Monday 9/28/15

Do AlfI digestion of ethanol precipations from last week.
  1. SS2_3
  2. HC1_1
  3. NF1_1
  4. NF1_2
  5. SS2_6
  6. SS2_7
  7. SS2_8
  8. HC1_5
  9. HC1_6
  10. HC1_7
  11. HC1_8
  12. HC1_9
  13. HC1_10
  14. NF1_4
  15. NF1_5
  16. NF1_6
  17. NF1_7
  18. NF1_8
1x
20x
10x buffer R
1.2 uL
24
150 uM SAM
.8
16
AlfI (2 U/uL)
.5
10
water
1.5
30

Did ligation of same samples. Left for 2.5 hours at 16degC and did 10 minutes at 65degC following Matz protocol.

  • 1st make new adaptors.
  • Adaptor 1
    11x
    20x
    5ILL-NR
    .6 uL
    1.1
    Anti-ILL
    .6 uL
    1.1
    NFW
    58.8 uL
    106.91
    Adaptor 2
    11x
    3ILL -NR
    .6 uL
    Anti-ILL
    .6 uL
    NFW
    58.8 uL
    Master mix
    1x
    20
    2 uM Adaptor 1
    5 uL
    100
    2 uM Adaptor 2
    5 uL
    100
    T4 ligase
    1 uL
    20
    T4 ligase buffer with 10 mM ATP
    4 uL
    80
    NFW
    25
    500

    40 uL master mix + 10 uL digestion product

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A taste of data analysis

Playing around with some data on protoshell length taken by Ryann (PSRF intern) from the salinity experiment.

In R:

 > plot(Length~Population, data = sal30df)
sal30_length

Boxplot of protoshell length in 7 day old Olympia oyster larvae from 3 populations in Puget Sound. n = 20 per population

Did an anova:

> aov.sal30 <- aov(Length~Population, data=sal30df)
> summary(aov.sal30)

Df Sum Sq Mean Sq F value Pr(>F)
Population 2 9281 4641 14.81 6.61e-06 ***
Residuals 57 17861 313

Signif. codes:  0 ‘***’ 0.001 ‘**’ 0.01 ‘*’ 0.05 ‘.’ 0.1 ‘ ’ 1

Indicates that the means of protoshell length are significantly different (yay!).

> pop <- sal30$Population
> len <- sal30$Length
> ptest <- pairwise.t.test(len, pop, p.adjust = "bonferroni")

Pairwise comparisons using t tests with pooled SD

data: len and pop

Fidalgo Bay Hood Canal
Hood Canal 0.0913             –
Oyster Bay 0.0069          3.9e-06

P value adjustment method: bonferroni

Very significant p-value between Hood Canal and Oyster Bay.

Wednesday 9/23/15

  • Did more extractions of broodstock samples. I think once this test library is done the broodstock libraries should be 1st priority, with samples from the stressor experiment 2nd and other larvae samples 3rd.
    • Used the EZNA kit with a 4 hour digestion time. All the tissue for HC1_16 was used up.
  1. HC1_11
  2. HC1_12
  3. HC1_13
  4. HC1_14
  5. HC1_15
  6. HC1_16
  7. HC1_17
  8. HC1_18
  9. HC1_19
  10. HC2_1
  11. HC2_2
  12. HC2_3
  13. HC2_4
  14. HC2_5
  15. HC2_6
  16. HC2_7
  17. HC2_8
  18. HC2_9
  19. HC2_10
  20. HC2_11
  21. HC2_12
  22. HC2_13
  23. HC2_14
  24. HC2_15
  • Finished the ethanol precipitation started on Tuesday 9/22/15. Left samples in 10 uL of water in 4degC overnight.
  • Qubit of samples for phylogeographic study. Made list of samples for GBS library.
      1. BC1_12: 25.2
      2. BC1_8: 6.44 (8)
      3. BC2_7: 10.6
      4. BC2_13: 11.5
      5. BC2_9: 13 (8)
      6. BC3_13: 108
      7. BC3_12: 15 (8)
      8. BC4_2: 8.84
      9. BC4_17: 11.4 (8)
      10. CA1_18: 24.4
      11. CA1_1: 36
      12. CA1_2: 17.4 (8)
      13. CA2_9: 43.8
      14. CA2_12: 18.8 (5)
      15. CA3_6: 62.3 (5)
      16. CA4_1: 5.1
      17. CA4_7: 19.8 (8)
      18. CA5_10: 48.8 (8)
      19. CA6_8: 5.14
      20. CA6_15: 25.2 (7)
      21. CA7_11: 7.4
      22. CA7_16: 19.1
      23. CA7_15: 14.9 (8)
      24. OR1_ 1: 7.66
      25. OR1_5: 29.2 (7)
      26. OR2_6: 12.9
      27. OR2_12: 15.8
      28. OR2_15: 18.2 (7)
      29. OR3_15: 9.24
      30. OR3_20: 13.7 (7)
      31. WA10_16: 25.4
      32. WA10_13: 9.52 (8)
      33. WA11_10: 18.7
      34. WA11_22: 17.4
      35. WA11_4: 11.7
      36. WA11_17: 4.78 (8)
      37. WA12_15: 24.4
      38. WA12_11: 4.18  (8)
      39. WA13_12: 7.2
      40. WA13_5: 4.4
      41. WA13_15: 45.1 (9)
      42. WA9_2: 11.4 (7)
      43. WA1_16: 62 (8)
      44. Conch_1: 12.4
      45. conch_2: 12.3
      46. conch_4: 9.65
      47. CA5_12: 21.8 (repeat with 2)
      48. CA5_10 repeat within

Tuesday 9/22/15

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

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

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

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

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