5 22 2014 oyster bay repro

Oyster bay, WA

High mid 70s.

Participants Katie Jackson and Jake heare.

Tested for reproductive active today. Very bad news.

There has been a huge mortality event at oyster bay. Numbers for brooding, gaping, dead, and alive but not open are as follows.

1n13-16
Brood. 0
Gaping. 3
Dead 93
Alive 5

1s5-8
Brood 0
Gaping 2
Dead 68
Alive 4

1h5-8
Brood 0
Gaping 47
Dead  51
Alive. 9

Clearly dabob has survived whatever mortality event much better than north and south sounds populations. We will be sampling this site tomorrow to ensure fresh samples.

5 21 2014 Manchester repro

Did a brood check today at manchester. Found no brooders. 

Numbers as follows. 

4S5-8

Brood. 0

Gaping. 76

Dead.  3

4H13-16

Brood. 0

Gaping. 16

Dead.  2

4N13-16

Brood. 0

Gaping.  40

Dead.  3

Enjoy this picture of an eagle.

Capsized boat at Fidalgo Marina

Apparently a boat capsized roughly 500 feet from the samples at Fidalgo Bay. Depending on what leaked into the water, the size of the wake and other disturbances caused by rescue efforts. This might cause some issues with the animals. You can watch a time lapse video of the recovery effort here.

5-14 to 15-16 2014 Reproduction Work Up

5-14-2014

Manchester WA

Participants: Brent Vadopalas and Jake Heare

We retrieved the next set of trays in the dosing sequence and allowed each tray to dessicate for 45 minutes. Trays were then treated in a 10 gallon bath of 50% sea water to 50% freshwater mixed with 7 lbs of epsom salt. The tray was treated for 45 minutes, at which point it was removed and gaping animals were examined for signs of brooding. After examination the trays were then placed into a recovery tub with 100% sea water until the last tray examined had be in the recovery tub for 45 minutes. After each treatment the treatment water was replaced with fresh treatment to reduce temperature flux.  Then trays were rebuilt into a stack and hung off the dock. 

We counted the number of gaping animals and brooders for each tray. 

They are as follows:

4H1-4

brooders 0

Gaping 25

Closed 72
% Open  25.8%

4S9-12

Brooders 0

Gaping  43

Closed  55
% Open  43.9%

4N9-12

Brooders 0

Gaping 31

Closed  28
% Open 52.5%

5-15-2014

Oyster Bay WA

Participants: Katie Jackson and Jake Heare

We followed a procedure similar to that at Manchester. The difference being that the treatment water was not replaced after each treatment. We also counted the dead in each tray.

1H1-4

Brooders 0

Gaping    49

Dead      7

Closed   33
% Open  59.8%

1N5-8

Brooders 0

Gaping    46

Dead      7

Closed  14
% Open  76.7%

1S13-16

Brooders 0

Gaping     59

Dead        8

Closed    26
% Open  69.4%

5-16-2014

Fidalgo WA

Participants: Steven Roberts and Jake Heare

Same procedure as at Oyster Bay. 

2N1-4

Brood 0

Open 53

Dead  0

Closed  46
% Open  53.5%

2S13-16

Brood 0

Gaping  55

Dead   0

Closed 39
% Open  58.5%

2H5-8

Brood 0

Gaping 48

Dead  0
Closed  52
% Open   48%

DNA Extraction Attempt 2 5-12-14

Attempting to extract DNA from 5-10 mm whole oysters minus the shell. This is needed so that sequencing can occur in the future with completed extractions. Unlike the last attempt I will be switching out 95% EtOH for Isopropanol to get better extraction results.

Protocol is as follows:

Incubation for 24 hours at room temp:

1	ml	DNAzol
2.35	ul	Proteinase K (100ug/ml)
1	whole	oyster 5-10 mm
X5		Reactions

1. Pipette 1 ul DNAzol and 2.35 ul Proteinase K in tube containing oyster
2. Homogenize each oyster with disposable pestle.
3. Centrifuge briefly
4. Incubate overnight at ROOM TEMP.

Isolation:

500	ul	Isopropanol
1	ml	75% EtOH
1	ml	75% EtOH
200	ul	Sterile H2O

1. Centrifuge @ 10,000 g for 10 minutes.
2. Collect and Transfer Supernant; Discard pellet
3. Add 500 ul Isopropanol
4. Mix through inversion
5. Incubate for 1 minute at ROOM TEMP.
6. Spin down DNA precipitate
7. Remove supernant
8. Wash with 1 ml 75% EtOH
9. Spin down
10. Remove supernant
11. Wash with 1 ml 75% EtOH
12. Spin down until solid pellet
13. Elute with 200 ul Sterile H2O
14. Quant.

Spawning Oly Predictions

After looking through the new temp data I pulled off the temp loggers yesterday I notice a few trends that have lead to some predictions as to which site will spawn first.

Oyster Bay

Temps are trending upward to the 12.5 C mark that the animals require to begin spawning. It seems that they will have hit that temperature this week and will possibly begin spawning next week. The forecast for next week is also encouraging with the daily ambient air temps to be 75 or higher. This will bring these animals to fruitition. The only problem being that it will be during a spring tide which is not good for spawning. 

If they don't spawn next week it will occur during the following neap tide during the second to last week in May.

Manchester

While these animals have experienced a warming trend, it seems as in recent weeks there has been a stalling of the warm temps. Water temperatures need to hit the 12.5 C mark and maintain them for several days to iniate spawning. It looks like it might be a slow start at Manchester or even a possible abortive spawn if the temps don't stay high enough long enough. I expect these guys to spawn in the last week of May, first week in June. 

Fidalgo

Temps here are colder than elsewhere but are experiencing consistent increases throughout time. It will probably be the last week of May when they spawn. 

5-8-2014 Anesthetization Issues

Dr. Roberts recently asked me why I was getting such a low response rate from the treatments. I was seeing between 5 and 50% of the animals become anesthetized. Monitoring the situation I have a few ideas on why. Which I stated in a response to him in a message which is copied below.

I think it was a combination of the warm ambient air temps that caused an increase in treatment temperature which led to animals not opening in the treatment.

I also checked my treatment calculations and the epsom salt concentration was about half. I calculated it fo 5 gallons instead of the full ten. I'll just double the amount of epsom salt to make it the proper concentration. Hopefully it will increase response from the animals.

Brent has also suggested doing sub samples instead of dosing the whole tray to limit exposure by all the animals and reduce the amount of epsom salt we are using. I think its better to dose the whole tray because I can't guarantee that the percentage of animals that open will stay the same. On top of this variable percentage, there will only be 10-20% of the animals brooding at any given time. If we don't treat enough animals we will be more likely to miss spawning effort. 

Brent also had some useful suggestions about previous work with the treatments as well as some recalculations of the treatment solution to increase treatment concentration.

Katie consistently saw about 30% response (without dessicating them first. I don't have her data on dessication, but response rate was higher).  If we stick with the 5% threshold to see brooders and a 30% response,  you'd need to treat 67 oysters to = 1.  If the response with dessication is closer to 60%, you can decrease the subsample by half.  
It sounds like the oysters may have been too warm to respond--important to bring a thermometer, and probably some ice packs to control treatment temp.  
85g/L = ~7 lbs of MgSO4 for 10 gallons.

It can be argued that we only treat half the tray at any given time to limit downstream effects from the treatment which may affect spawning including abortive spawns or gonadal regression. The problem with this is that too see any signs of spawning we would be looking for 1-2 animals in any group to be brooding. I don't think this is a reliable way to determine if the populations are spawning. We would be better served in treating the whole tray to avoid mixups and hopefully have a higher number of brooding animals to strongly suggest that they animals have spawned.