PBS recipe

1X Phosphate Buffered Saline (PBS Buffer) Recipe

Dissolve in 800ml distilled H2O:

8g of NaCl
0.2g of KCl
1.44g of Na2HPO4
0.24g of KH2PO4

Adjust pH to 7.4 with HCl.

Adjust volume to 1L with distilled H2O.

Sterilize by autoclaving.

Symbiodinium cp23S Re-PCR

Yesterday I completed some re-do PCRs of Symbiodinium cp23S from the branching Porites samples Sanoosh worked on over the past summer. Some of the samples did not amplify at all, so I reattempted PCR of these samples (107, 108, 112, 116). Sample 105 amplified last summer but the sequence was lousy, so I redid that one too. After the first PCR, I obtained 1 ul of the product and diluted it 1:100 in water. I then used 1 ul of this diluted product as the template for a second round of PCR. PCR conditions were the same Sanoosh and I used last summer (based on Santos et al. 2002):

Reagent Volume (µl)
water 17.2
5X Green Buffer 2.5
MgCl2 25 mM 2.5
dNTP mix 10 mM 0.6
Go Taq 5U/µl 0.2
primer 23S1 10 µM 0.5
primer 23S2 10 µM 0.5
Master Mix volume 24
sample 1
total volume 25

Initial denaturing period of 1 min at 95 °C, 35 cycles of 95 °C for 45 s, 55 °C for 45 s, and 72 °C for 1 min, and a final extension period of 7 min.

Samples were then run on a 1% agarose gel for 30 min at 135 volts.


Surprisingly, the first round of PCR amplified samples 107 and 112 (note: two subsamples of each were run; one that was the original extraction diluted (d) and another that was the original cleaned with Zymo OneStep PCR Inhibitor Removal kit (c)). The cleaned samples were the ones that amplified. I believe Sanoosh had tried these cleaned samples with no success.

The second round of PCR produced faint bands for both of the 108 samples. Sample 116 still did not amplify.

I cleaned the samples with the NEB Monarch Kit and shipped them today to Sequetech. I combined the two 108 samples to ensure enough DNA for sequencing.

RAD library prep

This is a belated post for some RAD library prep I did the week of January 23rd in the Leache Lab. I followed the same ddRAD/EpiRAD protocol I used in August. Samples included mostly Porites astreoides from the transplant experiment, as well as some geoduck samples from the OA experiment, and a handful of green and brown Anthopleura elegantissima. Sample metadata can be found here. The library prep sheet is here. The TapeStation report is here. Below is the gel image from the TapeStation report showing that the size selection was successful. However, the selection produced fragments with a mean size of 519-550 base pairs, as opposed to the size selection in August which produced ~500 bp fragments. While there will obviously be some overlap between libraries, combining samples from the two libraries may be problematic. This occurred despite identical Pippen Prep settings targeting fragments 415-515 bp. Libraries were submitted to UC Berkeley on 1/31/17 for 100 bp paired-end sequencing on the HiSeq 4000.

Library JD002_A-L


DNA extraction: Anthopleura elegantissima

In the interest of comparing methylation levels between symbiotic states in Anthopleura elegantissima, I extracted DNA from three zooxanthellate and three zoochlorellate individuals. These were anemones that were collected last summer at Point Lawrence, Orcas Island, and had been residing in indoor sea tables at Shannon Point since then. For each specimen, I excised part of the tentacle crown with scissors and deposited the tissue directly into a microfuge tube. I opted to freeze the tissue in the -80ºC freezer since earlier attempts with fresh tissue did not seem as effective (i.e., the tissue seemed resistant to lysis). After a day or two in the freezer, I pulled the samples out, rinsed them with PBS, and proceeded with the Qiagen DNeasy assay.  After addition of proteinase K, I used a small pestle to homogenize the sample. An overnight lysis period at 56ºC was used. DNA was eluted via two passes with 50 µl AE buffer (100 µl total volume). To further clean the DNA, samples were subject to ethanol precipitation using this protocol. Samples were re-eluted in 50 µl AE buffer.

To assess DNA quantity and quality, samples were tested with the Qubit BR DNA assay followed by electrophoresis on a 1% agarose gel with 1X TBE, 135 volts for 25 min.

sample ng/µl ng/µl post-dilution (100 µl buffer) total DNA
Ae-B-1 256 128 12800
Ae-B-2 183 91.5 9150
Ae-B-3 304 152 15200
Ae-G-1 163 81.5 8150
Ae-G-2 166 83 8300
Ae-G-3 274 137 13700


DNA extraction – naturally bleached Porites spp.

I was able to collect some naturally bleached Porites astreoides and Porites porites specimens while in Belize in November. I hope to use these as reference samples without symbiont DNA for RADseq. These samples were flash frozen instead of preserving in SS-DMSO. To extract DNA, a small fragment was crushed with a mortar and pestle and divided between three 1.5 ml tubes for extraction using the Qiagen DNeasy assay.  An overnight proteinase K lysis period at 56ºC was used. DNA was eluted via two passes with 50 µl AE buffer (100 µl total volume). To further clean the DNA, samples were subject to ethanol precipitation using this protocol. Samples were re-eluted in 100-200 µl AE buffer.

To assess DNA quantity and quality, samples were tested with the Qubit BR DNA assay followed by electrophoresis on a 1% agarose gel with 1X TBE, 125 volts for 30 min.

sample ng/ul (qubit) total vol total DNA ng
Past_bleached_1 15.8 100 1580
Past_bleached_2 7.02 100 702
Past_bleached_3 6.32 100 632
Ppor_bleached_1 16.7 100 1670
Ppor_bleached_2 6.74 200 1348
Ppor_bleached_3 13.7 100 1370


DNA looks great but note that RNase was not used and there may be some RNA in there. Also, note that the pellet was brown and there may be other contaminants.


DNA extraction – transplant expt.

This week I extracted DNA from Porites astreoides samples collected the week of 11/8/16 in Belize. These samples represent the endpoint of a year long common garden experiment in the backreef at Carrie Bow Cay. Detailed sample info can be found in the initial post here. The samples were stored at room temperature in salt saturated DMSO. Tissue was scraped off the skeletons with forceps into 1.5 ml tubes. Tissue was washed three times with PBS via centrifugation before beginning the Qiagen DNeasy extraction method. An overnight proteinase K lysis period at 56ºC was used. DNA was eluted via two passes with 50 µl AE buffer (100 µl total volume). To further clean the DNA, samples were subject to ethanol precipitation using this protocol. Samples were re-eluted in 50 µl AE buffer.

To assess DNA quantity and quality, samples were tested with the Qubit BR DNA assay followed by electrophoresis of the majority of them on a 1% agarose gel with 1X TBE, 135 volts for 30 min.

Sample DNA ng/µl
19 37.6
8 302
17 256
11h 304
7 180
18 326
16 242
2 151
11 178
10 306
12 163
4h 191
5 66.8
15 50.6
94 167
14 66.8
9 91.6
3 382
1 92.2
13h 197
5h 98.4
7h 184
6 195


GOALS for November

November will be an important month. Here are my goals:

1. Prep for Belize trip

  1. Prepare DMSO and RNA later in 50 ml tubes
  2. Order supplies: decitabine, magnetic stirrers, heaters, etc
  3. Anything else??? Bring waterpik, hemacytometer?

2. Belize trip goals

  1. Collect transplanted P. astreoides, save separate samples for DNA and morphology
  2. Run short term (3-7 days) gene expression experiment in backreef
  3. Run two week heat stress experiment in aquaria
  4. Possibly collect more P. porites samples
  5. Consider other side projects

3. Continue analysis of RADseq data

  1. Things to consider: allow missing data? test for outlier loci? how to deal with EpiRAD data?
  2. Goal to have mostly complete analysis by the end of the month (also end of class)

RADseq day 5

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.

RADseq days 3&4


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


Barcode# Bases



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

IMG_20160804_123903438 (1)

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


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

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:


5uL CutSmart buffer x n

1uL PstI x n

1uL MspI x n


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