Here you will find a protocol adapted from Coral Histology to be use for Excavating Sponges Histology.
You can reference this protocol as: Chaves-Fonnegra, A. Increase of excavating sponges on Caribbean coral reefs: reproduction, dispersal and coral deterioration. Doctoral Dissertation, Nova Southeastern University, 2014. pp. 195.
--------------------------------------------------------------------------------------------------------------------------------------------------- STEP 1- FIXATION: Fix tissues overnight, 24h (or for extended periods up to 3 weeks) in Bouin´s fixative . Use gloves and a fume hood, or if in a boat or in the field station, do it outside. --------------------------------------------------------------------------------------------------------------------------------------------------- STEP 2- RINSE: Rinse in distilled water 2 times for 10 to 15minutes. --------------------------------------------------------------------------------------------------------------------------------------------------- STEP 3- DECALCIFICATION: Use Hydrochloric acid (HCl) 10% plus EDTA. Leave specimens for 2 to 3 days under the fume hood, use glass beakers of 200ml per each sample (Coral lab do it in a big plastic tray and separate samples with plastic rings)
To prepare 3L (3000ml) of 10% HCl use: 300ml of HCl (Concentrated which usually is 37%) 3g of EDTA 2700ml of dH2O. Mix well in a glass bottle, and put with a stir bar on a mixing plate for 20min to 1h. Depending of the quality of the EDTA it can take longer. ---------------------------------------------------------------------------------------------------------------------------------------------------STEP 4- RINSE: After decalcification rinse specimens 3x30 min to 1hour in Distilled Water prior to ethanol. --------------------------------------------------------------------------------------------------------------------------------------------------- STEP 5- DEHYDRATION Transfer specimens to a glass vial, add and remove each of the following :
50% ethanol – 1 hour (can be stored in 50% ethanol for longer periods).
70% ethanol – 1 hour (then stop here for Desilicification: remove most of 70% ethanol and add 4% hydrofluoric acid prepared in 70% ethanol to plastic 50ml falcon tubes (Prepare fresh every time under a fume hood) and add until covering the specimen (usually 3-5ml, but depends on the size of the tissue). Leave the specimen in the 4 °C refrigerator overnight. The following day, remove the HF/Ethanol to a waste container and add fresh 70% Ethanol, enough to ensure the HF has been rinsed out, 20 min). Continue with dehydration series. [HF waste was stored in a 4 °C refrigerator until it could be safely disposed as chemical waste, while carefully following all MSDS protocols].
95% ethanol – 15 min x 2
100% ethanol – 10 min x 2-3. Depending on the size of the specimen; longer for larger specimens). Note: absolute 100% ethanol causes shrinkage and hardening, so prolonged time in absolute is not good. --------------------------------------------------------------------------------------------------------------------------------------------------- STEP 6- CLEARING Draw off the 100% Ethanol and add Xylene or Toluene – 15 min x 2. Note: Xylene and Toluene melt plastics, use glass beakers or vials. I used Xylene.
--------------------------------------------------------------------------------------------------------------------------------------------------- STEP 7- EMBEDDING Draw off the Xylene and place the tissue in molten paraffin in a beaker on the oven at 60 °C– 15min x 2.
Transfer the tissue to a metallic or plastic mold with fresh molten paraffin. Orient the tissue on the bottom of the mold, press the tissue softly to release any trapped bubbles. Place the embedding rings. Pour in more wax, and move the boat off the hot plate to an ice cold plate. --------------------------------------------------------------------------------------------------------------------------------------------------- STEP 8- SECTIONING
Perform sections between 0.4 and 0.5 µm, thicker sections at 10 µm can also be used, but usually are too thick to see the structures clearly. Use good microtome knifes, and be cautious that even desilicifing and decalcifying sometimes does not eliminate other minerals that can break the tissue. Sections should be done slowly and if you find minerals take a dissecting needle and pull them out. This can leave holes in the block and in your sections, so be careful. Sometimes holes are unavoidable for certain species or samples of sponges. Use slides with a space to mark and proper markers or pencil that will not get erased during the staining procedure. ---------------------------------------------------------------------------------------------------------------------------------------------------