 5-10 min
8. Remove supernatant (125 uL)
9. Add 195 uL of 75% EtOH (do not remove from stand)
8. Incubate for 30 sec
9. Remove supernatant (205 uL)
10. Add 195 uL of 70% EtOH (do not remove from stand)
11. Incubate for 30 sec
12. Remove supernatant (205 uL)
13. Remove blobs of EtOH with sterile toothpick & make sure beads dry (but not too much)
14. Add 40 uL EB buffer; pipette to mix (remove from stand)
15. Incubate for 5 min in the dark; then place back on stand for 5-10 min
16. Pipette supernatant (40 uL) to second set of 1.5 ml tubes
Qubit
1. Qubit the samples using the HS protocol and record the values in spreadsheet in ng/uL.
Here are the data:
| SAMPLE | Qubit DNA ~40μl Post-PCR Enrich. (ng/μL) | Total DNA ~40μl Post-PCR Enrich. (ng) | ||||||||||
| POOL 1 | 1.93 | 77.2 | ||||||||||
| POOL 2 | 10.10 | 404.0 | ||||||||||
| POOL 3 | 5.18 | 207.2 | ||||||||||
| POOL 4 | 5.41 | 216.4 | ||||||||||
| POOL 5 | 5.29 | 211.6 | ||||||||||
| POOL 6 | 8.44 | 337.6 | ||||||||||
| POOL 7 | 2.71 | 108.4 | ||||||||||
| POOL 8 | 5.04 | 201.6 | ||||||||||
| POOL 9 | 3.57 | 142.8 | ||||||||||
| POOL 10 | 3.38 | 135.2 | ||||||||||
| POOL 11 | 2.15 | 86.0 | ||||||||||
| POOL 12 | 2.56 | 102.4 | ||||||||||
| TEST St. #2 | 9.20 | 368.0 | ||||||||||