Another week in review…6/29-7/2

So track record for daily posts aren’t great, but I’m working on it. This was a pretty eventful week in the hatchery- setting up a system for settlement, some unfortunate mortality, and getting locked out of the molecular lab (now I know to keep a stash of ethanol/RNALater/tips in the hatchery!).

Monday 6-29-15

Since I cleaned out the 100L larval tanks on Sunday, on Mon-Wed-Fri  I’m now only flushing the line with bleach, cleaning all the drippers, cleaning all the banjo filters, collecting any spawned larvae, and cleaning the broodstock/larvae catch buckets. On Tues-Thurs-Sat/Sun I’ll filter out the larvae from the 100L tanks and clean those, as well as the daily banjo filter and dripper cleaning. I’ll also check for newly spawned larvae on those days and filter those out to count and add to a 100L tank.

Checking for new larvae:

  • Some: NF1(?), HC3
  • Lots: SSS1, HC5, SS2, SS3, HC4, SS4, HC2
  • Got about 222,000 larvae from the SS buckets to add to SS_Tank2 and 230,000 from the HC buckets to add to HC_Tank2
  • None from NF

Had time to filter out what was spawned in the SS group bucket during cleaning but not the HC bucket, although there was no noticeable larvae in that one.

There’s been an issue with the algae cultures since Friday 6/26, where there hasn’t been any diatoms to feed the animals. The recommended diet is 50% diatoms and 50% flagellates, so the larvae might have a deficiency of necessary amino acids.

Tuesday 6-30-15

Checking for larvae:

  • Some/Maybe: HC3, NF1(?), NF2(?), SS1(?), NF5(?)
  • Lots: HC4
  • none in SS1 or NF families, just poop

Today I decided to try out doing “weight counts” as well as my usual counts for the 100 L larval tanks. With this method, you filter out a 100 L tank over a few different sized mesh screens. The larvae of each size class is put in a small (approx. 15cm diameter, 10 cm height) pvc silo with a 100 micron mesh screen at the bottom, where the weight of each silo when empty is known. You weigh the silo+larvae, subtract the weight of the silo, and use a conversion sheet to get the estimated number of larvae based on their size. I’ll add a picture of the conversion sheet later, but you can see my data from the counts on the 4th sheet of my Google doc.

Larval Counts Data Sheet

I screened each of my 5 tanks over 200, 160, and 100 screens. Since I’m not weighing every size class possible, I calculated a range for # of larvae- assuming that my 160 sample had larvae sized between 160 and 180 and my 100 sample had larvae between 100 and 150.

After counting, I threw out larvae from SS_Tank1 that was left on the 100 micron screen as these are mostly dead or sick.

When doing drop counts, I accidentally knocked over a tray with SS_Tank2 160, 100 and NF_Tank1 100 on it. I had already put the larvae back in their tanks, so except for weight counts I don’t have data for those.

Comparing the weight counts to drop counts, the weight counts usually overestimate compared to drop counts- particularly if there are less than 10,000 larvae.

Some Chagra (a diatom) is now available.

Wednesday 7-1-15

Steven Roberts came in to help today. I did all of the filtering of larval catch buckets, and he did all of the counts, helped with bucket cleaning, and helped me set up a growth rate experiment.

Checking for larvae:

  • Some: NF3, NF5, SS3, HC1,HC5
  • Lots: SS1,NF2,SS4
  • Filtered out all buckets
  • These were counted almost immediately after filtering out, but still saw a higher proportion of dead larvae in buckets than usual.

Growth rate experiment

As multiple families in each group spawned today, I started an experiment to look at differences in growth rate. Silos with 100 micron screens are put in beakers filled with seawater and algae. I have 3 replicate silo/beakers for each population, with ~900 larvae added inside of the silo.

Growth Rate Experiment

Cheesin’ hard next to the growth rate experiment

This is referred to as a “static system”, as water is not flowing through it. Every day I have to rinse off the silo and transfer it to a clean beaker filled with seawater/algae. I’m taking samples of the water that goes into the beakers and samples after the larvae have been in there for a day to get an idea of feeding rate. To calculate the algal cell density in these samples, I put 10 mL into a 15 mL centrifuge tube and spin them down. I then take out the seawater and add some back so that they are all at a volume of .5mL. With this concentrated sample, I can use a hemocytometer to count algal cells and scale up to estimate the density in my original samples.

Notes: algae is 50% Tiso, 50% Chagra

Hemocytometer

Hemocytometer

Thursday 7-2-15

On Wednesday night I bought a 24″ x 40″ sheet of PVC to make into 4″ x4″ tiles for oysters to settle on, as they were close to holding on a 224 micron screen on Tuesday (meaning they were almost large enough to be at the metamorphosis stage). However, when I got to the hatchery on Thursday there was no one around to show me how to cut it up. I noticed some larvae were piled up on the bottom of the SS_Tank1 and HC_Tank1. I thought this might be due to them being ready to settle. With the tiles not ready, I rigged a settlement system using large silos and cultch (ground up bits of shell around 450 microns in size). This system is static, like the growth rate experiment, so I have to change the water every other day and add in algae every day. I added 6,000 SS_Tank1 larvae to each of two silos.

I filtered out all of the 100L larval tanks over 224, 160, and 100 screens except for NF_Tank1 where I just did 160 and 100 (ran out of time). Based on the age of the HC_Tank1 and SS_Tank1 tanks, there shouldn’t be any screening below 160, but a significant number were in both cases. The larvae at all sizes did not move very much, even without ethanol added to the well. I took 20,000 of the SS_Tank1 larvae that were filtered on the 224 screen and added them to my settlement system. Not knowing what to do with the rest, I added them back into the SS_Tank1 tank. I threw out the 100 micron sized SS_Tank1 and HC_Tank1 larvae though.

Growth Experiment

Notes:

  • salinity: 29 ppt
  • temp: 20C
  • Food: Tiso, 609

Larval Count Sheet

Monday 7-6-15

Today I talked to Ryan, the hatchery manager, a little bit about the possible causes of the mortality I saw over the weekend. In his experience, mortality events are not uncommon when rearing larvae in the summer. He thinks it may have to do with water quality issues (which I can’t really do anything about). He recommended I be more vigilant about culling dead or sick larvae from the tanks.

As I emptied out HC_Tank1 and SS_Tank1 over the weekend to be cleaned for new larvae, I adopted a new labelling scheme on my data sheet. The first attempts in a tank have the letter “a” (i.e. HC_Tank1a, SS_Tank1b) and after totally culling or emptying out a tank of larvae I change the letter in the tank name. I’ll see how this scheme works when it comes to actual data analysis, but for now it’s helping with data entry and keeping track of how many are in a tank at a time.

SS_Tank2a smelled disgusting and had sheets of larvae on the bottom so was also emptied out entirely. For my NF and HC larvae, I made a “sick” tank and a “new” tank. In the sick tanks (NF_Tank1b and HC_Tank2b), I added back larvae that were swimming in the beaker after being filtered out from the tanks. I may have to throw out these too, but until I have too many larvae for the new tanks I figured I would try to save them.

Filtering out larvae tanks:

  • HC_Tank2a (160,120) -> swimmers into HC_Tank2sick
  • HC_Tank1 (100) -> swimmers into HC_Tank2sick
  • NF_Tank2a (160,120,100) -> swimmers into NF_Tank2a
  • NF_Tank1 (100) -> swimmers into NF_Tank1sick

Checking for larvae:

  • Lots: NF1,HC1,HC2,HC4,HC5,SS4,SS5
  • Some: HC3

Set up growth rate experiment #2:

Since I had lots of larvae from every family, I decided to start another growth rate experiment. I combined the larvae from all families in a population, took samples for DNA, and added 1,100 larvae to 3 replicate silos.

Friday 7/3/15 and Sunday 7/5/15

Friday 7/3/15

My fiancé Michael Alcorn had a vacation day, so was able to come out to the hatchery and finally find out what I do all day. He did some of the counts, but quickly found those to a bit too mind numbing for him so mostly helped out with cleaning. We also tried to cut up the PVC sheet into tiles, but weren’t able to do so with the tools in the hatchery workshop.

Became really concerned about the mortality going on in the 100L larval tanks, as all of them had a lot of larvae sitting on the bottom. With it being the holiday weekend, most of the hatchery staff were out of town so I didn’t have much advice on how to proceed. One of the hatchery technicians, Alice, said they were having mass mortality with their larvae as well making me think it was a hatchery-wide water quality issue. She gave me some advice to minimize time the larvae are spent in small containers or out of the water which I started implementing. I decided for today to empty out the HC1 larval tank, do counts for the different size classes, save some for a DNA sample, and then put all the newly spawned HC larvae in that tank.

Piles of dead oyster larvae on the bottom of the tanka

Piles of dead oyster larvae on the bottom of the SS2 larval tank

Checking for larvae

  • Lots: HC5, NF2, HC3
  • Some: HC2, HC1

Larval counts

Notes on growth experiment

  • CGW/Tiso/Chagra
  • 19degC
  • 30 ppt salinity

Sunday 7/5/15

I filtered out SS_Tank1, NF_Tank1, and HC_Tank2 over various sizes to look at how far they grew and the estimated mortality. I did not put SS_Tank1 larvae back in the tank, and instead cleaned it out for the newly spawned larvae. I also changed the water on my growth experiment, filtered out new larvae, and changed my banjo filters.

Checking for larvae

  • Some: SS4,NF5,HC5
  • Lots: HC2,HC3,HC4,SS5

Notes on growth experiment

  • CGW/Chagra
  • 19degC
  • 29 ppt salinity

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.

20150615_133654

*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).

http://www.nwfsc.noaa.gov/news/features/hatchery/

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!)

Advancement to Candidacy

For the past two years, my email signature has read “Graduate Student” instead of the slightly more illustrious “Doctoral Candidate”. Every program has slightly different hoops to jump through in order to reach this distinction. In the Darwinian Sciences at UChicago, it’s a Dissertation Proposal Hearing- where one tries to convince their faculty committee that they are capable of carrying out a 3+ year independent research project. Students form these committees themselves by choosing faculty with expertise pertaining to their dissertation research, so that they may act as resources as well as qualified judges.

Check out my written proposal and the presentation I gave on May 21- I’d love comments on either!

Proposal hearing

Deciding WTF To Do (Nov. 2014)

This was a stressful time in my graduate school career. I felt torn by indecision about what the best wet lab method was to get the data I wanted- given the fact I had very little research funds. When I wrote a few grant proposals in Spring 2014, I had chosen to do the ezRAD method with pooling of 20 individuals from a site per library with one unique barcode per library. As the name suggests, this method is technically straightforward as it uses standard Illumina TruSeq preparation kits, with additional benefits of eliminating PCR-induced bias and not requiring sonication. I had enough money for one kit of 24 barcodes and 1 lane of Illumina HiSeq sequencing. Perfect! But in October, I met with a couple faculty members (one doing plant phylogeograhy using high throughput sequencing, the other a bioinformatics/population genomics guy) and they strongly discouraged against the pooling idea. While there was some support in the literature (Molecular Ecology 2013 Gautier) for the ability to get accurate allele frequencies from pooled data, numerous other papers (such as Molecular Ecology 2014 Anderson) cast doubt. By pooling individuals, I would limit the information I could glean from my sequencing data (ie observed heterozygosity or any form of haplotype analysis) and also be making an a priori assumption that the sites I collected were indeed separate populations.

Alright, so pooling was out. But if I could barely afford 24 barcodes, how could I possibly afford enough unique barcodes in order to sequence 96 individuals on a lane?? Fortunately, a curator at the Field Museum offered to share her lab’s Genotype-by-Sequencing (Elshire-2011-A Robust, Simple Gen) adaptors and barcodes for free. The only problem was they only had 48 barcodes, and my goal was to sequence at least 96 individuals per lane (each lane costs $1100-$1800). This act of scientific kindness led me down another path of obsessive pros and cons lists. Numerous grad students, postdocs, and professors (some I had never met and only stalked on the internet) kindly put up with my frantic emails as I tried to figure out wtf to do. Long story short, I decided to accept the 48 GBS barcodes and use a combinatorial index approach as in Double Digest RADSeq. Excessive pros/cons lists attached.

Cost of Pooling with Different Methods

Method:
  1. Make 40 libraries, with 2 for each population each containing 20 individuals. Sequence on 1 lane. Then resequence individuals from a subset (ie 8 pops) on another lane, will get more loci and better coverage. Can be pops that were not sequenced well previously or pops of interest.
    1. Use ApeKI and GBS
    2. Use ezRAD and REs of choice
    3. ddRAD
  2. Make 40 libraries, with 2 for each population each containing 20 individuals. Sequence on 1 lane. Resequence 8 pools that were poorly sequenced or of greater interest. OR sequence 20 each lane.
    1. Use ApeKI and GBS.
    2. Use ezRAD and REs of choice
    3. ddRAD
GBS (vs ezRAD)
Pros
Cons
$2000 cheaper
uneven sequencing of loci: may need to throw out loci. 2nd run def required
Advice on protocol
Not sensitive to methylation
blocked by some CpG methylation
PCR bias
~65,000 fragments
PCR cleanup vs Ampure beads
1a)40, then 96 GBS
Item
Estimated Cost
Sequencing
2200 (if joined with another group, otherwise 3600)
Adapters
~$200 for Y
RE
~$168
Ligase
80-200
PCR purification kit
$220-$524
Bionalyzer
$1088
Total
$3956-$5780
1b)40, 96 ezRAD
Item
Estimated Cost
Sequencing
3600
Adapters
2880
RE
236
PCR purification kit
0
Bionalyzer
$1088
Total
$7104

1c) and d) 40(48) to 40+96 ddRAD

Item
Estimated Cost
Sequencing
3600
Adapters
$4350
RE
$300
Ligase
80-$201
Ampure beads
$945
Pippin Prep
40-120
Bionalyzer
320-$1088
Total
9636-10,604
Not Pooling:
 
Method:
  1. Use GBS with ApeKI.
    1. 96 libraries of individuals using GBS with ApeKI on one lane. 5 from each of 19 populations or 6 from 16 pops. Look at sequencing, then do another lane with 96 individuals.
    2. 48 on two lanes (so as not to fiddle with adaptors).  ~7 from 13 populations.
  2. 96 libraries of individuals on one lane using ezRAD. 5 from 19 populations or 6 from 16.
    1. Adjust sequencing of additional individuals/pops
  3. 96 on one lane using ddRAD. Resequence additional on 2nd lane.
1a) 96 then 96 GBS
Item
Estimated Cost
Sequencing
1800-3600
Adapters
$200
RE
0-150
Ligase
80-$201
PCR Cleanup
330-524
Bionalyzer
$768-$1536
Total
$3178($33/96)-6211($32/192)
1b) 96 then 96 GBS (real)
Item
Estimated Cost
Sequencing
3600
Primers
$200
RE
$256
Ligase
$256
PCR Cleanup
$279-$389
Bionalyzer
$16
NEB Taq 2X Master Mix
$56
Total
$4663-4773 ($24.5/192)
2)96 then 96 ezRAD
Item
Estimated Cost
Sequencing
1800-3600
Adapters
$2880
RE
$150-300
Bionalyzer
$768-$1536 (12($96)-24($192)
qPCR
72-144
Total
5000($52/96)-7116($37/192)
3) 96 then 96 ddRAD
Item
Estimated Cost
Sequencing
1800-3600
Adapters
$4350
RE
$300
Ligase
80-$201
Ampure beads
$945
Pippin Prep
80-160
Bionalyzer
$768-$1536
Total
8323-11,092($86/96-$57/192)

Sticks in the Mud

As mentioned previously, one of the aims for this project was to compare allele frequencies between life stages of the Olympia oyster to see if there are any significant shifts and at which loci. These shifts could be due to stochastic processes, such as Sweepstakes Reproductive Success, or strong selection during a life stage that causes high mortality for individuals with certain alleles. Discriminating between the two is veryyyy difficult. As this field season is exploratory, I designed an experiment to determine if allele frequencies might indeed be shifting between life stages and whether this varies by geographic location and environmental conditions.

Larval Olympia oysters preferentially settle on oyster shells, so I made sticks with shells on them to place in various locations at the beginning of my field season. Olys (the affectionate term for Olympia oysters) start spawning as early as mid June and can go until September. After settling, the spat is about 0.5mm in size and under the microscope faintly resembles a contact lens. At the end of my field season I planned to gather some of the oyster sticks and carefully scrape off the spat for subsequent genetic analysis. The remaining oyster sticks would be left to allow the spat to mature to juveniles and then sampled at a later date.

Drilling shells

Drilling holes into Pacific oyster shells…somehow I finished with all 10 fingers still intact…

 

 

Shell sticks after being left in the water for almost a year

Shell sticks after being left in the water for almost a year.

Due to Jennifer Ruesink’s longterm monitoring of larval settlement in the area and recent characterization of pH variability, I chose Willapa Bay as one of the geographic regions for this project. The pH in the southeast part of the bay remains relatively stable at 8 during the summer, while the pH in the north part of the bay can range between 7.6 to 8.6 (a huge difference when you consider the pH scale only goes to 14!!). This difference in variability between the sites is likely due to the Willapa River input in the north and other oceanographic features of the bay. To see if these environmental differences affect allele frequency shifts, I decided to place shell substrates in both areas. While setting out oyster sticks in the southeast was easy, the olys in the north part tend to settle on substrate 30 feet underwater. To address this, Alan (introduced here) helped me construct the concrete version of shell sticks, because apparently oly larvae are not very discriminating between shell and cinder blocks. The blocks were tied to buoys and then deployed at a later date by Alan to be picked up again sometime in October.

The other region I chose was Puget Sound, WA, which also demonstrates environmental heterogeneity between different subregions. Dr. Brent Vadopolas and others at the University of Washington are doing some really interesting work looking for signals of local adaptation in 3 populations of Olympia oysters by outplanting oysters from each population to 4 locations around Puget Sound. During the course of this study, they observed extreme mortality in one location (Dabob Bay) for all transplanted populations. Additional observations from the region indicate that natural populations of olys in south Puget Sound are doing well, while those in the north are starting to grow despite previously low numbers. Some questions these observations raise: was the mortality event stochastic, or was it due to consistent ecological factors in the area? If oysters are locally adapted to their specific subregion of Puget Sound, where at the northern oysters recruiting from? To tie in my project with theirs, I placed two shell sticks next to their experiment in the North Sound and two near the South Sound location. Stephanie Valdez, a recent graduate of UW, is monitoring Olympia and Pacific oyster settlement in Hood Canal (body of water that offshoots Puget Sound) and agreed to give me the oly recruits from her Dabob Bay shell sticks for genetic analysis.

Locations of Puget Sound shell sticks

Locations of Puget Sound shell sticks

Sounds like a great plan, right? As I’m writing this post facto, I can say that this aspect of my field season was essentially a failure from the get go, at least with the Puget Sound shell sticks. First, I was unable to find the oyster transplant site in South Sound due to a miscommunication about the GPS coordinates and my own stubbornness about asking for clarification. Instead, I decided to find my own spot in the vicinity to put my shell sticks. The area of the sound I was in, Oyster Bay, is surrounded by private homes with “No Trespassing” signs and actual commercial oyster farms. After an unsuccessful attempt to ask a homeowner for permission to access the bay from their backyard, I gave up on that idea and decided to park at a wildlife viewing area near the tip of the estuary and cross the (deceptively) short distance to the water’s edge. It took an hour of slogging through the mud, losing my boots, being laughed at by raccoons, and eventually breaking one of my shell sticks for me to accept defeat.

10463887_10152988725794128_5124704887406444502_n

That night, after a campground shower and a couple IPAs, I found the resolve to try again the next day. This time, instead of directly cutting across the soul-crushing mud to the water, I decided to walk along the rocky edge of the bay until I reached an area that would remain underwater even during minus tides.

Oyster Bay

Green: path to stick placement
Red: path back to car

This Google Earth image was obviously taken during a normal tide, because when I trekked out there with a 1.3 ft low tide the water didn’t reach nearly as far inland. From the car to the shell stick placement site was about 1.2 miles of sweating, slipping, cursing, and  overall misery. To avoid such a return trek, I cut through someone’s backyard and walked 2 miles on the backroads- no doubt looking like the creature from the black lagoon. The placement in North Puget Sound was considerably easier. I quickly found the UW oyster transplant site and wedged my shell sticks about 50 ft away.

Fast forward 5 weeks to the beginning of August. By that time, I had finally internalized the value of asking for help and simply drove up to the house in Oyster Bay I had previously trespassed by and asked if I could walk through their backyard to the bay. They found the request strange, but mildly amusing and let me go by. When I get to the coordinates for my shell sticks, however, they were gone. Both of them. I frantically waded around the entire area to no avail. Talking to Stephanie later, she said that in her work shell sticks do occasionally go missing. Maybe a nearby oyster farmer saw the pink flag tape sticking out of the water and thought it was trash. Maybe I didn’t hammer them in deep enough and they washed out with the tides. Maybe otters came and made a shell necklace out of them. Whatever the cause, by not having one of the sites it took away any interesting comparisons I could do with the others in the region so I decided to focus on my phylogeographic collections for the remainder of my field season. Also, I still have hope to get something interesting out of the recruits I’ve collected from Willapa Bay.

While totally a bummer, the entire experience taught me a lot about field work, experimental design, and myself.

  1. When setting out an experiment, talk to all of the nearby landowners and workers so they know to keep an eye out and not disturb anything. They may even be interested in the science!
  2. Don’t put all of your eggs in one basket, i.e. put your shell sticks in multiple sites for a sampling area.
  3. Ask for help! You don’t get any points for being stubborn and trying to work something out alone. In fact, you’re more likely to get covered in mud.

 

See You in Seiku

I’m going to be a little anachronistic for this next post, because I’m waiting on GPS coordinates to make a pretty map for my last couple of days in Willapa. So fast forward to the evening of June 29….

WBtoSekui

 

Off Highway 101 near Kalaloch, WA

Off Highway 101 near Kalaloch, WA

After a beautiful 3 hour drive along the Olympic peninsula, I arrived late at the house of my advisor Cathy Pfister and her husband Tim Wootton, another ecologist and faculty at UChicago. For most of the year they live in Chicago, but during the summer stay out in Seiku, WA so that they are much closer to their primary field site, Tatoosh Island.

tatoosh_aerial

A tiny seaside community of 27 year-round residents, where cell service was negligible and kids were more likely to be running around outside than sitting in front of a TV, Seiku had the feeling of quintessential Small Town America without all the commercial trappings. Being a 5 minute walk from the Pacific wasn’t too bad either. My purpose for this visit was not particularly oyster related, but instead to catch up with Cathy and explore some of her field sites around the area to see if a particular organism, ecosystem, or potential research question tickled my fancy. While oysters are certainly my species du jour, it’s a good idea at this stage of my PhD to keep an open mind. Choosing a thesis project requires discovering a relatively unfilled niche so that other researchers aren’t scooping your findings and publishing them before you, making all the hard work seem for naught!

The next morning, Cathy drew me out directions to Slip Point, a rocky intertidal habitat a few miles away.

Slip Point

Armed with pH and dissolved oxygen sensors, a waterproof notebook, and a rain bib I spent a few hours poking around in tide pools and climbing over slippery rocks. Having previously lived in Texas and Florida, I had never actually seen a Pacific Northwest rocky intertidal habitat in person before. Similar to those I was familiar with on the east coast, there was a very visible gradient in different species leading away from the water. This follows the gradient of stresses an inhabiting organism might be subjected to, from being constantly submerged by water in the subtidal zone to experiencing huge, daily fluctuations in salinity, temperature, and desiccation in the upper intertidal. Tide pools dotting the shore offer respite from drying out, but even adjacent tide pools can vary drastically in species composition and environmental factors.

http://sky.scnu.edu.cn/life/class/ecology/chapter/Chapter3.htm

Intertidal zonation

What struck me the most was the diversity of seaweeds covering the rocks (struck being the operative word as I slipped and fell on my ass because of them quite a few times).

seaweed

While I may know more at this point about oysters than my adviser, she is much more familiar with the ecology of kelp in the PNW than me. I can’t even tell my reds from my greens! Despite that I was intrigued about the possibility of local adaptation in these sessile species, for in many ways oysters are similar to some plants in the way they produce large numbers of offspring and release them into the water/air with a small chance of finding suitable habitat to settle. Another PhD student in my lab, Courtney Stepien, is doing a lot of work in this area and I resolved to talk to her more about her research once I returned to Chicago.