ROD challenge protocol (version 2)

ROD = Roseovarius Oyster Disease

Protocol for an Roseovarius Oyster Disease (Juvenile Oyster Disease) Challenge using Juvenile Oysters Exposed to Diel-cycling Hypoxia and Acidification Pathogen prep and inoculation are details are documented in Version 1 of this protocol from 8/24/23.

Experimental maintenence

  • Aim for oyster exposure to CV for at least 24 hours before first water change/feeding
  • 1/2 water changes + feeding every day & full water changes on Fridays to avoid weekend water change
  • Mortality estimates on MWF

See Google calendar for maintenance schedule

Water change protocol

Notes:

  • Start with control group to avoid contamination
  • change gloves in between treatment groups (CON/ROD)
  • Ethanol gloves and surfaces to avoid contamination, but ensure it is dry before handling juveniles
  • Rinse containers and tools with ASW before using them

Materials needed:

  • Artificial Seawater (ASW, 28-30ppt)
  • Shellfish Diet (SD)
  • 70% ETOH in spray bottle
  • beakers
  • green trays - for transporting cups
  • ASW squirt bottles
  • clean baskets (if needed)

Procedure:

  1. Ethanol bench and trays and wipe dry with paper towels.
  2. Make ASW+SD. Dilute 1.5ml of SD into 3.2L of ASW (for a full water change make up 3 3.2L jars like this)
  3. Turn off airstones before moving any baskets, to avoid contamination via aerosols
  4. Remove cups for water change - start with control cups
  5. Remove basket and place it in it’s treatment-specific red bin.
  6. Into the “OLD WATER” beaker, pour out either 1/2 the volume or the full volume (depending upon if you’re doing a 1/2 or full water change).
  7. Attempt to rinse out the built up algae and poop from the basket- if possible. Tilting the basket will help to separate oysters from debris.
    • If baskets need to be changed, rinse juveniles into new labeled basket with ASW squirt bottle. The red bins can be used to catch seawater.
  8. Top off the cup with ASW + SD (measured in the “NEW WATER” beaker) for a total volume of ~650ml.
  9. Return basket to it’s cup and place back in the incubator with airstone.
  10. Turn on airstones after all water changes are complete.
  11. Bleach ROD contaminated seawater for 15 mins and pour out.
  12. Dry, bleach, and ethanol the benchtop.
  13. Bleach all glassware, bins, caps, squirt bottles, baskets, etc. following the Puritz Lab bleaching protocol (see below).
  14. Set glassware and other materials to dry on the drying pad.
  15. Double check the air bubbles and make sure the incubator is propped open before leaving for the day.
  16. Check ASW stock, bleach, and sodium thio and text Megan/Caitlin if any needed to be replenished.
  17. Record water change and any notes in lab notebook (located on Puritz Lab bench).

Bleaching protocol

Materials needed:

  • bleach
  • sodium thiosulfate
  • water
  • DI

bleach

Procedure:

  1. Bleach with dilute bleach solution (a quick pour of bleach + water should be good). Be sure to bleach the outside of jars/beakers too with 10% bleach squirt bottle (on Puritz bench).
  2. Let sit for at least 15mins
  3. Pour out bleach
  4. Neutralize remaining bleach with sodium thiosulfate crystals dissolved in water (pour some crystals into each container and add hot water & mix)
  5. Rinse everything very, very well 5-6x in hot water
  6. Rinse in DI water

Mortality estimates

Materials needed:

  • Magnifying glass or dissecting scope
  • Petri dishes
  • tweezers or paint brush
  • counters

Note: This dissecting scope works really great!

scope

Procedure:

  1. Transfer all oysters from 1 cup to their treatment-specific dish
  2. Under magnification, assess whether they are alive or dead keeping count with counters
    • living oysters will be closed and have some color to them
    • dead oysters can look gapped open, often have no color, and sometimes they float a little
  3. Set dead oysters to the side and record alive/dead counts in notebook
  4. Place all of the living oysters back into their basket and place the dead oysters into a petri dish for imaging
  5. Imaging the dead oysters.
    • Place the dish with dead oysters on a good background with good lighting so you can easy see the margins of the shells
    • Add a paper tag with the date, cup ID (ex. ROD-CADO1), and your initals on or near the dish (to help keep track of the dead from each cup)
    • Lastly, place a ruler or calipers near the dish for scale and take a good quality image of the dish, tag label and scale ruler
    • Save image to google drive folder (“ROD images”)
    • Example images:
      mortimage1 mortimage2
  6. Preserve dead oysters/shells in 95% ethanol
    • If you see any oyster with tissue still inside, carefully dissect out the tissue and place it in its own labeled tube and fill tube with ethanol
    • Place remainder of dead oysters/shells in a 2ml tube and fill tube with ethanol (we should be able to use just 1 tube/replicate for the whole challenge)
    • Place tubes in a box in the 4C fridge
  7. Repeat steps 1-6 for at least 1 cup from each treatment group
  8. Update Megan with % mortality estimates for the day (we are aiming to get 60-70%)
  9. Dishes can be wiped out with a kim wipe – do the control dishes first to avoid contamination!

Final mortality counts and sampling

When mortality is around 60-70%, get good final counts of all of the replicates in each treatment group using the above procedure.

Sample remaining, living juveniles from each replicate for genomic sampling.

  • Add oyster to tube
  • Flash freeze the tube in liquid nitrogen
  • Store at -80C
Written on October 19, 2023