Histology Protocol

For processing in Mo’orea

Contents

• Materials
• Protocol
• References

1. Materials
   • Tripour beaker
   • 1000 mL graduated cylinder
   • 500 mL graduated cylinder
   • Hydrochloric acid (100% acide hydrochlorique from Polymat in plastic bottle with blue and black label)
   • Plastic bins (CSUN has pre-labeled bins designated for HCl processing if available)
   • PBS tablets (phosphate buffered saline)
   • DI water
   • Plastic cups
   • Tweezers
   • Fume hood
   • Scoopula
   • Sharpie
   • Gloves
   • Mask
   • Protective eyewear
   • Lab coat
   • Bucket
   • Coral rubble

2. Protocol

   1. Wear proper PPE (mask, gloves, glasses, close-toed shoes, pants, and a lab coat).
   2. Turn on the fume hood light and exhaust.
   3. In the fume hood, fill a trifold beaker with 800 mL PBS solution (DI water and PBS tablets) (fill with DI water found in LTER backlab), using a 1000 mL graduated cylinder to measure.
   4. Using the 500 mL graduated cylinder, measure 80 mL of 100% HCl and add it to the trifold beaker. Mix the solution well with a scoopula. This will make a 10% HCl diluted solution.
   5. Always make sure the water is added first before the HCl.
   6. Label all the plastic cups with the month and sample ID number twice before beginning, or read out the sample number to a labmate to label each cup during processing.
      • EX: ACR 464 April 2022 ACR 464
   7. Using the 500 mL graduated cylinder in the fume hood, measure out 40 mL of the 10% diluted HCl solution from the trifold beaker before beginning each sample.
   8. Pour the 40 mL 10% diluted HCl solution into the labeled cup.
   9. Using the tweezers, carefully remove the histological clipped fragment (specified in the same day sample processing protocol) from its designated 10% formalin fixed 5mL tube. Allow any drips to drip off into the origin tube before moving the sample carefully into the labeled cup filled with 40 mL of the 10% diluted HCl solution.
      • Keep the remaining ~2-3 mL 10% formalin solution in the 5 mL tubes for later preservation.
   10. The coral should fizz or bubble immediately.

   Coral Histology Video
   

   1. Leave the coral in the HCl solution to decalcify. This may be for several hours or days. For ~0.5 to 1 inch Acropora pulchra fragmented samples, some were completely decalcified 24 hours later with the HCl solution still being acidic (pH of 0-3 in sample cups).
   2. Check each individual sample with the tweezers gently to see if any skeletal structure remains. The sample should be squishy and there should not be any resistance in the middle of the sample when using the tweezers. You should also not see any white structures in the middle of the sample if you look from the cut area top or bottom.
      • You can immediately tell if the sample needs more time in the solution if it is not floating. The floating coral fragments are more likely to be completely decalcified or to have a small bit of skeletal structure remaining.

   

   

   1. If the sample is completely decalcified, bring it to the fume hood and find its original 5 mL labeled test tube with the residual 10% formalin solution. Carefully use the tweezers to move the coral sample back into its origin tube. Make sure the sample is as submerged as possible, they will now float.

      

   2. If not dissolving, carefully pour half of the 10% HCl solution holding the coral into a designated bucket holding dead coral rubble to further neutralize the HCl solution.
   3. Add another 40 mL of the HCl solution to any samples that need it and check back every six hours.
   4. Once all coral pieces are decalcified, put dead coral rubble and skeletons into a designated bucket outside and dispose of the remaining HCl into the bucket. The coral rubble will dissolve and neutralize the acid so that we do not have any leftover waste.
   5. Check pH level in the bucket with pH strips over the next two days to make sure the solution is around 7 before discarding the waste.
3. References