TITLE: Histological processing of octocoral tissue 
AUTHORS: Maria Rakka, Gala G Edery, Marina Carreiro-Silva

[DESCRIPTION]
In recent years, there has been a significant focus on coral habitats, encompassing both shallow and deep ecosystems. Historically, most studies concentrated on scleractinian corals, however contemporary research acknowledges the ecological importance of octocoral species, and the need to improve our understanding of their biology. Histology, which describes the internal morphology of animal tissues, is an essential tool for a wide range of biological studies, including reproductive biology, life history, taxonomy and phylogeny. Processing octocoral tissue histologically can be a complex task, given the unique skeletal structure of these organisms. Octocorals have internal skeletal structures called sclerites, which are composed of calcium carbonate and are embedded into the coral tissue. Decalcification of these structures creates air pockets in the tissue that hault further histological processing. Moreover, many octocorals possess a skeletal axis that is predominantly composed of organic matter, but may contain varying degrees of calcium carbonate. Including the axis in histological analysis may require experimentation to adjust the processing times. Although several studies report processing times for histological analysis, it is difficult to find detailed information on how to process octocoral tissue. Here, we present a detailed protocol that separates octocoral polyps from the skeletal axis, and utilizes vacuum to address issues with decalcification. The use of single polyps, or small polyp groups, decreases the processing time, and allows for manual processing without the use of automated systems. Vacuum removes trapped air from decalcification and increases the quality of the histological sections. We also include an optional step for specimens that were directly preserved in ethanol without prior fixation. Lastly, we demonstrate simple ways to section paraffin blocks that do not require extensive experience in histological sectioning.

[BEFORE_START]
Read timings and plan the protocol beforehand. It includes several long steps (<6h) and if these steps are not planed beforehand, the protocol might have to continue through the night

[GUIDELINES]
Decalcification causes entrapment of air in the coral tissue, therefore steps 10-15 have to be performed in a vacuum hood. If a vacuum hood/machine is not available, use a vacuum pump attached to a desiccator.
The protocol is for small pieces of tissue or individual polyps without the coral axis. It will still work with larger specimens, or with the axis included, but the times in processing will have to be adjusted (increased)
We include a step of Rehydration and post-fixation, that can be used if your specimens were not fixed in a fixative that is commonly used in histology (e.g. formalin, Bouin), but were preserved directly in alcohol. If you fixed your specimens, you do not need to use this step.

[STEPS]
SECTION: Fixation
1. Use 10% seawater formalin to fix the samples
SECTION: Dissection
3. Using a scalpel and forceps, make an incision parallel to the coral axis. Carefully separate the tissue from the axis by using the forceps, and remove the axis completely.
SECTION: Decalcification
7. Place tissue in eppendorfs (or other plastic/glass vials), use stickers to label
SECTION: Dissection
4. Dissect polyps or small groups of polyps by using the scalpel. If working with corals from the family Primnoidae, do not separate individual polyps from each whorl, i.e. you can work with entire whorls.
 

SECTION: Fixation
2. After 24-48 hours transfer specimens in 70% ethanol to preserve. This step might be omitted (the protocol will work with specimens preserved in formalin) but leaving specimens for too long in formalin will make the tissue harder, and may complicate sectioning.
SECTION: Staining
49. Dry excess of xylene with paper towel, mount slides with coverslips and DPX
 

SECTION: Staining
48. Pass from Xylene two times
SECTION: Staining
47. Pass from Ethanol 100% two times
SECTION: Staining
46. Pass from Ethanol 95% quickly two times
SECTION: Staining
45. Transfer to Eosin 2.5 min
SECTION: Staining
44. Transfer to Ethanol 95% 1 min
SECTION: Staining
43. Transfer to Ethanol 70% 1 min
SECTION: Staining
42. Transfer in  Distilled water 1 min-two times
SECTION: Staining
41. Transfer in warm tap water until blue
SECTION: Staining
40. Wash in HCL 1%
SECTION: Staining
39. Transfer slides to Harris haematoxylin 5 min
SECTION: Staining
38. Transfer slides to Distilled water for 1 min-two times
SECTION: Staining
37. Transfer slides to Ethanol 70% for 1 min
SECTION: Staining
36. Transfer slides to Ethanol 95% for 1 min
SECTION: Staining
35. Transfer slides to Ethanol 100% for 1 min
SECTION: Staining
34. Place slides into Xylene for 20 min
SECTION: Staining
32. Prepare the staining table. Place a label with the content on the staining vials. It is also usefull to write the time of each bath on the label.
 

SECTION: Sectioning
31. Let slides to dry overnight
SECTION: Sectioning
30. Place histological slide in warm bath to stretch the ribbon, be careful not to lose the ribbon in the water bath and do not leave for long
 

SECTION: Sectioning
29. Pick ribbon with histological slide
SECTION: Sectioning
28. Place ribbon in cold water bath
SECTION: Sectioning
27. Detach ribbon from microtome with scalpel, pick it carefully with forceps
SECTION: Sectioning
26. Perform cross sections (5-10 μm), creating a ribbon
 

SECTION: Sectioning
25. Trim the paraffin block until the area you want to section is clean
 


 

SECTION: Sectioning
24. Name slides with pencil
SECTION: Sectioning
23. Prepare cold water bath and hot water bath ~50 ºC
SECTION: Sectioning
22. Prepare microtome

SECTION: Embedding
21. Using the edge of a knife, remove carefully the block from the mold.
SECTION: Embedding
20. Move mold to a cold area (fridge) and subsequently place in refrigerator until completely solid 

SECTION: Embedding
19. Make sure there are no bubbles
SECTION: Embedding
18. Let it cool down slightly and top up with paraffinRoom temperature

 

SECTION: Embedding
17. Pick up sample with warm forceps and position it into the mold
SECTION: Embedding
16. Pour a small amount of warm paraffin into the mold
SECTION: Embedding
15. Place paraffin on a magnetic stirring hot plate at 50 ºC, keep the stirrer on at low speed to keep the paraffin temperature homogenous 50 °C 
SECTION: Embedding
14. Leave paraffin, molds and forceps in an oven at 50 ºC for at least 5-6h 50 °C 

SECTION: Processing
13. Transfer tissue into paraffin and leave in oven for 1 hour  50 °C 

SECTION: Processing
12.6. Transfer tissue in Xylene for 20 mins   
Samples will turn transparent, any white remnants are sclerites that did not get decalcified properly, discard these samples. This step needs to be done in clean vials, do not just remove alcohol and put xylene.
SECTION: Processing
12.5. Transfer tissue into Ethanol 100% for 20 mins x 3 times   

SECTION: Processing
12.4. Transfer tissue into Ethanol 95% for 15 mins   

SECTION: Processing
12.3. Transfer tissue into Ethanol 90% for 15 mins   

SECTION: Processing
12.2. Transfer tissue into Ethanol 80% for 30 mins (or just change the solution in the same vial)  
SECTION: Processing
12.1. Transfer tissue into Ethanol 70% for 30 mins   
SECTION: Processing
11. Leave paraffin in 46-50 ºC overnight to melt
SECTION: Decalcification
10. Transfer decalcified samples in clean vials with distilled water. Rinse 2,3 times, leave at least 30 mins between each rinse
SECTION: Decalcification
9. Remove water completely, add formic acid solution (10%). Do not close the lid! Solution will start bubbling. Leave samples in formic acid until no traces of calcified material is left (typically 20-45 mins, will depend on the species)
 

SECTION: Decalcification
8.3. In the morning, change the water one more time and leave for another 30-60 mins
SECTION: Decalcification
8.2. Leave specimens overnight in distilled water
SECTION: Decalcification
8.1. Using a pipette, ideally with a thin tip, remove as much fixative as possible. Fill vial with distilled water. Leave for 2-3 hours. Repeat at least 3 times 
SECTION: Decalcification
8. Rinse fixative very well with distilled water (see steps 9.1-9.2)
SECTION: Rehydration and post-fixation
5. For samples that were fixed in alcohol, these should be rehydrated before starting decalcification
 

SECTION: Rehydration and post-fixation
5.1. Transfer tissue to Ethanol 70% for 15 min
SECTION: Rehydration and post-fixation
5.2. Transfer tissue to Ethanol 45% for 30 min
SECTION: Rehydration and post-fixation
5.3. Transfer tissue to Ethanol 30% for 45 min
SECTION: Rehydration and post-fixation
5.4. Transfer tissue to Ethanol 15% for 1h
SECTION: Rehydration and post-fixation
5.5. Transfer tissue to distilled water for 2h
SECTION: Rehydration and post-fixation
6. Transfer tissue to 10% formalin for 12h
SECTION: Processing
12. Dehydration of the tissue in ethanol series in a vaccum chamber
 

SECTION: Staining
50. Put slight pressure on slides with the forceps and guide bubbles to the margins of the coverslip. Make sure there are no bubbles left on the DPX, especially on tissue. This will ensure good visualization and long-term preservation of the section.
SECTION: Staining
33. Place slides into an oven for 5 mins. This will help to remove the paraffin 50 °C