Kaitlyn’s Notebook: Differential Protein Expression

I’ve been trying to identify diferentially expressed proteins in Rhonda’s oyster data, however there has been significant difficulty finding a way to compare the different treatments or days. Initially I would take the differences between the proteins however this did not consider the majority of proteins in the ABACUS data which means I could be missing important enriched genes. Furthermore, finding the differences between two days was simple when looking at the differences, but impossible when comparing all 13 days with each treatment.

Working with Sean, we tried a PCA plot and K means clustering however neither worked effectively. Yaamini is using MSstats to analyze her Skyline data. I am not sure if ABACUS data can be analyzed using MSstats but I know MSstats can analyze DDA so I am going to look into it further.

I helped Yaamini label some tubes for her and Laura’s DNR a little today as well!

Laura’s Notebook: May 29th & 30th, at Manchester

May 29th, 2017

  • Collected and counted larva today; not too many new.
  • Steven imaged plate #1 & #2 in my backed-up well plates.
  • Setting tank location is TBD- Stuart will discuss with Ryan to see if I can use one of the tanks inside the hatchery, as it is much cleaner and easier to feed.
  • Sampled flow from drippers on larval table and used hemocytometer to count algae. Average concentration is ~113k cells/mL, which is slightly above the 100k cells/mL target, as outlined in the FAO manual. None are below the target. Dosing rate when I sampled was 70, so I reduced to 65. Concentration will, however, depend on how concentrated the algae is in the header tank.


May 30th, 2017

Screened & counted larval buckets today.

Began by screening through 180um (started with bucket #16, SN-6 Ambient). Nothing held on a 180, and the larvae did not look good. I screened through another group, and saw similar poor quality. It became obvious that I needed to separate live from dead. I began setting up a double-bucket system, where the “swimmers” flow into a small bucket w/ banjo, with the assumption that the dead oysters will remain at the bottom of the first bucket. I tested this method a few weeks ago, using the 26L/hr drippers and waiting 6 hours, but there were still many live larvae remaining. I’m hoping that if I increased the time considerably, I will have better success. So, instead of screening through 180 & 100 today I screened through a 224 to remove debris, onto a 100um, rinsed the larvae with FSW, let soak in fresh FSW for a few minutes while I cleaned, then sampled for counts, and returned ALL to the cleaned 5gal bucket, but with the banjo removed, the outflow directed into a 2-gal bucket with a banjo. I installed air stones in both the 5-gal and the 2-gal. I used the 8L/hr drippers. Tomorrow, I will collect the live larvae, clean the 5-gal bucket of the dead larvae (inspecting it to confirm 100% mortality), then return the live to the 5gal. If this method of culling is successful, I will incorporate it into the weekly schedule moving forward.

Ryan provided some insight into potential causes: This time of year is notorious for die-offs due to extreme fluctuations in dissolved gases from algae blooms & crashes. High mortality can create a breeding ground for ciliates. I saw lots of ciliates in my poorest looking samples. Also, many of the SN & NF groups have looked lazy.

Interestingly, the SN & NF groups, which are all being reared on the small table against the wall, had much fewer live as compared to the K & HL groups, which are being reared on the center table. Overall average %Live for the larval rearing table was 43%, and for the center table it was 68%. The K & HL groups also appeared much more active, swimming and feeding. Because of the apparent poor conditions on the larval table, I’m going to move all larvae to the middle table tomorrow. I’ll move the K and HL broodstock to the small table.


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Yaamini’s Notebook: DNR BCA Assay Round 3


Once again, Jose and I are using the protocol generated in December to quantify the protein concentrations my samples and Laura’s samples. We’ll also quantify the protein in Rhonda’s larval geoduck samples. It’s a lot to keep track of! The end goal is to obtain the µL of protein required for 100 µg of protein in our samples, and the corresponding amount of 50 mM NH4HCO3 in 6M urea.

Step 1: Prepare reagents for 90-100 samples

Step 2: Plan microplate arrangement

Figure 1. Microplate arrangement

Step 3: Pipet 10 µL of either standard or sample into the corresponding microplate well

Step 4: Prepare BCA working reagent

The BCA working reagent should only be prepared right before plate incubation and reading.

  • ((8 standards x 5 plates) + 100 samples) x (3 replicates each) x (200 µl of working reagent per well) = 104,000 µL working reagent = 104 mL working reagent
  • Used 110 mL Reagent A and 220 µL Reagent B to make a 50:1 Reagent A to B ratio.

Step 5: Added BCA working reagent to each well

Figure 2. Microplate 1

Figure 3. Microplate 2

Figure 4. Microplate 3

Figure 5. Microplate 4

Figure 6. Microplate 5

Step 6: Used Genome Sciences incubator and plate reader

Step 7: Calculated volume of protein needed for 100 µg per sample

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Grace’s Notebook: May 30, 2017

Today I helped with Yaamini and Laura’s DNR sampling.

We started out the day by halving tissue (I worked on Laura’s Geoduck, while Yaamini worked on her oysters) in order to save some for future use if need be.

The samples then went through sonication after they had ____ added and were vortexed.

The samples I used today were:

Screen Shot 2017-05-30 at 2.25.30 PM



Yaamini’s Notebook: DNR Sample Preparation and Sonication Round 3

Here’s to hoping this is the last round of DNR sample preparation!

The plan for the week:

  • Tuesday (that’s today!): Prepare tissue and sonicate with Grace
  • Wednesday: BCA analysis all by myself
  • Thursday: Mini-trypsin digestion with Jose and Kaitlyn
  • Friday, Saturday and Sunday: Speed vacuum all samples. The speed vacuum can hold a maximum of 60 samples, so Laura, Jose and I all have to speed vacuum our samples separately.
  • Monday: Desalt all samples
  • July: Mass spec this shit :sunglasses:

This is what Grace and I did today:

Step 1: Label centrifuge tubes

  • Tubes for the sample I would extract proteins from (“1/2”)
  • Tubes after sonication (“11 µL”)

Step 2: Obtain samples for extraction from the -80ºC freezer.

Table 1. Proteins to extract, based on table from a previous lab notebook entry.

Site Condition 1 2 3
PG B N/A O51 O52
PG E O78 O56 O30
FB B O43 O40 O35
FB E N/A O24 O49
WB B N/A O121 O122
WB E N/A O131 O144
SK B O99 O96 O113
SK E O106 O102 O91
CI B N/A O21 O22
CI E O10 O06 O04

Step 3: Cut samples in half.

  • Obtained dry ice from Biochemistry J Wing
  • Placed sample centrifuge tubes (original vials and vials labelled “prot”) in dry ice to keep them from thawing
  • Created a 10% bleach solution (4 mL Clorox bleach in 40 mL nanopure water) to sanitize equipment
  • Obtained tweezers, weigh boats and razor blades
  • Placed weigh boat in dry ice to keep chill
  • Using tweezers, removed gill tissue from centrifuge tube and placed in weigh boat
  • Cut gill tissue in half with razor blade
  • Disposed of razor blade
  • Placed one gill tissue half back in original tube, the other in the tube labelled “1/2”
  • Put both tubes back in dry ice to stay cold
  • Tweezers cleaned by dipping in 10% bleach solution, and then rinsing in nanopure water
  • Repeated for all samples

Step 4: Made 50 mM NH4HCO3 + 6M urea solution

Protocol used can be found here, or viewed below. This solution must be used no later than 24 hours after it is made, or it is no longer viable.

  • Measured 10 mL of nanopure water in a graduated cylinder, and poured into falcon tube
  • Weighed out 79.06 mg of ammonium bicarbonate (NH4HCO3) (0.0793g measured)
  • Added NH4HCO3 to falcon tube, vortexed until mixed
  • Weighed out 7.21g Urea (7.21g measured)
  • Added Urea to falcon tube, vortexed until mixed
  • Poured falcon tube contents into graduated cylinder
  • Topped of contents in graduated cylinder with nanopure water up to 20 mL
  • Poured contents of graduated cylinder into falcon tube

Step 5: Added 50 mM NH4HCO3 + 6M urea solution to samples

  • Moved samples to wet ice. Samples remained in wet ice for the duration of the sonication process.
  • Pipetted 100 µL 50 mM NH4HCO3 + 6M urea solution into one sample tube
  • Repeated for all samples + 1 blank

Step 6: Homogenized samples

  • An up-and-down motion worked better than a side-to-side motion (pounding > smushing)
  • Homogenized as much as possible
  • Resulting mixture was cloudy, difficult to see tissue fragments
  • Repeated for all samples + 1 blank

Step 7: Vortexed samples

  • 3 pulses on vortex set to speed 10 (maximum speed)
  • Repeated for all samples + 1 blank

Step 8: Sonication preparation

  • Filled containers with 1000 mL nanopure water each and labelled containers “1” and “2”
  • Made an ethanol dry ice bath
  • Filled a beaker with ethanol
  • Added one piece of dry ice at a time
  • Continually added ice over the course of sonication
  • Ensured all samples needing sonication were in a wet ice bath
  • Cleaned sonicator tip
  • Dipped in ethanol for 5 seconds while on
  • Dipped in first container of nanopure water (1) for 5 seconds while on
  • Dipped in second container of nanopure water (2) for 5 seconds while on
  • Wiped down with additional ethanol and a kim wipe

Step 9: Sonication

  • Sonicated one sample
  • Placed sonicator tip in one sample centrifuge tube for 10 seconds
  • Immediately placed sample in ethanol dry ice bath for 5 seconds
  • Moved sample to wet ice bath
  • Cleaned sonicator tip using procedure explained above
  • Repeated for the rest of the samples sequentially, and the blank
  • Repeated sonication procedure for all samples and a blank 3 times total
  • Sonication must proceed sequentially (one sample sonicated, then another sonicated)
  • Cannot sonicate one sample immediately after it has already been sonicated

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Yaamini’s Notebook: DNR Mini-Trypsin Digestion Round 2

I need to become a faster pipetter.

If there were a pipetting Olympics, I wouldn’t come in last. But I wouldn’t come in first either, or even medal. This was slightly problematic when pipetting every few minutes for the trypsin digestion. It took a while, but we finished!

Last time we performed a digestion, our quantities were calculated based on having 30 µg protein/100 µL. This time, we had 100 µg protein/100 µL. Therefore, we used the quantities of reagents specified in the original protocol.

The first thing Jose and I did were set up heating blocks at 37ºC. Because the heating block in the Roberts Lab did not have the capacity to hold all of our samples, Jose brought one from his lab. We placed additional thermometers in the heating blocks to ensure the temperature was what we wanted.

Based on my calculations for Plate 1 and Plate 2, I pipetted the correct amount of sample and 50mM NH4HCO3 + 6M urea into newly labelled tubes (ex. OBLNK D). Then, I added 6.6 µL of 1.5 M Tris pH 8.8 and 2.2 200 mM TCEP and vortexed the tubes. For our 51 samples, we needed 2-200 µL aliquots of TCEP from the -80ºC freezer. At this point, I went to class. Jose tested the pH of our samples to ensure the pH was still basic, than began the digestion in the heating blocks for one hour.


Figure 1. pH strips for samples. The blue color of the strips indicates the sample pH was still basic.


Figure 2. First heating block set-up.


Figure 3. Temperature confirmation for first heating block.


Figure 4. Second heating block set-up.


Figure 5. Temperature confirmation for second heating block.

After the digestion, I got 6-200 µL of IAA from the -80ºC freezer and covered them with aluminum foil. We added 20µL of IAA to each sample and vortexed the tubes. We covered all of our samples with aluminum foil and let them digest for one hour. Halfway through the digestion set-up, I left for Merril’s Ph.D defense. When I was gone, Jose added 20 µL of DTT to each of our samples, vortexed, and started the digestion. I got back in time to add 1.65 µL of LysC to each sample and started the one hour digestion.

Because Jose needed to leave early, he prepared 25 mM NH4HCO3 solution for all of our samples. After the digestion was over, I added 800 µL of 25 mM NH4HCO3 and 200 µL of HPLC grade methanol to each sample and vortexed them. I then needed to add 3.3 µL of Trypsin to each sample, meaning I needed 9 bottles of 20 µg Trypsin. We only had five bottles, so I needed to borrow 4 bottles from Genome Sciences.

Before using each bottle, I added 20 µL of nanopure water and vortexed. This made a 20 µg/20 µL solution, allowing me to add 3.3 µL of Trypsin to six samples/bottle. After adding Tryspin to all oyster and geoduck samples, I vortexed the tubes and let them sit overnight. I started the digestion at 4 p.m. on Friday evening, and stopped it at 10 a.m. Saturday morning.

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Yaamini’s Notebook: DNR BCA Assay Round 2

We have lots of protein!

Using the protocol we generated in December, Jose and I quantified the amount of protein in my oyster samples and Laura’s geoduck samples. Because we used 100 µL 50 mM NH4HCO3 + 6M urea this time, our protein concentrations were generally much higher.

The first thing we did was prepare reagents. Based on the recipes in Rhonda’s original protocol, we made enough 30 mL of 50 mM NH4HCO3 and 6.6 mL of Lysis Buffer. We also made one set of standard solution vials.

Because there were 51 samples total including blanks, we needed three separate microplates.


Figure 1. Microplate arrangement.

To make each plate, Jose first pipetted each standard in triplicate in the first microplate. I added 22 µL of 50 mM NH4HCO3 to each sample vial and vortexed the solution. Jose then pipetted 10 µL of the sample vial contents into the corresponding microplate wells. The only sample that was not run in triplicate was GBLNK. At this point, I needed to get to class. We covered each micrplate with parafilm and placed it in the fridge. The plates were in the fridge from about 9:30 a.m./10:00 a.m. until 1:30 p.m. Right before I came back from class, Jose made the BCA Working Reagent and used a multichannel pipet to add 200 µL of the reagent to each microplate well.


Figure 2. Microplate 1.


Figure 3. Microplate 2.


Figure 4. Microplate 3.

We took our completed microplates to the Genome Sciences Building. Microplate 3 was placed in the Varioskan Flash plate reader. It was incubated at 37 ºC for 30 minutes, shaken, then read at 562 nm. While microplate 3 ran, I incubated Microplate 2 in a separate plate incubator, similar to the one used in December. I kept it in there for 30 minutes at 37 ºC. When it was finished, I took it out and placed Microplate 1 in the incubator. Microplate 2 was out for five minutes before I could place it in the plate reader. In the Varioskan Flash plate reader, Microplate 2 was shaken, then read at 562 nm. Microplate 1 was also incubated at 37ºC for 30 minutes. The incubation period finished after Microplate 2 was read, so Microplate 1 went directly from the incubator to the plate reader. After being shaken, it was read at 562 nm. I sent Laura the data she needed for calculations and kept the data I needed.

Here’s what I used to calculate the concentration of protein in each well: Plate 1 data

Plate 2 data

I used formulas from my previous calculations to make spreadsheets to analyze my BCA data. The final results can be found below, including the amount of protein and 50 mM NH4HCO3 + 6M urea needed for trypsin digestion.

Plate 1 calculations


Plate 2 calculations


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Laura’s Notebook: May 27th, 2017

Here are some charts of the Oly larvae that I’ve collected & counted to date



Some documentation

Arrival inspection

Larval buckets:

  • Test bucket Check:
    • Not overflowing, despite not changing banjo yesterday.
    • T = 17.25 (T1)
    • pH = 7.88 (Durafet 2)
  • Aeration looks good
  • Algae concentration looks a bit weak – increased dosing rate from 60 -> 70
  • New larvae in following catchment buckets:
    • K-10 Low – a bit
    • HL10 Low – a ton!
    • SN-6 Low B – a lot
    • SN-10 Low B- a lot
    • NF-6 Low B – a bit
    • NF-6 Amb A- a bit
  • Maybe some in:
    • NF-10 Low B
    • NF-10 Low A
    • SN-6 Amb A
    • NF-10 Amb A
    • NF-6 Amb B

Need to buy:

  • 15 feet 1/2 pvc flex hose (for freshwater flush)
  • 1/4” tube connectors – straight
  • 1/8” tube connectors- Y’s and straight

Need to make:

  • Fresh water connection
  • Pipe with lots of holes for catch freshwater flush outflow
  • Microculch
  • Assemble setting tank silos

Tasks Today

  • Collect new larvae, stock, sample excess – DONE
    • This went smoothly, except that the banjo was left off group HL-10 low pH yesterday, so I therefore likely lost larvae. there still was quite a bit on the bottom of the bucket, but because of this likely fewer # of larvae in the bucket I added extra from today’s spawn.
  • Make freshwater connection – NOT COMPLETE, but I figured ou what I need to purchase
  • Clean top row broodstock & all components – DONE
  • Record location of top shelf broodstock on manifold – DONE
  • Change all banjos, drippers – DONE
  • Feed – DONE. used – 1/2 Ciso, 1/2 CGW
    • Increased algae capacity so I can take Sunday off:
      • 200L for larval tank
      • 200L for other tank
      • 100L for gigas
  • Image plates – NOT DONE

Discovered a flat tire on my bike, called an Uber- only $17, and he arrived within 15 minutes. Pretty sweet.

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Laura’s Notebook: May 23rd – 26th, 2017

May 23rd

First full day of screenling larval buckets to assess survival, growth, and determine stocking numbers for next 3 days. Katie Davidson helped out, as did Steven.


  • Screen larvae for different size classes, count, re-stock buckets:
    • Screen through…. 140, 160, 180 … ?
      • Decided on 2 size classes, <180, >180. Screened through 160 today on first 2 groups, and nothing held; will screen through 180 next week. Only screened onto 100 today, counted live, and used that info to re-stock buckets.
    • Sample for counts & images
    • Sample for freezer
    • Stock buckets – method depends on # surviving larvae- Replace drippers – DONE
  • Replace banjos – DONE
  • Collect new larvae, count, stock, and sample excess
  • Measure dripper rate @ larval tank – NOT DONE
  • Collect water sample for algae counts from each dripper – NOT DONE
  • Record broodstock & larvae group locations on tables/manifolds – NOT DONE

May 24th

  • Arrival inspection
  • Image yesterday’s larvae – DONE
  • Change banjos – DONE
  • Change drippers – DONE
  • Collect & Sample for counts – DONE
  • Sample rest for -80 – DONE
  • Talk to Ryan about:
    • Larvae health – help trouble shooting?
      • Larvae looked sluggish when screening yesterday; this could simply be a response to the handling, however it is markedly different from the new larvae that are collected in a similar manner Ryan said that “lazy” larvae can be a sign of supersaturation of Nitrogen or Oxygen. He suggested raising the drippers out of the bucket to allow for degassing, and increasing the aeration. I did so.
    • Setting tank
      • Ryan showed me which silos I can use in the setting tanks outside; I checked with Joth (as he may have had plans for the tanks), but he said that I could use them through the month of June.
    • PSRF and other help while I’m at FHL?
      • Stuart has agreed to oversee the project
      • Jade and Dana can help out
      • Olivia to work Mondays (1x per week)

May 25th, 2017

Arrival Inspection:

  • Larval Table:
    • Aeration did not hold overnight.
    • Banojs very dirty, likely due to use of Tetraselmis in algae cocktail. I will request that PSRF not feed my larvae Tet.
    • I can see larvae swimming in water column, however there are a significant amount at the bottom of the bucket.
  • Larvae present in:
    • K-10 Low
    • HL-10 Ambient
    • K-10 Ambient
    • NF-6 Ambient A
    • NF-6 Low B
    • NF10 Ambient A


  • Collected Larvae, sampled and counted most (not al- need to finish)
  • Sampled immediately after collecting, then stocked. 6 buckets = 1.75 hrs by mysefl.
  • Recorded location of bottom shelf broodstock and sensors on manifold
  • Cleaned! the following took ~2 hrs
  • NF & SN broodstock thoroughly, including tubes, tube conenctors
  • Replaced all drippers, banjos & air stones
  • Set up 2 “test” buckets to measure T & pH in both larval systems, and to test whether banjos will clog over 2 days.
  • Sketched out possible June/July schedule coverage

May 26th, 2017

Arrival Inspection:

  • Larval Buckets
    • Aeration looks good
    • Very few bubbles accumulated on the inner walls of buckets- elevating the drippers and increasing aeration appeared to have been effective.
    • Banjos not consisently dirty; perhaps corresponding to the # larvae in bucket? Need to measure dripper rate on manifold to confirm equal flow rates.


  • NF-6 Ambient A group is foamy- male(s) likely spawning today
  • Water color looks light; increased algae dosing rate to 120 pulses/min

Larvae present in:

  • NF-6 Low B
  • NF-6 Ambient B
  • HL-10 Ambient
  • K-10 Low
  • K-6 Low
  • HL-10 Low (ID’d in afternoon)
  • SN-10 Low B (ID’d in afternoon)


  • Screen larval buckets, count, restock – 3 hrs for 16 buckets
    • Possibly use 2 different buckets per group, if needed – this was needed for SN-10 Low, & SN-10 Ambient
    • I was pleasantly surprised at the survival rate! Despite larvae consistently pooling at the bottom of the buckets, my survival rate for the SN & NF groups appears adequate (calculations TBD). The K group, which has been consistently stocked since Friday, had poorer survival rate. Need to refine data sheet to auto-calculate stocking #’s and survival rate
  • Collect new larvae, stock in buckets – 2 hrs
    • sample excess larvae & freeze
  • Image larvae from 5/23 & 5/24 & finish counts – 30 minutes -not done.
  • Daily maintenance: – 30 minutes – done.
    • Replace banjos
    • Replace drippers
    • Spray down filters
  • Clean top row broodstock if there’s time – 1 hr – not done.

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Laura’s Notebook: May 22nd, 2017

Arrival Inspection

  • Larval table, top row:
    • Lots of bubbles on side of buckets; after speaking with PSRF this is likely a sign of supersaturated water (N or O).
    • Color very light – it appears that I have the same un-even algae distribution problem on this table.
    • Air stones not bubbling enough- I opened their valves a bit more and that worked. Seems like this is another depressurization issue. Not sure why.
    • Lots of larvae pooled on bottom of buckets- likely due to a combination of the above issues.


  • Larval table, bottom row:
    • banjos very dirty, but not clogged
    • air stones working well
    • algae concentration appears appropriate
  • Broodstock Table:
    • Things appear stable!
    • Very few bubbles on the sides of buckets
  • Larvae present in:
    • SN-10 amb B
    • NF-6 amb A
    • SN-6 low A
    • SN-6 amb B
    • NF-10 low A
    • K-6 low
    • K-10 amb

Today’s Tasks:

  • Moved all buckets that were on the top shelf of the larval table to the bottom shelf. Buckets that were on top shelf were: 11, 12, 13, 16.
    • Took photo of buckets 16 & 5 next to each other-obvious difference in algae concentration. img_8020
  • Cleaned banjos in larvae
  • Spoke with Ryan about June:
    • They do not know whether staff will be working weekends through June. If they don’t, then people w/o access cards/keys won’t have access to the lab. I asked whether, in that situation, PSRF would be able/willing to work my project on weekends. Ryan said maybe Jade or Stuart could- or perhaps one of them could run my project…
    • Ryan is going to talk to Stuart and consider whether Stuart or Jade could take the lead for my project while I’m at FHL. Will get an answer tomorrow.
  • Counted larvae collected yesterday
  • Imaged larvae collected yesterday
  • Collected new larvae, sampled, counted, stocked
    • Not too many larvae today. I’m streamlining this process, but currently takes a couple hours. Collection from today: 56999bb1-7cdc-4898-a156-e92e2c0fbaa4
  • Collected excess larvae in 2mL vials froze. Labeling scheme will be numerical:
    • Date, Oly, Vial # e.g. “5/22/2017 Oly 1-A”
    • I had saved excess larvae from 5/20 & 5/21, so I sampled those and the excess from today: fc88fe67-90d9-4733-bd5a-0f66f7f880fb
  • Determined how I should collect larval samples, and how frequently
    • I will collect larval samples daily when there are >10k excess larvae
    • Collection will occur after stocking buckets – I will save tripours with remaining larvae, then pull 2 samples each of 10k
  • Figure out how to fix the over saturation situation
    • Ryan said I could install small degassing columns on each bucket, that the water would flow through prior to entering the bucket. Hmm….
  • Figure out how to fix the algae concentration situation on the larval tank – not enough space to plumb algae input further back. Options could be:
    • Connect top and bottom manifold at other end to create a circular flow
    • Plumb in 2 pumps for each shelf

By the way, this is my banjo setup, which works very nicely in the quick-change: img_7926

To Do Tomorrow:

  • Replace drippers
  • Replace banjos
  • Measure dripper rate @ larval tank
  • Collect water sample for algae counts from each dripper – yikes!
  • Record broodstock & larvae group locations on tables/manifolds
  • Screen larvae for different size classes, count, re-stock buckets:
    • Screen through…. 140, 160, 180 … ?
    • Sample for counts & images
    • Sample for freezer
    • Stock buckets – method depends on # surviving larvae

To Do with time:

  • Make connection for freshwater – to do Thursday

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