Thursday 6-25-15

Went to the hatchery in the morning to check for larvae, clean/replace banjo filters, rinse drippers, and replenish algae. As there was a hatchery-wide mortality event after the temperature drop over the weekend, I started 2 new 100 L larval tanks: SS_Tank2 and HC_Tank2. This is because it’s pretty hard to separate out live from dead larvae until the live larvae have grown up a bit. In case the dead larvae still in SS_Tank1 and HC_Tank1 cause additional mortality, I want to start a 2nd replicate where none of the larvae experienced the temperature drop.

  • Checking for larvae in larval catch buckets
    • Maybe/Some larvae: SS1
    • Lots/Noticeable larvae: HC3, HC4, HC1, SS4
    • Also screened NF5 but did not get any larvae
    • Added to HC_Tank2 and SS_Tank2
  • Counting larvae
    • need to get better at estimating ~100 per mL. Some of these counts had over 400 per mL which makes it harder to count and more stressful for the larvae when they’re in the tripour beaker

Larval counts data DNA samples

  • As per my conversation with S. Roberts yesterday, I’m keeping all of the samples from my larvae counts, including those I don’t think are necessary for future genotyping. I’m trying 2 different preservation methods, and will do test DNA extractions next week to look at quantity and quality.
    • 75%/90% ethanol: this was suggested by Jake Heare. I pipet up one of the 1 mL larvae count samples and put it in a 1.5 mL tube. Since I used ethanol when counting, all of the larvae sink quickly to the bottom and I pipet out as much seawater as I can. I continue adding the other larvae count samples and decanting off the seawater in the same fashion. If needed I spin them down briefly with a centrifuge. Then I add 75% ethanol to the tube. A day or 2 after I take out the 75% ethanol and add 90% ethanol. These are stored in -20C.
    • RNALater: As described above, I take out as much seawater as I can and then add about .75 mL of RNALater to the tubes. These are stored in a -80C freezer.

In the afternoon, I stopped by the Roberts lab on the UWashington campus to find some bench space for the test DNA extractions and pick up a HOBO temperature logger to stick in one of my broodstock buckets so I can get a better idea of what they’re experiencing in case a temperature malfunction happens again.

Wednesday 6-24-15

Steven Roberts came to help out today, which made the Wednesday cleaning/collecting larvae only take about 2.5 hours. He did all of the larval counts, filtered out the SS larval catches, and cleaned many of the buckets. We decided in the interest of time to just qualitatively note individual family output of larvae and then combine the larval output from each family in a population for counting.

Families that were filtered for larvae:

  • very little/some larvae: HC2, SS5, SS2, SS4, SS3
  • lots: NF5, HC3
    • for both of these the larvae seemed to be clustering near the bottom (dead??)
SS5 seemed to release some sperm when put in the bucket with all the other SS families.
I’ve organized my datasheet for larval counts a little differently.
  • The 1st sheet, “Day 1”, is the raw data from counting the larval output from the broodstock.
  • The 2nd sheet, “Larval tank counts”, is the raw data from counting the number of live/dead larvae filtered out of the 100 L larval tanks. It also lists the estimated ages of the cohorts in the tanks.
  • The 3rd sheet- “Larval tanks running total”- is formatted like a bank account transaction list, with larval tanks for each population the “accounts”. This incorporates data from the first 2 sheets to let me know approximately how many larvae I have in the 100 L tanks at a time. The differences between counts is also my estimated mortality (although some of this is due to leakage during cleaning).

Update 6-25-15

  • On Tuesday 6-23-15, the seawater was shut off for about 3 hours. Also, there was an issue with the airline so that tanks may not have been bubbling for a while. When I went in on Wednesday, all tanks seemed to be bubbling fine.

Monday 6/22/15

Larvae collection, counting, and sampling

I’m being really paranoid at this point as I want to try and “catch ’em all”. I’ve modified my larvae collecting a little bit by stacking a 400 micron screen on top of the 100 micron screen to catch the poop/gunk.

  • SS3, SS5, SS4, HC4, and HC5 had low or suspect number of larvae
  • SS1, SS4, SS2, HC2 had a definitely noticeable or large number of larvae
  • HC4 and HC5 did not have any larvae when checked with the dissecting scope
  • Data sheet with larval counts

After I counted the larvae samples, I dumped them into into the appropriate HC or SS 100 L tanks, after they had been cleaned and filled up with water. In the afternoon, Ryan (the hatchery manager) told me that there had been a malfunction with the heated seawater line sometime between Friday afternoon and Saturday when he came in. All of the tanks were at ~15C, and they observed severe mortality in their larvae tanks. The larvae I had collected from the larval catch buckets were mostly healthy. There were a few empty shells (from now on I’ll count those too), but they were definitely less than 5%. The problem might be if there was a severe mortality event in my 100 L tanks. Those dead larvae would have been filtered out with the healthy ones while I was cleaning and put back in, this could make the rest of the larvae sick from swimming around with dead ones. Unfortunately I had already put my newly collected larvae in those same tanks.

There were 29,000 larvae in the HC tank and 87,850 in the SS tank over the weekend. There are now 83,600 larvae in the HC tank and 363,437 in the SS tank. On Wednesday when I clean these out, I will filter them over a 160 micron screen stacked on a 100 micron screen. I’ll do counts of both sizes, and in particular look at the proportion of dead larvae in the smaller size. Any larvae that didn’t die or get sick on Saturday would have grown since, while those that did would still be passing through a wider screen.

June 15, 17, and 19 2015

I’m planning on this being my last “week in review” post, as my fiancé and I finally moved into our place and are no longer drifting homeless through Seattle. I also have oyster babies now, so the data collection can officially begin!

Monday 6-15-15

Checked the buckets for larvae-did not see any. No dead broodstock.

Did the M-W-F cleaning

  • an oyster from a South Sound family (SS5) had fallen out of the bag and was in the bottom of the bucket while cleaning. May have been exposed to very dilute bleach. I put it back in the bag and will keep an eye on that family.

Started taking pictures of the adults for size measurements in Image J. To do this, I take one family at a time out of it’s bucket and gently take the oysters out of the bag. An oyster is placed next to a ruler and I took pictures of dorsal/ventral sides using my Galaxy S6 phone (which should be more than adequate quality). I’ll load these on to Dropbox and put the link here this weekend.


*note: HC = Hood Canal, SS = South Sound, NF = Fidalgo Bay (central sound)

  • Took photos of HC1_1-20, HC4_1-20, HC5_1-15, SS1_1-20, NF1_1-21
  • Originally counted 20 in NF1, but found a very small (

Ryan helped me get out the six 100 L larvae tanks and showed me how to set them up and clean them once I start getting larvae.

Wednesday 6-17-15

Checked the buckets for larvae.

  • Saw little black specs in HC2 so filtered them out over a 240 micron mesh screen onto a 100 micron screen. Sure enough they were larvae! I felt a little bit like a chicken with its head cut off- running around trying to remember what to do and figure out the best order for all the cleaning tasks so that no one was out of the water or crowded together for too long. There was a big spawn going on for another experiment, so I wasn’t able to take a larval count. I filled up a 100 L tank and put the HC2 larvae in there, planning on counting them on Friday.

Did M-W-F cleaning.

  • No dead broodstock

Ryan gave me some advice on how to deal with cleaning, filtering out larvae, and counting larvae. He suggested:

  1. Cleaning the line out with freshwater/bleach
  2. Taking the larvae out of the 100 L tanks and clean those
  3. Refill the 100 L tanks and replace the larvae
  4. Filter out any newly spawned larvae from the buckets. Leave them in a tripour, mixing frequently.
  5. Clean out the broodstock and larval catch buckets.
  6. While those are refilling, do the larvae counts. Pour the larvae in the appropriate 100 L tank after counting.

Friday 6-19-15

Checked the buckets for larvae.

  • SS1 and SS3 had noticeable larvae in the broodstock buckets only. They were lighter in color than the HC2 larvae, which has been observed previously in South Sound oysters.
  • Filtered them out over 240/100 micron screens and put them in 700 mL of seawater in separate tripour containers.

Flushed out the line.

Emptied out the HC 100 L larvae tank onto a 100 micron screen and put those larvae in a separate tripour with 800 mL seawater.

Cleaned the broodstock/larval catch buckets and filled up two 100 L tanks.

To count larvae, I mix up the tripour with the larvae, take four replicates of 1 mL samples and put them each in a clear well plate. I then count these under a microscope. After I had finished counting, someone recommended that I put a little ethanol on them 1st to keep them from swimming around. I’ll do this from now on, as trying to keep track of which ones I’ve already counted while they were swimming might explain the variance in my counts.

How To Raise an Oyster

Keeping oysters alive in a hatchery requires almost daily care. Monday, Wednesday, and Friday I flush fresh water and bleach through the line that brings them seawater. On these days I also give them a little rinse with fresh water, check for any “gapers” (dead oysters), clean all the poop out of their buckets, clean all of the tubing, and clean the larval catch buckets.

Broodstock bucket and larval catch bucket

Broodstock bucket and larval catch bucket

Oysters hanging out

Oysters hanging out






All 15 families

All 15 families

Every day I’m at the hatchery, I make sure they have enough algae, clean and replace the banjo filters in the larval buckets, empty out any places in the line where water might stagnate, and clean the water drippers. Much of this is to prevent unwanted algal and bacterial growth, as well as minimize the spread of disease.

100 micron banjo filer

100 micron banjo filer

Algae culturing

Algae culturing










Once I have larvae, this workload will increase. I’ll be filtering out larvae every day and putting them in 100 L tanks. I’ll have 2 tanks per population group, so 6 total. These tanks will also require cleaning 3 times a week, and with millions more oysters to take care of, I’ll need to harvest more algae. I’m still a bit slow at the cleaning process, but soon I should get it down to ~4 hours of general animal husbandry on M-W-F which will leave me plenty of time for data collection and molecular lab work.


100 L larval tank

100 L larval tank

A larger 100 micron banjo filter is used in the larval tanks

A larger 100 micron banjo filter is used in the larval tanks

Setting up an Oyster Garden

Monday (June 8) was my first day out at the NOAA Manchester Research Station in Washington State. Specifically, I’m working in the Kenneth K. Chew Center for Shellfish Research and Restoration. This shellfish hatchery is the result of collaboration between many groups and funding agencies, in particular the Puget Sound Restoration Fund (PSRF).

The hatchery (right) and algae greenhouse (left)

My project this summer is to raise oysters descended from three Puget Sound populations under common conditions in order to measure differences in fitness. This type of experimental design is commonly referred to as a “common garden”, and allows one to control environmental variables so that phenotypic disparities among individuals can be attributed to their genetic differences. My fitness metrics are reproductive output, survivorship at different life stages, and growth rate. I will also be taking DNA/RNA samples along the way to see if mortality is random in respect to genotype, or due to purifying selection. With the RNA, I plan to look at differences in gene expression to help detect cryptic differences in phenotype between these populations.

Three source populations for common garden experiment

Three source populations for common garden experiment

This project is a collaboration with Steven Robert’s lab at the University of Washington, who previously conducted a reciprocal transplant experiment with offspring of wild oysters from these same populations. For that experiment, they outplanted the young oysters from each group at four different sites and measured growth rate, mortality, and reproductive characteristics. They observed significant variation at these metrics among populations and sites (informative slides and manuscript preprint available here). My experiment will be following up on these results by testing if population-level differences are consistent in a second generation under controlled environmental conditions.

As I’ve never raised shellfish before, this week has had a bit of a learning curve. Fortunately for me, the staff at the hatchery have been super helpful in showing me the ropes and advising on how to set up my experiment. I’m starting with about 100 adult oysters for each group (see lab notebook entries for data). These are the first generation (F1) offspring of wild oysters, and have been living in common conditions their entire lives- mostly hanging off the docks near the hatchery. Their offspring will be 2nd generation (F2) from the original broodstock, and should have any influence from maternal effects erased.

The adults were brought in to the hatchery on May 28 and placed in three separate buckets to avoid cross fertilization. To maximize genetic diversity and minimize the chance that one male fertilizes all of the females, I split each group into 5 buckets of ~20 oysters. These “families” will be marked, so that I can genotype them later and follow their offspring’s success throughout the experiment. Their water temperature was switched to a balmy 20°C this week, which will encourage them to start spawning and producing larvae.

(sorry for the lack of pictures, I’ll take some and put them up soon!)