After getting satisfactory Qubit results post-PCR, my final step in the ddRAD/EpiRAD library prep was to run the samples on the Agilent TapeStation to make sure fragments of correct and consistent size were selected earlier by the Pippin Prep.

1. Spin down samples and vortex

2. Obtain High Sensitivity D1000 Screentape and reagents. Spin down and vortex reagents. Note that ladder is in an extremely small volume.

3. Obtain 13 strip tubes (12 plus one for ladder) and place on a tube rack

4. Label 1st tube “L” for ladder, then 1-12 for each pool (only need to indicate order, not every tube)

5. Pipet 2 uL buffer into each tube

6. Pipet 2 uL ladder into “L” tube

7. Pipet 2 uL sample into each respective tube

8. Place caps on tubes, spin down, and vortex

9. Remove caps from tubes **Important – do not run TapeStation with lids on tubes

10. Place tubes in instrument, making sure ladder is in well position #1 (top left)

11. Place High Sensitivity D1000 Screentape in instrument with label facing towards the front of the instrument and the barcode facing right.

12. Ensure there are new pipet tips in machine and no used ones remaining

13. Open Agilent Tapestation Controller software

14. Select the tubes that will be used on the diagram

15. Rename the samples. These need to be the final names that will be sent to the sequencing facility. For example, my run was named JD001, and each pool was named A-L, such as JD001_A, JD001_B, and so on.

16. Press “START”

17. Save to JayDimond folder on desktop

18. When run is done, save output file as a MS Word document by creating a report under the file menu; click option to create thumbnail of each sample. This needs to be saved using the final plate name sending to the sequencing facility. For example, my file was named 2016-08-08_JD001_A-L.docx. If the run contains only part of a plate or parts of different plates name accordingly.

My results looked good, showing consistent size selection and quantities that matched well with the prior Qubit results. The fragment sizes were a bit larger than they should have been, averaging about 495 bp instead of 465 bp. If the resulting fragments are to be compared to fragments generated in another set of libraries, it is important that the fragment sizes are as consistent as possible, because the loci chosen will vary depending on their size. Thus it is particularly essential that the Pippin Prep settings are the same.

These libraries will be sent to UC Berkeley within the next two weeks or so along with Greg’s frog libraries. Once there, the facility will perform qPCR to ensure the libraries are combined in equimolar ratios in a single lane.

Library metadata:

Prep No.	Conc. post digestion (ng/ul)	New Pool (column)	Well	enzymes	Barcode	Illumina Index	Post Size Selection ng/ul	Post PCR ng/ul	Mean fragment size (bp)	Library name
past_80	15.00	Pool 1	A	PstI-HpaII	TGCAT	ATCACG
past_80	15.00	Pool 1	B	PstI-MspI	AAGGA	ATCACG
116	15.00	Pool 1	C	PstI-HpaII	GCATG	ATCACG
108	15.00	Pool 1	D	PstI-HpaII	AACCA	ATCACG	0.264	1.93	502	JD001_A
past_98	15.00	Pool 1	E	PstI-MspI	CAACC	ATCACG
past_82	15.00	Pool 1	F	PstI-HpaII	TCGAT	ATCACG
116	15.00	Pool 1	G	PstI-MspI	CGATC	ATCACG
102	15.00	Pool 1	H	PstI-HpaII	GGTTG	ATCACG
past_84	15.00	Pool 2	A	PstI-MspI	TGCAT	CGATGT
past_w1	15.00	Pool 2	B	PstI-MspI	AAGGA	CGATGT
past_90	15.00	Pool 2	C	PstI-MspI	GCATG	CGATGT
past_w1	15.00	Pool 2	D	PstI-HpaII	AACCA	CGATGT	0.39	10.10	493	JD001_B
past_91	15.00	Pool 2	E	PstI-HpaII	CAACC	CGATGT
past_88b	15.00	Pool 2	F	PstI-MspI	TCGAT	CGATGT
past_90	15.00	Pool 2	G	PstI-HpaII	CGATC	CGATGT
past_89	15.00	Pool 2	H	PstI-HpaII	GGTTG	CGATGT
past_98	18.57	Pool 3	A	PstI-HpaII	TGCAT	TTAGGC
127	18.06	Pool 3	B	PstI-HpaII	AAGGA	TTAGGC
102	17.78	Pool 3	C	PstI-MspI	GCATG	TTAGGC
past_84	17.02	Pool 3	D	PstI-HpaII	AACCA	TTAGGC	0.411	5.18	498	JD001_C
past_w3	16.96	Pool 3	E	PstI-HpaII	CAACC	TTAGGC
past_86	16.83	Pool 3	F	PstI-MspI	TCGAT	TTAGGC
103	16.22	Pool 3	G	PstI-HpaII	CGATC	TTAGGC
past_w3	16.15	Pool 3	H	PstI-MspI	GGTTG	TTAGGC
106	15.90	Pool 4	A	PstI-HpaII	TGCAT	TGACCA
114	15.31	Pool 4	B	PstI-HpaII	AAGGA	TGACCA
past_91	15.28	Pool 4	C	PstI-MspI	GCATG	TGACCA
past_85	15.22	Pool 4	D	PstI-HpaII	AACCA	TGACCA	0.411	5.41	493	JD001_D
131	14.51	Pool 4	E	PstI-HpaII	CAACC	TGACCA
115	14.24	Pool 4	F	PstI-HpaII	TCGAT	TGACCA
past_95	14.10	Pool 4	G	PstI-MspI	CGATC	TGACCA
past_86	13.98	Pool 4	H	PstI-HpaII	GGTTG	TGACCA
128	13.94	Pool 5	A	PstI-MspI	TGCAT	ACAGTG
121	13.78	Pool 5	B	PstI-MspI	AAGGA	ACAGTG
past_87	13.73	Pool 5	C	PstI-HpaII	GCATG	ACAGTG
111	13.54	Pool 5	D	PstI-HpaII	AACCA	ACAGTG	0.35	5.29	494	JD001_E
129	13.39	Pool 5	E	PstI-MspI	CAACC	ACAGTG
120	13.37	Pool 5	F	PstI-HpaII	TCGAT	ACAGTG
123	13.37	Pool 5	G	PstI-HpaII	CGATC	ACAGTG
118	13.06	Pool 5	H	PstI-HpaII	GGTTG	ACAGTG
128	12.92	Pool 6	A	PstI-HpaII	TGCAT	GCCAAT
131	12.83	Pool 6	B	PstI-MspI	AAGGA	GCCAAT
past_87	12.71	Pool 6	C	PstI-MspI	GCATG	GCCAAT
112	12.61	Pool 6	D	PstI-MspI	AACCA	GCCAAT	0.321	8.44	496	JD001_F
122	12.56	Pool 6	E	PstI-MspI	CAACC	GCCAAT
101	12.50	Pool 6	F	PstI-MspI	TCGAT	GCCAAT
122	12.50	Pool 6	G	PstI-HpaII	CGATC	GCCAAT
105	12.29	Pool 6	H	PstI-HpaII	GGTTG	GCCAAT
101	12.22	Pool 7	A	PstI-HpaII	TGCAT	CAGATC
past_85	12.19	Pool 7	B	PstI-MspI	AAGGA	CAGATC
107	12.17	Pool 7	C	PstI-MspI	GCATG	CAGATC
past_89	12.12	Pool 7	D	PstI-MspI	AACCA	CAGATC	0.372	2.71	491	JD001_G
past_95	11.68	Pool 7	E	PstI-HpaII	CAACC	CAGATC
125	11.56	Pool 7	F	PstI-HpaII	TCGAT	CAGATC
105	11.50	Pool 7	G	PstI-MspI	CGATC	CAGATC
past_96	11.43	Pool 7	H	PstI-MspI	GGTTG	CAGATC
121	11.39	Pool 8	A	PstI-HpaII	TGCAT	ACTTGA
past_88b	11.18	Pool 8	B	PstI-HpaII	AAGGA	ACTTGA
123	11.11	Pool 8	C	PstI-MspI	GCATG	ACTTGA
126	11.11	Pool 8	D	PstI-MspI	AACCA	ACTTGA	0.302	5.04	500	JD001_H
120	10.83	Pool 8	E	PstI-MspI	CAACC	ACTTGA
117	10.78	Pool 8	F	PstI-MspI	TCGAT	ACTTGA
115	10.72	Pool 8	G	PstI-MspI	CGATC	ACTTGA
past_96	10.68	Pool 8	H	PstI-HpaII	GGTTG	ACTTGA
130	10.61	Pool 9	A	PstI-MspI	TGCAT	GATCAG
111	10.56	Pool 9	B	PstI-MspI	AAGGA	GATCAG
past_w11	10.50	Pool 9	C	PstI-MspI	GCATG	GATCAG
125	10.44	Pool 9	D	PstI-MspI	AACCA	GATCAG	0.42	3.57	503	JD001_I
112	10.35	Pool 9	E	PstI-HpaII	CAACC	GATCAG
110	10.28	Pool 9	F	PstI-HpaII	TCGAT	GATCAG
past_w11	10.25	Pool 9	G	PstI-HpaII	CGATC	GATCAG
103	10.17	Pool 9	H	PstI-MspI	GGTTG	GATCAG
117	9.97	Pool 10	A	PstI-HpaII	TGCAT	TAGCTT
127	9.72	Pool 10	B	PstI-MspI	AAGGA	TAGCTT
130	9.69	Pool 10	C	PstI-HpaII	GCATG	TAGCTT
106	9.61	Pool 10	D	PstI-MspI	AACCA	TAGCTT	0.358	3.38	498	JD001_J
124	9.51	Pool 10	E	PstI-HpaII	CAACC	TAGCTT
114	9.06	Pool 10	F	PstI-MspI	TCGAT	TAGCTT
109	8.89	Pool 10	G	PstI-MspI	CGATC	TAGCTT
109	8.85	Pool 10	H	PstI-HpaII	GGTTG	TAGCTT
126	8.85	Pool 11	A	PstI-HpaII	TGCAT	GGCTAC
124	8.78	Pool 11	B	PstI-MspI	AAGGA	GGCTAC
110	8.33	Pool 11	C	PstI-MspI	GCATG	GGCTAC
107	8.30	Pool 11	D	PstI-HpaII	AACCA	GGCTAC	0.236	2.15	501	JD001_K
129	7.99	Pool 11	E	PstI-HpaII	CAACC	GGCTAC
past_99	7.58	Pool 11	F	PstI-MspI	TCGAT	GGCTAC
past_99	7.58	Pool 11	G	PstI-HpaII	CGATC	GGCTAC
108	6.94	Pool 11	H	PstI-MspI	GGTTG	GGCTAC
118	6.56	Pool 12	A	PstI-MspI	TGCAT	CTTGTA
104	6.06	Pool 12	B	PstI-MspI	AAGGA	CTTGTA
past_81	5.78	Pool 12	C	PstI-HpaII	GCATG	CTTGTA
past_81	4.97	Pool 12	D	PstI-MspI	AACCA	CTTGTA	0.201	2.56	493	JD001_L
113	4.83	Pool 12	E	PstI-HpaII	CAACC	CTTGTA
104	4.34	Pool 12	F	PstI-HpaII	TCGAT	CTTGTA
113	1.33	Pool 12	G	PstI-MspI	CGATC	CTTGTA
past_82	0.03	Pool 12	H	PstI-MspI	GGTTG	CTTGTA

Note that two samples were compromised due to pipetting errors: sample 102, digested by PstI/MspI, pool 3, well C; and sample 120, digested by PstI/MspI, pool 8, well E. These samples were completely excluded early in the process and no sequences should be obtained from them.

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.

1. Use spreadsheet to calculate master mix volumes
2. Start up thermocycler – choose Kevin -> Ligation. Start program, then pause until ready
3. Get out reagents and put on ice
   1. T4 Ligase
   2. T4 buffer
   3. Barcodes (8)
   4. P2 adapter
   5. Annealing buffer
4. Obtain chilled plate/tube holder and two strip tubes (would only need one for a single master mix, but I made two)
5. Label strip tubes (mostly for correct orientation (1-8)
6. Add annealing buffer to each tube
7. Add different barcode to each tube
8. Mix P2 master mix in 1.5 mL tube (2 separate ones in my case)
9. Mix T4 master mix in 1.5 mL tube (2””)
10. Reagents are kept on ice at all times; keep in mind they will be viscous when pipetting
11. Add P2 and T4 mixes together and mix well
12. Divide combined volume by 8.2 to figure out quantity to add to each tube of barcodes
13. Add T4+P2 to the adapters, mix well by pipetting, spin down, and vortex
14. Now add 7 uL master mix to samples in plate using 8-channel pipettor, mix by pipetting up and down several times
15. Spin down and vortex plate
16. 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

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).

 

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

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

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