Feb 25, 2026
Version 2
Making fresh frozen and formalin-fixed paraffin embedded sections for Hydractinia V.2
- Zachary Lane1
- 1University of Florida
Protocol Citation: Zachary Lane 2026. Making fresh frozen and formalin-fixed paraffin embedded sections for Hydractinia. protocols.io https://dx.doi.org/10.17504/protocols.io.n92ld6x2xg5b/v2Version created by Zachary Lane
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 25, 2026
Last Modified: February 25, 2026
Protocol Integer ID: 243959
Keywords: tissue sections for hydractinia, sections for hydractinia, hydractinia researcher, hydractinia, staining tecdhnique, colormetric staining, based spatial transcriptomic, spatial transcriptomic, useful for fluorescent, ffpe section, ffpe section creation, tissue section, fluorescent, ff section, ffpe
Funders Acknowledgements:
Christy Schnitzler, Ph.D.
Grant ID: NIH R35 GM138156
Abstract
These protocols describe methods for producing fresh frozen (FF) and formalin-fixed paraffin embedded (FFPE) tissue sections for Hydractinia. FFPE section were once a major part of research in this species, but have fallen out of typical use. This has lead to a generation of Hydractinia researcher not having access to the technique, which remains quite useful for fluorescent/colormetric staining and other applications. FF sections have not typically been used in this species, but the ability to generate these sample opens up new avenues of research including array-based spatial transcriptomics and staining tecdhniques that are incompatible with the harsh chemical used in FFPE section creation. In general, both techniques allow for better cell-type resolution that whole-mount imaging.
Guidelines
Generally speaking, Hydractinia researcher who hope to utilize these types of samples will need to generate them themselves, unlike mammalian researcher who can generally outsource sectioning to large-scale commercial enterprises. This is due to the extra care and specific know-how required to embed and section the tiny amounts of tissue that comprise each Hydractinia polyp. This extra care bars most high-throughput facilities from attempting the task.
The protocol and troubleshooting advice are a good start for those hoping to section Hydractinia, but it should be expected that high-quality samples will not be generated without a significant amount of practical experience (i.e., practice). Good luck! These techniques yeild very beautiful results.
Materials
Fresh Frozen Section Preparation
Menthol seawater
- Place a small menthol crystal (~100 mg) into 12 ml of filtered seawater.
- Incubate at room temperature on rocker overnight
MgCl seawater
- Dissolve 40 g of MgCl in 450 ml diH2O; add diH2O to a final volume of 500 ml
- Add 500 ml of filtered seawater.
Vannas Scissors
Cryostat
Cryostat blades
Cryostat chuck
Cryostat heat extractor
Cryo molds
OCT liquid
Glass petri dishes
Small camel hair brush
Forceps
Superfrost Plus microscope slides
Stereo Microscope
Formalin-fixed Paraffin Embedded Section Preparation
Sorensen's Phosphate Buffer 7.0 pH
- Solution A: dissolve 3.56 g of sodium phosphate monobasic monohydrate in 100 ml of H2O
- Solution B: dissolve 2.76 g of sodium phosphate dibasic dihydrate in 100 ml of H2O
- Combine 2 parts Solution A and 3 parts Solution B; Do not combine all of either solution.
- Measure pH. If pH is > 7.0, add some Solution A to lower it to 7.0. If pH is < 7.0, add some with Solution B to increase it to 7.0. This process is similar to standard titration.
MgCl seawater
- Dissolve 40 g of MgCl in 450 ml diH2O; add diH2O to a final volume of 500 ml
- Add 500 ml of filtered seawater.
Vannas Scissor
Paraffin wax (melt point 56-58°C)
Paraffin bath (preferrably with pour spout)
Peel Away Histology molds
Microtome
Microtome cassettes
Microtome blades
Paraformaldedyhe
100% Ethanol
Xylene
Cell strainers
Glass petri dishes
Glass slide staining rack
Glass slide staining jar (preferrably x 12)
Small camel hair brush
Forceps
Water bath (large tupperware is fine)
Superfrost Plus microscope slides
Microscope slide cover slips
Crafting clay
Heating plate capable of maintaining surface temperature of 70°C
Small flashlight or other adjustable lightsource
Alcohol lamp or Bunsen burner
Broad sided knife or other flat metal object with handle
Small cooler
Hard-sided plastic freezer packs
May-Grunwald staining solution
Giemsa staining solution
Troubleshooting
Problem
FFDirty or sticky working area
Solution
Try not to breath on or warm the working area. This is where most of the stickiness comes from while cryosectioning. If the area does become sticky, wipe it with a kim wipe or paper towel. It it remains sticky, wipe the area again and then close the lid of the microtome so that it can cool back to the desired temperature.
Problem
FF Curled sections
Solution
Turning the handle slowly, use a camel hair brush to catch the section as it is cut, preventing it from curling. Once caught, contine to turn the handle and use the paintbrush to pull the section very gently away from the blade. Once the section is fully cut, you should be able to pull it free from the blade afterwich it should lay flat. If it does not lay flat after it is separated from the blade, consider increaseing the temperature from -20C to -18C or -15C.
Problem
FFPE Dirty or sticky working area
Solution
Keep a small amount of xylene on hand. Clean the blade and sectioning area between each strip using a cotton tipped application soaked in xylene. Keep xylene cover with tin foil to avoid unnecessary inhalation.
Problem
FFPE Curled sections
Solution
Use a paintbrush to catch the section as it is cut, preventing it from curling. Once one section has come out flat, the rest should do so as well. Consider leaving one flat section attached to the blade to avoid future curling (i.e., when transferring section to the bath, always leave one behind).
Problem
FFPE Sections curving to one side
Solution
This usually indicates that some portion of the blade is dull. Moving the blade holder so that a uniformly sharp portion of the blade is being used. If you cannot find a uniformly sharp portion, change the blade.
Problem
FFPE Crumpled Sections
Solution
Put the cassette on ice for 5 minutes to harden the wax in order to get flatter sections.
Problem
FFPE Splitting sections
Solution
Remove all sections from working area and clean the blade. If problem persists move to an unused section of blade or replace the blade is too dull.
Safety warnings
Please refer to all SDS for the reagents used in these protocol. Standard PPE is recommended throughout. Be extra careful when working with microtome/cyrostat blades and when handling sub-freezing temperatures.
Ethics statement
These protocol do not require ethics approval by IACUC. Hydractinia is an invertebrate model without a centralized nervous system or body plan. Dissections of feeding polyps do not negatively impact the survival of the greater colony.
Before start
Read the entirety of the protocol you are executing. Some things will need to be prepared in advance (e.g., cyrostat cooling, dehydration/rehydration ethanol solutions, etc.).
Fresh Frozen Section Preparation
Dissection and Embedding
Place the colony you plan to use in MgCl seawater. Incubate for 15 min to relax and enlongate the polyps
Dissect the polyps you would like to section and transfer them to a petri dish containing MgCl seawater.
Fill cyro mold with OCT liquid, being careful to avoid introducing bubbles.
Transfer 1-3 poyps Menthol seawater incubate for 30 seconds.
This step ensures that the polyps will not contract once placed in the OCT liquid, but it is necessary that the polyps are exposed to the Menthol seawater for only a short time to avoid the tissue degradation that accompanies prolonged exposure to Menthol.
Transfer the polyps to the OCT liquid in the mold.
Under a stereo microscope use an insect pin or other fine tool to gently position the polyps along a singular plane at the base of the mold.
Flash freezing
Flash freezing of Hydractinia can be accomplish in several ways. Many laboratories usilize consumables (i.e., liquid nitrogen or dry ice) or speciallized equipment (e.g., cryobaths) to perform flash freezing. This protocol has been developed to utilized a standard -80°C chest freezer so that the process can be performed without time sensative consumables or uncommon laboratory equipment.
This protocol assumes that you have a flat surface with roughly 20 cm3 of space within the -80°C chest freezer to use as a working area.
Failure to flash freeze Hydractinia tissues result in the formation of water crystal during the freezing process which cause irreparable to the tissues.
Place a cyostat chuck and cryostat heat extractor into the -80°C chest freezer several hours before needed for flash freezing to allow both objects to reach -80°C. This should be accomplished well in advance of the Dissection and Embedding steps above.
Depending on cyrostat brand and shape of associated cryostat chuck, you may also need a cardboard freezer storage box for 1.5 ml microcentrifuge tubes in order to hold the cryo chuck in an upright position throughout the flash freezing process.
In quick succession:
1) Place the pre-chilled cryo chuck onto the mold containing the dissected polyps such that it contacts the exposed OCT liquid
2) Place the chuck, with mold attached, into the -80°C chest freezer so that the face of the chuck contacting the OCT liquid is facing upward (i.e., with the mold on top)
3) Place the pre-chilled heat extractor on top of the mold
4) Close the freezer
Flash freezing should only take a few moments. After a minute, remove the sample from the freezer and transfer it quickly into the cyrostat. If possible, transfer the sample with the heat extractor in place to ensure that sample is not exposed to temperatures above freezing at any point during the transfer.
Cryosectioning
Set the cryostat to -20°C before beginning work. Carefully remove the OCT block from the mold.
Put the chuck in the cryostat chuck holder and adjust the it so that:
A) the face of the wax block is as on-plane as possible with the blade
B) the top of the block is as close as possible without touching the blade
The polyps are very near to the front edge of the block, so proper aligment is critical to ensure that high quality sections are being cut as soon as possible once the block contacts the blade.
Begin to crank the handle until the machine begins to produce proper section (i.e., sections cut from the entire face of the OCT block). The polyps should be very near to the base of the block, so do not waste any more sections than necessary.
Hydractinia tissues should be visible in colorless (i.e., white) OCT liquid. Liquid can be bought in other colors if greater contrast is desired.
Once proper sections are being cut, use a camel hair brush to catch the sections as they are cut and pull them gently from the blade to get them to lay flat. An antiroll plate may be used instead, but with enough practice, using a brush give more consistent results in my experience.
Continue to section the block. Occassionally check your progress by adhering a flattened section to a Superfrost Plus Microscope slide by pressing the face of the slide directly onto the section as it lies on the working area. After the section sticks to the slide, remove it from the cyrostat and melt the section to the slide by pressing your finger to the underside of the slide directly below the section.
After melting, check the section under a stereo microscope and decide if you are yet sectioning the portion of tissue you would like to sample. If not, continue to section and occassionaly check your progress as outline above.
Once you are sectioning the portion of tissue you are interested in, begin to collect as many samples as needed by flattening and adhere the sections to Superfrost Plus slide (or other appropriate device as called for by downstream application) as outline above.
Proceed with protocol for your downstream application (e.g., live/fixed staining procedure, spatial transcriptomics tissue processing, etc.)
Formalin-fixed Paraffin Embedded Section Preparation
Dissection and Fixation
Place the colony you plan to use in MgCl seawater. Incubate for 15 min to relax and enlongate the polyps.
Dissect the polyps you would like to section and transfer them and a small amount of MgCl seawater to a microcentrifuge tube.
Remove as much supernatant as possible. Add 1.5 ml of 4% PFA in filtered seawater to the sample and incubate at room temperature for 2 hours on a rocker.
Dehydration and Xylenation
Pour polyps into cell strainer mounted on appropriately sized tube to ensure fixative is safely removed.
Perform the following serial dehydration/xylenation by transferring the cell strainer containing the polyps to petri dishes containing the specified concentration of ethanol or xylene.
1x10 min 50% Ethanol
1x10 min 70% Ethanol
1x10 min 80% Ethanol
1x10 min 95% Ethanol
2x10 min 100% Ethanol
2x10 min 100% Xylene
Keep the polyps in the final Xylene bath until you embed them.
Embedding
Turn a heating plate on to 70°C and light alcohol lamp to heat tools as needed throughout process.
Pour liquid paraffin into 1-4 molds such that only the pyramidal base of the molds is full. Transfer to molds to the heating plate. The entirety of the volume of wax in each mold should stay liquid. If it does not, the molds may be over filled or the heat may be too low.
Put 2-4 polyps into each wax-containing mold using forceps. Let sit for 10 min, then use forceps to arrange them in a radial pattern and then remove the mold from the heating plate to cool.
Note: Positioning the polyps is easier with the lights off in the room and a bright light (e.g., a flashlight) shining sideways through the mold. This allows you to see where you are positioning the unstained polyps by using their shadows.
Note: The alcohol lamp or bunsen burner should be used to occasionally melt the paraffin that solidifies on the forceps.
Once the blocks have cooled, remove them from their molds.
Place the block onto your broad-sided knife and gently hold in place using forceps. Be careful not to make any indentations on the face of the block with the forceps.
Position the knife over the alcohol lamp or bunsen burner until the paraffin block begins to melt. At this point quickly transfer the block and the liquid wax to a microtome cassette. If done properly, the block should be firmly affixed to the cassette once it cools back to room temperature.
Blocks can be stored indefinately at room temperature until it is time for sectioning
Sectioning
Fill water bath. Prepare microtome for use (e.g., install new blade, set to preferred thickness, etc.). Put ice in cooler and place hard-sided plastice freezer pack into the cooler.
Put the cassette in the cassette-holder and adjust the it so that:
A) the face of the wax block is as on-plane as possible with the blade
B) the top of the block is as close as possible without touching the blade
The polyps are very near to the front edge of the block, so proper aligment is critical to ensure that high quality sections are being cut as soon as possible once the block contacts the blade.
Remove the cassette and place block face down on the freezer pack in the cooler. Allow to cool for 5-10 mins with cooler lid shut. This is step may not be necessary if the ambient temperature in the workspace is adequately low (e.g., 65-68ºF).
Place the cassette back onto the microtome, being sure that it is oriented the same as when you adjusted the cassette holder in step 4.2.
Begin to crank the handle until the machine begins to produce a proper ribbon (i.e., strip of serial sections each taken from the full face of the block). The polyps should be very near to the base of the block, so do not waste any more sections than necessary.
Sectioning in general can be difficult. The camel hair brush will be of great use, as will patience. See the troubleshooting section for more specific advice.
Transfer ribbons of 3-5 sections at a time to the water bath using the forceps. Lay them gently on the surface of the water, where they should flatten out and float.
To attach the sections to the Superfrost Plus microscope slides, dip the slides vertically into the water bath and attach the sections by gently guiding them to the slide using forceps and then pulling the slides out of the bath. The sections should be pull onto the slide and lie flat as the slide is removed from the water bath.
Allow the slides to dry in a slide holder overnight. Store dried, wax-embedded section at room temperature for several months or longer. If prolonged storage is preferred, it is better to leave block unsectioned until needed.
Determinine when the you have fully sectioned through the tissue in the block is difficult do to a lack of contrast between the polyp tissue and the paraffin wax. You may be able to make out the presence of tissue by shining a flashlight through the wax. Continue to section until you are comfortable that you have finished the job.
Deparaffinization and Rehydration
Place 10 air-dried slides with paraffin sections attached into glass slide staining rack.
Perform the following serial deparaffinization/rehydration by transferring glass slide staining rack to glass slide staining jars containing the specified concentration of Ethanol, Xylene, or H2O.
1x10 min 100% Xylene
1x5 min 100% Xylene
1x5 min 50/50 Xylene/Ethanol
2x5 min 100% Ethanol
2x5 min 95% Ethanol
1x5 min 80% Ethanol
1x5 min 70% Ethanol
1x5 min 50% Ethanol
2x5 min 100% diH2O
Allow samples to air dry for 2 hours before staining.
Pappenheim staining for i-cell identification
Place slides in petri dish containing undiluted May-Grünwald solution using forceps or other appropriate device. Incubate for at room temperature in dark for 3 minutes.
Remove slide, pour off May-Grunwald, and transfer to 1:15 diluted Giemsa in Sorensen's phosphate buffer. Inbuate at room temperature in dark for 10 min.
Remove slide, pour off Giemsa, and transfer to Sorensen's phosphate buffer. Incubate for 3 min at room temperature. Repeat 1 time.
Remove slide and mount in Sorensen's phosphate buffer using cover slips with clay feet.
Protocol references
Müller, W. (1964). Experimentelle Untersuchungen über Stockentwicklung, Polypendifferenzierung und Sexualchimären bei Hydractinia echinata. Wilhelm Roux'Archiv für Entwicklungsmechanik der Organismen, 155(2), 181-268.
Müller, W. A., Teo, R., & Frank, U. (2004). Totipotent migratory stem cells in a hydroid. Developmental biology, 275(1), 215-224.
Acknowledgements
Thank you very much to Jessica Farrell and Paul Linser for their help with development of this protocol