Apr 09, 2024
Histological processing of octocoral tissue
- Maria Rakka1,
- Gala G Edery2,3,
- Marina Carreiro-Silva2,3
- 1Dalhousie University;
- 2Institute of Marine Sciences-Okeanos;
- 3IMAR-Institute of Marine Research
Protocol Citation: Maria Rakka, Gala G Edery, Marina Carreiro-Silva 2024. Histological processing of octocoral tissue . protocols.io https://dx.doi.org/10.17504/protocols.io.81wgbx38nlpk/v1
Manuscript citation:
Rakka, Maria, Iris Sampaio, Ana Colaço, and Marina Carreiro-Silva. "Reproductive biology of two deep-sea
octocorals in the Azores Archipelago." Deep Sea Research Part I: Oceanographic Research Papers 175 (2021): 103587. https://doi.org/10.1016/j.dsr.2021.103587
License: This is an open access protocol distributed under the terms of the Creative Commons Attribution License, which permits unrestricted use, distribution, and reproduction in any medium, provided the original author and source are credited
Protocol status: Working
We use this protocol and it's working
Created: February 21, 2024
Last Modified: April 09, 2024
Protocol Integer ID: 95481
Keywords: Octocorals, coral, decalcification, Histology, histological processing of octocoral tissue, processing octocoral tissue, octocoral tissue, coral tissue, octocoral polyps from the skeletal axi, scleractinian coral, internal morphology of animal tissue, octocoral species, many octocoral, ecological importance of octocoral species, unique skeletal structure of these organism, octocoral, animal tissue, significant focus on coral habitat, octocoral polyp, coral habitat, internal skeletal structure, unique skeletal structure, further histological processing, calcium carbonate, degrees of calcium carbonate, tissue, histology, internal morphology, histological processing, processing times for histological analysis, histological section, extensive experience in histological sectioning, air pockets in the tissue, histological sectioning, morphology, axis in histological analysis, histological analysis, quality of the histological section, skeletal axi, calcium, biological study, called sclerite, organism, phylogeny, speci
Funders Acknowledgements:
Direção Regional da Ciência e Tecnologia
Grant ID: M3.1.a/F/047/2015.
iAtlantic H2020 project
Grant ID: 818123
MIDAS H2020 project
Grant ID: 603418
Disclaimer
We have put great effort to include all relevant information and warnings, however the reader should make their own evaluation as to the appropriateness of any experimental technique for their objectives. Histological processing requires handling of toxic and corrosive substances, therefore the reader should take all essential precautionary measures. This output reflects only the author's view and the European Union cannot be held responsible for any use that may be made of the information contained therein
Abstract
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.
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.
Materials
Reagents
- 10% seawater formalin
- Distilled water
- Formic acid 10%
- Ethanol solutions (15%, 30%, 45%, 70%, 80%, 90%,95%,100%)
- Xylene
- Paraffin
- Hydrochloric acid (1%)
- Harris haematoxylin
- Eosin
- DPX
Equipment
- Oven
- Vacuum hood or similar
- Heating and stirring plate
- Refrigerator and freezer
- Microtome
- Forceps and scalpel
- Histological molds
- Holders and vials for histological staining
- Water bath (or other way to keep water at ~50 ºC)
Consumables
- Eppendorf vials
- Glass and/or plastic pipettes
- Beakers
- Histological slides
- Coverslips
Troubleshooting
Safety warnings
- Formic acid and hydrochloric acid are corrosive and toxic, use a lab hood, gloves and glasses and take care when handling them
- Xylene is toxic, use a lab hood and gloves during processing and especially during staining
- Formalin is toxic, use a lab hood and gloves
- DPX is toxic, use a lab hood and gloves
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
Fixation
Use 10% seawater formalin to fix the samples
1d
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.
Dissection
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.
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.
Note
There is a thin tissue layer separating the polyp and the coral axis. Make sure you don't cut through the base of the polyp, otherwise its content including gametes, may be lost.
Example of dissection that destroyed the base of the polyp, exposing and spilling coral gametes. (species: Viminella flagellum)
Rehydration and post-fixation
For samples that were fixed in alcohol, these should be rehydrated before starting decalcification
Note
This section (Rehydration and post-fixation) is only needed if the specimens were fixed and preserved in ethanol, i.e. if there was no fixation in formalin, Bouin solution, or other fixatives typically used in histology. The goal of the section is to reverse the dehydrating effect of ethanol that may have caused shrinkage of tissue and fixing it in formalin before proceeding with the rest of the protocol.
Attention: If you have fixed your specimens in formalin or other fixatives, and then transferred them to ethanol you do not need to perform this section. Jump directly to decalcification.
Transfer tissue to Ethanol 70% for 15 min
15m
Transfer tissue to Ethanol 45% for 30 min
30m
Transfer tissue to Ethanol 30% for 45 min
45m
Transfer tissue to Ethanol 15% for 1h
1h
Transfer tissue to distilled water for 2h
2h
Transfer tissue to 10% formalin for 12h
12h
Decalcification
Place tissue in eppendorfs (or other plastic/glass vials), use stickers to label
Rinse fixative very well with distilled water (see steps 9.1-9.2)
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
9h
Leave specimens overnight in distilled water
12h
In the morning, change the water one more time and leave for another 30-60 mins
1h
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)
45m
Transfer decalcified samples in clean vials with distilled water. Rinse 2,3 times, leave at least 30 mins between each rinse
1h 30m
Processing
Leave paraffin in 46-50 ºC overnight to melt
12h
Dehydration of the tissue in ethanol series in a vaccum chamber
Note
Ideally processing should be done in a vacuum chamber, to remove all the bubbles that were created during decalcification. Without the vacuum chamber, bubbles will not let the solutions to reach the tissue and the procedure will fail. A desiccator attached to a vacuum pump can work too.
Transfer tissue into Ethanol 70% for 30 mins 100 mbar
30m
Transfer tissue into Ethanol 80% for 30 mins (or just change the solution in the same vial)100 mbar
30m
Transfer tissue into Ethanol 90% for 15 mins 100 mbar
15m
Transfer tissue into Ethanol 95% for 15 mins 100 mbar
15m
Transfer tissue into Ethanol 100% for 20 mins x 3 times 100 mbar
1h
Transfer tissue in Xylene for 20 mins 100 mbar
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.
20m
Transfer tissue into paraffin and leave in oven for 1 hour 50 °C oven
1h
Embedding
Leave paraffin, molds and forceps in an oven at 50 ºC for at least 5-6h 50 °C oven
6h
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 stirring hot plate
Pour a small amount of warm paraffin into the mold
Pick up sample with warm forceps and position it into the mold
Let it cool down slightly and top up with paraffinRoom temperature
Note
During embedding, histological cassettes are often used to aid with securing the block during sectioning. We do not use them here (see sectioning for how we position and secure the block), but feel free to use them.
Make sure there are no bubbles
Move mold to a cold area (fridge) and subsequently place in refrigerator until completely solid
Using the edge of a knife, remove carefully the block from the mold.
Sectioning
Prepare microtome
Prepare cold water bath and hot water bath ~50 ºC
Name slides with pencil
Trim the paraffin block until the area you want to section is clean
Note
As shown in the images below, we use an unconventional way to section our blocks. Instead of using cassettes and taking frontal sections of each block, we rotate blocks and take transverse sections. This reduces the required time to to trim the blocks, and does not require histological cassettes.
Paraffin block with samples placed on the microtome to be sectioned.
Perform cross sections (5-10 μm), creating a ribbon
Paraffin ribbon after block sectioning on the microtome.
Detach ribbon from microtome with scalpel, pick it carefully with forceps
Place ribbon in cold water bath
Pick ribbon with histological slide
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
Let slides to dry overnight
Staining
37m 20s
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.
Staining set up.
Place slides into an oven for 5 mins. This will help to remove the paraffin 50 °C oven
5m
Place slides into Xylene for 20 min
20m
Transfer slides to Ethanol 100% for 1 min
1m
Transfer slides to Ethanol 95% for 1 min
1m
Transfer slides to Ethanol 70% for 1 min
1m
Transfer slides to Distilled water for 1 min-two times
2m
Transfer slides to Harris haematoxylin 5 min
5m
Wash in HCL 1%
10s
Transfer in warm tap water until blue
10s
Transfer in Distilled water 1 min-two times
2m
Transfer to Ethanol 70% 1 min
1m
Transfer to Ethanol 95% 1 min
1m
Transfer to Eosin 2.5 min
2m 30s
Pass from Ethanol 95% quickly two times
10s
Pass from Ethanol 100% two times
10s
Pass from Xylene two times
10s
Dry excess of xylene with paper towel, mount slides with coverslips and DPX
Final histological preparations.
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.
Protocol references
Price K, Peters E (2018) Histological techniques for corals. Draft2Digital