Category Archives: Puget Sound Oly Common Garden

Tuesday 9/22/15

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Lab Work

Set up a test PCR to determine the optimum # of cycles. You want to identify the minimum number of cycles required for a visible product at 166 bp. I chose 4 of the samples and tested them at 12, 17, 22, & 27 cycles as recommend in the protocol.

  • Made 100 uM stock of new ILL-Lib2 adaptor
  • Made 10 uM stocks of ILL-Lib1 and ILL-Lib2 primers
    • 1.5 stock + 13.5 uL NFW
  • Made 1 uM stocks of HT1 and BC1
    • 1uL stock + 99 NFW
1x
17x
NFW
4.6 uL
78.2
10 mM (each) dNTPS
.4
6.8
10 uM ILL-Lib1
.4
6.8
10 uM ILL-Lib2
.4
6.8
1 uM ILL-HT1
1
17
1 uM ILL-BC1
1
17
5X Q5 buffer
4
68
Q5 Taq polymerase
.2
3.4
Added 12 uL master mix to 8 uL ligation.
  1. SS2_3_12x
  2. HC1_1_12x
  3. NF1_1_12x
  4. NF1_2_12x
  5. SS2_3_17x
  6. HC1_1_17x
  7. NF1_1_17x
  8. NF1_2_17x
  9. SS2_3_22x
  10. HC1_1_22x
  11. NF1_1_22x
  12. NF1_2_22x
  13. SS2_3_27x
  14. HC1_1_27x
  15. NF1_1_27x
  16. NF1_2_27x

17 and 18 are 2 of the ligation reactions for comparison.

  • 17: SS_2_3
  • 18: NF_1_1
Programed PCR in thermocyclers 4,5,6. Called 2b12, 2b17, 2b22, and 2b27.
  •  (98°C 5 sec, 60°C 20 sec, 72°C 10 sec) X N cycles

9_22_15

It looks like 22 cycles is best, and worked on all samples so will be using that in the preparatory scale.

Ethanol precipitation

  • Set up an ethanol precipitation of all the broodstock samples I’ve extracted so far, as well as repeats of of the 4 samples used in the test scale PCR:
    • Population Sample Date extracted ng/uL Volume for 1 ug Volume sodium acetate Vol ethanol
      Oyster Bay BS_2_3 19.8 50.50505051 5.05 113.64
      Hood Canal BS_1_1 12.3 81.30081301 8.13 182.93
      Fidalgo BS_1_1 9.13 109.5290252 10.95 246.44
      Fidalgo BS_1_2 17.4 57.47126437 5.75 129.31
      Oyster Bay BS_2_6 23.4 42.73504274 4.27 96.15
      Oyster Bay BS_2_7 16.4 60.97560976 6.10 137.20
      Oyster Bay BS_2_8 16.8 59.52380952 5.95 133.93
      Hood Canal BS_1_5 12.1 82.6446281 8.26 185.95
      Hood Canal BS_1_6 23.5 42.55319149 4.26 95.74
      Hood Canal BS_1_7 14 71.42857143 7.14 160.71
      Hood Canal BS_1_8 22.8 43.85964912 4.39 98.68
      Hood Canal BS_1_9 13.9 71.94244604 7.19 161.87
      Hood Canal BS_1_10 27.7 36.10108303 3.61 81.23
      Fidalgo BS_1_4 8.9 112.3595506 11.24 252.81
      Fidalgo BS_1_5 18.1 55.24861878 5.52 124.31
      Fidalgo BS_1_6 16.1 62.11180124 6.21 139.75
      Fidalgo BS_1_7 35.8 27.93296089 2.79 62.85
      Fidalgo BS_1_8 22.7 44.05286344 4.41 99.12

      Left in ethanol and sodium acetate in -20C overnight.

Friday 9/18/15

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Ligation of adaptors to digested samples.

Need to make new adaptors each time. Recipe below is for 10 samples (+1 for pipet error) to make adaptors at 2 uM.  Combine and let sit for 10 minutes at room temp – I made sure stocks were completely melted.

Adaptor 1
11x
100 um 5ILL-NR
.6 uL
100 um Anti-ILL
.6 uL
NFW
58.8 uL
Adaptor 2
11x
100um 3ILL -NR
.6 uL
100um Anti-ILL
.6 uL
NFW
58.8 uL
Ligation reaction master mix
1x
11x
2 uM Adaptor 1
5 uL
55
1 uM Adaptor 2
5 uL
55
T4 ligase
1 uL
11
T4 ligase buffer with 10 mM ATP
4 uL
44
NFW
25
275

*Note: the Meyer protocol calls for 1.0 uL rATP  and 4.0 uL T4 ligase buffer. My buffer had 10 mM rATP included, so I just used 25 uL of NFW instead of 24uL and did not add more rATP.

40 uL master mix with 10 uL of digested DNA. Held at 16degC for 2 hours and 15 minutes. Put in -20C.

Wednesday 9/16/15

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2-B RAD Library Prep -Common Garden

Setup AlfI digestion of the 10 samples that were dried down and reconstituted on Monday following the Meyer Lab’s protocol. The SAM that was ordered was at .5 mM, so I made a dilution to 150 uM by adding 30 uL of stock to 70 uL of water.

1x
11x
10x buffer R
1.2 uL
13.2
150 uM SAM
.8
8.8
AlfI (2 U/uL)
.5
5.5
nuclease-free water
1.5
16.5
 Protocol calls for at least 1 hour at 37degC and up to overnight, so I set it for 2 hours followed by 15 minutes at 65degC.
Ran out gel of 2 uL digest and 1 uL of concentrated DNA. Meant to run out the digest next to the DNA for easier view, but forgot. I think I overloaded the DNA, will do .5 uL next time. What to expect:

  • “The original gDNA should appear as a single high-molecular weight band (>10kb) while the digested samples should be visibly degraded. An effective AlfI digestion produces a slight downward shift in the original HMW band and a subtle smear trailing downward from that band. If initial gDNA is degraded, this test is not useful and may be skipped.” -Meyer protocol
  1. SS_2_3 digest
  2. SS_2_4 digest
  3. SS_2_5 digest
  4. HC_1_1 digest
  5. HC_1_2 digest
  6. HC_1_3 digest
  7. HC_1_4 digest
  8. NF_1_1 digest
  9. SS_2_3
  10. SS_2_4
  11. SS_2_5
  12. HC_1_1
  13. HC_1_2
  14. HC_1_3
  15. HC_1_4
  16. NF_1_1
  17. NF_1_2 digest
  18. NF_1_2
  19. NF_1_3 digest
  20. NF_1_3
 gel_9_6_15
Some digests look good (SS_2_3, SS_2_5,HC_1_2,HC_1_3,NF_1_2, HC_1_4), some don’t look very degraded (SS_2_4) and some the DNA is degraded making it hard to tell (HC_1_1,NF_1_1,NF_1_3). For some reason NF1_3 dna does not show up at all- must have pipeted incorrectly. NF_1_1 and NF_1_3 looked degraded on the initial gel run, but weirdly HC_1_1 looked fine. I had to keep drying some of the samples after 10 minutes in the centrivap, which may have impacted DNA quality. Will talk to lab tomorrow, but my gut is to go ahead with library prep and sequence these on the MiSeq. If the 3 degraded samples have low read coverage, we’ll know how DNA quality effects library quality.

Monday 9/14/15

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Quantified broodstock samples that were extracted 8/19/15 and 9/11/15 with Qubit and ran out gel. Concentrations in ng/uL are below with score of quality based on gel.

9_14_15 Gel

  1. HC1_10: 27.7 ng/uL; 5
  2. NF1_8: 22.7 ng/uL; 3 (some degrad)
  3. HC1_9: 13.9; 3
  4. SS2_8: 16.8; 4.5
  5. HC1_8:22.8; 5
  6. NF1_2: 17.4; 4.5
  7. HC1_6: 23.5; 5
  8. HC1_1: 12.3; 4.5
  9. NF1_5: 18.1; 4.5
  10. SS2_7: 16.4; 5
  11. SS2_4: 10.2; 4
  12. SS2_3: 19.8; 5
  13. SS2_5: 7.83; 5
  14. HC1_5: 12.1; 5
  15. NF1_3: 27.8; 3 (some degrad)
  16. NF1_4: 8.9; 4
  17. HC1_4: 13.8; 4.5
  18. NF1_1: 9.13; 2.5 (low, some degrad)
  19. HC1_7: 14; 4.5
  20. NF1_7: 35.8; 5
  21. NF1_6: 16.1; 5
  22. HC1_2: 15.2; 2 (low)
  23. HC1_3: 43.5; 5
  24. SS2_6: 23.4; 5
  25. SS2_1; 5
  26. SS2_2; 1 (rerun)

Ethanol extraction of 10 broodstock DNA samples to prepare for test 2b-RAD library, following this protocol.

Population Sample Date extracted ng/uL Volume for 1 ug Volume sodium acetate Vol ethanol
Oyster Bay BS_2_3 19.8 50.50505051 5.05 113.64
Oyster Bay BS_2_4 10.2 98.03921569 9.80 220.59
Oyster Bay BS_2_5 7.83 127.7139208 12.77 287.36
Hood Canal BS_1_1 12.3 81.30081301 8.13 182.93
Hood Canal BS_1_2 15.2 65.78947368 6.58 148.03
Hood Canal BS_1_3 43.5 22.98850575 2.30 51.72
Hood Canal BS_1_4 13.8 72.46376812 7.25 163.04
Fidalgo BS_1_1 9.13 109.5290252 10.95 246.44
Fidalgo BS_1_2 17.4 57.47126437 5.75 129.31
Fidalgo BS_1_3 27.8 35.97122302 3.60 80.94

After drying samples for ~10 minutes at 37degC in a Centrivap Concentrator, I added 10 uL of nuclease-free water and left to rehydrate in 4degC fridge.

Friday 9/11/15

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Thought I’d be switching my online notebook over to the Open Notebook Science network, but when trying to write this post I found that I could not add a new category. Will try to figure that out, but until then will keep posting here.

Took my samples and reagents from the freezer in my lab on campus to the Pritzker DNA Lab at the Field Museum, where I do the bulk of my molecular work. Spent the day alternating between writing my Doctoral Dissertation Improvement Grant (DDIG) and some lab work.

Common Garden Experiment

  • Extracted DNA from 14 broodstock samples using the EZNA Mollusc kit (protocol here). Left to digest for 2.5 hours.
    1. SS2_6
    2. SS2_7
    3. SS2_8
    4. HC1_6
    5. HC1_7
    6. HC1_8
    7. HC1_9
    8. HC1_10
    9. NF1_3
    10. NF1_4
    11. NF1_5
    12. NF1_6
    13. NF1_7
    14. NF1_8
  • Ran out a gel from the DNA extractions of larvae samples done on 8/11/12 at Manchester.
    • Population Tank Family Size Date Storage Est. # Date extracted
      1 Hood Canal LC >100 7/13/2015 75/95 EtOH 8/11/2015
      2 Fidalgo Bay NA LC 100 7/13/2015 75% EtOH 8/11/2015
      3 South Sound NA LC 100 7/13/2015 RNALater 8/11/2015
      4 Hood Canal NA HC2 100 7/17/2015 RNALater 8/11/2015
      5 Hood Canal NA LC 100 7/17/2015 RNALater 8/11/2015
      6 South Sound NA LC 100 7/17/2015 RNALater 8/11/2015
      7 Hood Canal NA LC 100 7/20/2015 RNALater 8/11/2015
      8 Fidalgo Bay NA LC 100 7/20/2015 RNALater 8/11/2015
      9 South Sound NA LC 100 7/20/2015 RNALater 8/11/2015
      10 Hood Canal HC_Tank1_160 NA 160 7/20/2015 RNALater 8/11/2015
      11 Fidalgo Bay NF_Tank1_new NA 160 7/20/2015 RNALater 8/11/2015
      12 South Sound SS_Tank1_new NA 160 7/15/2015 RNALater 8/11/2015
      13 Hood Canal HC_Tank1_new NA 160 7/24/2015 RNALater 8/11/2015
      14 Fidalgo Bay NF_Tank1_new NA 160 7/24/2015 RNALater 8/11/2015
      15 South Sound SS_Tank1_new NA 160 7/24/2015 RNALater 8/11/2015
      16 Hood Canal HC_Tank1_new NA 160 7/27/2015 RNALater 8/11/2015
      17 Fidalgo Bay NF_Tank1_new NA 160 7/27/2015 RNALater 8/11/2015
      18 South Sound SS_Tank1_new NA 160 7/27/2015 RNALater 8/11/2015
      19 Hood Canal HC_Tank2_160 224 8/3/2015 RNALater 8/11/2015
      20 Fidalgo Bay NF_Tank2_160 224 8/3/2015 RNALater 8/11/2015
      21 South Sound SS_Tank2_160 224 8/3/2015 RNALater 8/11/2015
      22 Hood Canal HC_Tank2_160 >224 8/7/2015 RNALater 350 8/11/2015
      23 Fidalgo Bay NF_Tank2_160 >224 8/7/2015 RNALater 504 8/11/2015
      24 Oyster Bay SS_Tank2_160 >224 8/7/2015 RNALater 641 8/11/2015
    • The computer hooked up to the UV camera wouldn’t recognize a USB drive, so I just have a phone picture of a printed out picture for now. 12 and 13 are cut off, but they did not show up on gel. For gels of DNA extracts, I give a ranking from 1 (did not work at all) to 5 (bright band of high molecular weight DNA). These are put of the Sample Master Sheet.
      • gel_9_11_15
        1. 3.5
        2. 4
        3. 4
        4. 1 (rerun)
        5. 4
        6. 4.5
        7. 3.5
        8. 3 (low)
        9. 5
        10. 4
        11. 1 (rerun)
        12. 1 (rerun)
        13. 1 (rerun)
        14. 1 (rerun)
        15. 5
        16. 3 (degrad)
        17. 2.5 (degrad)
        18. 2 (degrad)
        19. 5
        20. 1 (rerun)
        21. 4.5
        22. 3 (degrad)
        23. 2.5 (low)
        24. 5
  • Playing around with data!
    • Looked at larvae release from each population across time. Did this partially out of burning curiosity and because it would be nice to have some preliminary data to put in my DDIG proposal.
    • First, I had to edit the Google Sheet to include zeros for days when a population released no larvae and to have total counts across families. Then in R:
    •  > larvae = read.csv("Larval counts - Day 1 (1).csv", header = TRUE) 
> names(larvae)

      [1] “Date” “Population” “Family” “Tank.added.to” “Volume.of.tripour..mL.” “Vol.of.drop.counts” “Ethanol.used.”
      [8] “Live.Count.1” “Live.Count.2” “Live.Count.3” “Live.Count.4” “Total.Live.Larvae” “X…” “Notes”
      [15] “Dead.count..1” “Dead..2” “Dead..3” “Dead..4” “Total.Dead” “Total.Larvae” “Total.by.date”

 > pop_Total_Date_na <- na.omit(pop_Total_Date)
> ggplot(data=pop_Total_Date_na, aes(x=Date, y=Total.by.date, \
group=Population, colour=Population)) + geom_line() + geom_point()

Day1Larvae_date

  • Interestingly, the South Sound population produced more larvae earlier. This mirrors the reciprocal transplant experiment, where SS oysters reached their maximum percentage of brooding females sooner at two of the 4 sites.

Oly Population Structure

  • In addition to making libraries for samples from the common garden, I need to start getting DNA ready for one more sequencing run for my project looking at rangewide population structure in Olympia oysters. Did 10 extractions concurrently with the common garden extractions.
  1. WA1_16
  2. WA1_14
  3. BC1_19
  4. BC1_18
  5. CA6_16
  6. CA6_17
  7. CA6_18
  8. CA7_13
  9. OR3_7
  10. OR3_20

Monday 8/24/15 – Thursday 8/27/15 (Last week of fieldwork!)

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Monday 8/24/15

Had a long chat with my advisor, Cathy, about the status of my fieldwork- what data I had collected, what I was leaving behind, plans for a stressor experiment- as well as plans for funding next summer. Puget Sound Restoration Fund has approved me to conduct a common garden experiment next summer with olys from California, Puget Sound, and British Columbia. My hope is to use this rangewide common garden to contextualize the regional-scale one I did this summer.

Tile culling

  • SSA_7: 9
  • SSA_8: 83
  • HCB_1: 160
  • HCB_2: 310
  • HCB_3: 384
  • Took pictures of tiles that were culled last Thursday but didn’t have pictures for.

Growth Rate Experiment

Started taking pictures of larvae from the growth rate experiments conducted earlier in the summer. This was done using an adaptor to attach a phone to the eyepiece of a microscope. Labelling scheme (until I think of a better one) is Population+Replicate_Growth rate experiment (1 or 2)_date sample was taken. Also did live/dead counts of the entire sample.

  • SS3_G1_7/12: 25L/15D
  • SS1_G1_7/12: 57L/8D
  • SS3_G1_7/15: 38L/14D
  • SS2_G1_7/12: 64 L/9D

Tile Set B

  • Cleaned out the tanks from tile set B. Cut off the tiles and randomized them among the wire “cages” in the large tank with the other tiles from Set_A.

Tuesday 8/25/15

Cultch Set

  • SS>450A + SS>450B -> SS> 450B; SS>1000
  • SS_new -> SS>450B; dump
  • HC > 450 ->  HC > 1000
  • HC_new -> HC > 450; dump

Tile culling

  • HC_B_1: 42
  • HC_B_2: 103 from front, 164 from back
  • HC_B_3: 67 front, 42 back
  • HC_B_4: 4
  • HC_B_7: 4 front, 13 back
  • HC_B_8: 12 front, 3 back
  • HC_B_10: 9 back
  • HC_B_11: 1 back
  • HC_B_12: 2 back
  • HC_B_14: 86 front, 5 back
  • SS_B_1: 7 back (none on front)
  • SS_B_2: 4 back (none on front)
  • SS_B_3: 0/3
  • SS_B_7: 0/10
  • SS_B_8: 0/2
  • SS_B_9: 0/5
  • SS_B_10: 0/7
  • SS_B_11: 0/6
  • SS_B_13: 0/1
  • NF_B_1: 0/27
  • NF_B_2: 95/120
  • NF_B_3: 0/13
  • NF_B_4: 77/270
  • NF_B_5: 8/12
  • NF_B_6: 208/92
  • NF_B_7: 0/2
  • NF_B_8: 18/10
  • NF_B_9: 8/19
  • NF_B_10: 70/48
  • NF_B_11: 11/17
  • NF_B_12: 10/27
  • NF_B_13: 1/12
  • NF_B_14: 0/2

Wednesday 8/26/15

  • Dropped off samples and borrowed materials at the Roberts lab.

Thursday 8/27/15

The little oyster babies are leaving the nest for the big open ocean! I started out the day finishing up any culling that was needed. I also looked over previously culled tiles to make sure that at least 1 cm of space was around each oyster.

Culling

20150911_121151

Steven Roberts came to help with the deployment. While I finished up culling, he made labels for the trays but cutting small PVC pipe into 1 inch pieces and etching numbers onto them. We’re also putting waterproof paper in a tube with the tray number.

12 tiles (4 per population) were attached to each tray with zipties. The populations were ordered in the same way for each section of each tray (NF, HC, SS).

Tray with tiles attached

Tray with tiles attached

Pictures were taken of each tile next to a ruler to measure size at deployment of the oysters.

HCA11_150827

7 trays were filled up with with tiles. To minimize the effects of location within a stack of trays, we put 4 trays in a stack with a 2′ spacer between the top 2 and bottom 2 trays. An additional tray was used as a cover to keep out predators. We made 2 of these stacks, with an empty tray in Stack 2.

Stack of trays with oyster tiles ready for deployment

Stack of trays with oyster tiles ready for deployment

Order of tiles and trays

20150911_12201820150911_122037

The trays were hung of the dock at Manchester by ~20 foot rope. One of the stacks seemed to float (which was odd), so we tied on clam bags with rocks to both stacks.

20150827_15371020150827_153657IMG_3008

Wednesday 8/19/15 and Thursday 8/20/15

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Wednesday 8/19/15

There was a silo making party at the hatchery today for the rock scallop project. With little animal husbandry to do, I decided to do a DNA extraction and help out making silos while the tissue was digesting.

Tile Culling

  • HCA_7: 33 removed
  • NFA_1: 123
  • NF_A_2: 134
  • NF_A_3: 102
  • NF_A_4: 32

Cultch Set

NF_A + NF_New -> NF> 450_B; NF_New

DNA Extraction

Took tissue out of RNALater onto sterile weigh boat and cut off ~30 mg of tissue with scalpel (have previously weighed out chunks of adductor muscle tissue to help eyeball this amount). Replaced rest of tissue in RNALater. Wiped scalpel and forceps with 85% ethanol and sterilized over benchtop bunsen burner in between samples. Otherwise, followed EZNA protocol as described in a previous post.

  1. SS2_1
  2. SS2_2
  3. SS2_3
  4. SS2_4
  5. SS2_5
  6. HC1_1
  7. HC1_2
  8. HC1_3
  9. HC1_4
  10. HC1_5
  11. NF1_1
  12. NF1_2

Thursday 8/20/15

Tile Set B

  • bleached line
  • cleaned tanks
  • did counts for HC_SetB and NF_SetB

Tile Set A

Had a discussion with Ryan on the ferry about outplanting my tiles. He recommended transitioning the setters slowly to ambient temperature (they are still on heated at 18degC). My room at the hatchery is not really set up for this, so Alice helped me set up one of the large tanks outside that has a heated and an ambient spigot. I tried to just set the tiles on the large 400um mesh screens, but they did not all fit so I just randomized the tiles among 3 poultry wire setups (the same that were in the setting system tanks). The spigots are at one end of the tank and the outflow is at the other, promoting water flow. Food is splashed in by hand and then added in through a dropper. Some of the outflow is actually pumped back in to the side of the tank with the tiles to reduce algae waste.

Tile culling

  • NFA_5: 29 removed
  • SSA_3: 15
  • SSA_5: 61
  • SSA_2: 10
  • SSA_6: 13

Cultch Set

  • NF >1000 -> NF >1600; NF> 1000
  • NF > 450 ->  NF > 1000; NF > 450
  • HC > 1000 -> HC > 1600; HC > 1000
  • HC> 450  ->  HC > 1000; HC > 450

Sunday 8/16/15 – Tuesday 8/18/15

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Sunday 8/16/15

Came in on a Sunday to knock out some more dissections and transition to a Sunday/Tuesday/Thursday cleaning for this week since I’d be out of town on Friday.

Dissections

  • NF3 and NF5

Husbandry

  • Bleached the line
  • Rinsed tiles and cultch
  • Cleaned all of the A tanks, HC_SetB, and NF_SetB
  • Counts for HC_SetB and NF_SetB

Monday 8/17/15

Cultch Set

  • HC SetA + HC Set New -> HC >450; HC Set New
  • HC > 450 -> HC >1000; HC > 450

Dissections

  • HC1, HC2, HC2, HC3

Husbandry

  • rinsed tiles and cultch
  • cleaned SS_SetB tank

Tile Culling

Started culling for density off of tiles from A sets that were very crowded. Using a clean scalpel, I would remove spat so that only ~30 were left on a side of a tile. I put the spat in a 1.5 mL tube, rinsed with freshwater Millepore water, then stored in RNALater. I kept track of how many were taken from a tile, but in some cases it is an estimate when I was scraping off many that had settled on top of each other. My primary goal was to reduce the time the tile was out of the water.

  • HC_SetA_1: 300 removed
  • HC_SetA_2: 160
  • HC_SetA_3: 17 from front, 8 from back
  • HC_SetA_4: 420 from front

Tuesday 8/18/15

Dissection

  • HC4, HC5 (finished!)

Husbandry

  • Bleached the line
  • Cleaned all tanks
  • Counts for B set

Cultch Set

  • SS_SetA + SS_Set_New -> SS >450; SS_Set_New (on 1000 cultch)

Tile Culling

  • HC_SetA_4: 400 front
  • HC_SetA_5: 150
  • HC_SetA_6: 76

Wednesday 8/12/15- Friday 8/14/15

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Wednesday 8/12/15

Larvae tanks

  • NF_Tank2_160 (224) -> NF cultch set
  • NF_Tank2_160 (100) -> swimmers only
  • HC_Tank2_160 (224) -> HC cultch set
  • HC_Tank2_160 (100) -> swimmers only
  • SS_Tank2_160 (224) -> barely any, put back in tank
  • SS_Tank2_160 (100) -> swimmers only

Tile Set

  • Did live/dead counts for both A and B
  • Did not add the unset larvae back to A
  • Rinsed tiles

Cultch Set

  • NF >450 -> NF >1000
  • SS > 450 -> SS>1000
  • SS_SetA -> SS > 450; SS_SetA

Thursday 8/13/15

Started dissections of tissue from the broodstock for subsequent DNA extractions with the help of Brent Vadopalas and the PSRF intern Ryann. First, we recorded the weight of each oyster in a family and placed it on a numbered pad.

Broodstock dissection setup

Broodstock dissection setup

A picture was taken of the entire family with a ruler for later measurement in ImageJ. Effort was made to hold the phone level to avoid the impact of tilt on apparent size. We made a little assembly line, with Brent shucking before joining Ryann and I in dissecting out adductor muscle tissue and storing in 1.5 mL tubes with 1 – .75 mL RNALater. If there was not very much muscle tissue, mantle or the entire oyster were taken as well. Scalpels and forceps were rinsed sequentially in soapy water, bleach, and freshwater between each oyster. Fresh scalpels were used between populations. We got through SS2, SS1,SS3, SS4, SS5, and NF4, averaging about 45 minutes per family by the time we got the hang of it.

Husbandry

  • Rinsed tiles and cultch
  • Feeding

Friday 8/14/15

Larvae tanks

  • NF_Tank2_160 (224) -> NF_New cultch set
  • NF_Tank2_160 (100) -> dumped
  • HC_Tank2_160 (224) -> none
  • HC_Tank2_160 (100) -> dumped
  • SS_Tank2_160 (all) -> dumped

Tile Set Counts

  • NF_SetA
  • SS_SetA
  • HC_SetA
  • HC_SetB
  • SS_SetB

Dissections

Did NF2 and NF1 with a little help from Ryann on NF1

Monday 8/10/15 and Tuesday 8/11/15

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Monday 8/10/15

Larvae tanks

  • NF_Tank2_160 (224) -> 10,500 total: 500 for DNA, rest to NF_SetB
  • NF_Tank2_160 (100) -> swimmers only
  • HC_Tank2_160 (224) -> 9,187 total: 500 for DNA, 5,187 added to HC_SetB, rest to cultch set
  • HC_Tank2_160 (100) -> swimmers only
  • SS_Tank2_160 (224) -> 11,900 total, added to SS cultch
  • SS_Tank2_160 (100) -> swimmers only

Tile Set

  • Did live/dead counts for both A and B
  • Rinsed tiles (have been doing this everyday I’m in)
  • New totals
    • NF_SetB: 45,137
    • HC_SetB: 60,000

New larvae

  • no new larvae

Tuesday 8/11/15

Animal husbandry

  • Cleaned broodstock buckets
  • Rinsed cultch and tiles

Lab work

Extracted DNA from 24 larvae samples that would be good candidates for test 2b-RAD libraries. I chose sets of new larvae (“LC” for larvae catch), “160”s, and “224”‘s with 3-5 days in between.

Population Tank Family Size Date Storage Est. # Date extracted
1 Hood Canal LC >100 7/13/2015 75/95 EtOH 8/11/2015
2 Fidalgo Bay NA LC 100 7/13/2015 75% EtOH 8/11/2015
3 South Sound NA LC 100 7/13/2015 RNALater 8/11/2015
4 Hood Canal NA HC2 100 7/17/2015 RNALater 8/11/2015
5 Hood Canal NA LC 100 7/17/2015 RNALater 8/11/2015
6 South Sound NA LC 100 7/17/2015 RNALater 8/11/2015
7 Hood Canal NA LC 100 7/20/2015 RNALater 8/11/2015
8 Fidalgo Bay NA LC 100 7/20/2015 RNALater 8/11/2015
9 South Sound NA LC 100 7/20/2015 RNALater 8/11/2015
10 Hood Canal HC_Tank1_160 NA 160 7/20/2015 RNALater 8/11/2015
11 Fidalgo Bay NF_Tank1_new NA 160 7/20/2015 RNALater 8/11/2015
12 South Sound SS_Tank1_new NA 160 7/15/2015 RNALater 8/11/2015
13 Hood Canal HC_Tank1_new NA 160 7/24/2015 RNALater 8/11/2015
14 Fidalgo Bay NF_Tank1_new NA 160 7/24/2015 RNALater 8/11/2015
15 South Sound SS_Tank1_new NA 160 7/24/2015 RNALater 8/11/2015
16 Hood Canal HC_Tank1_new NA 160 7/27/2015 RNALater 8/11/2015
17 Fidalgo Bay NF_Tank1_new NA 160 7/27/2015 RNALater 8/11/2015
18 South Sound SS_Tank1_new NA 160 7/27/2015 RNALater 8/11/2015
19 Hood Canal HC_Tank2_160 224 8/3/2015 RNALater 8/11/2015
20 Fidalgo Bay NF_Tank2_160 224 8/3/2015 RNALater 8/11/2015
21 South Sound SS_Tank2_160 224 8/3/2015 RNALater 8/11/2015
22 Hood Canal HC_Tank2_160 >224 8/7/2015 RNALater 350 8/11/2015
23 Fidalgo Bay NF_Tank2_160 >224 8/7/2015 RNALater 504 8/11/2015
24 Oyster Bay SS_Tank2_160 >224 8/7/2015 RNALater 641 8/11/2015

I still have to go back over my notes to get estimated number for some of them. Storage was mostly in RNALater. A lot of the samples had some white precipitate on the bottom. If larvae weren’t  in the precipitate I sucked it out before adding the lysis buffer/proteinase K. Also had the same issue previously with larvae being buoyant in the RNALater even after a spindown. Halfway through trying to siphon off as much liquid as I could, I did some research and found that addition of some ice-cold PBS will change the density of the liquid and allow the larvae to settle out. This worked really well and was done for 2,10,13,15,19,20,21,24. It also dissolved most of the white precipitate.

Followed the protocol I listed here, with a 2.5 hour digest. Could not findd the gel rig set-up (found out later it’s in a different building).