Jun 08, 2026

Coral cell dissociation and cell sampling protocol 

DOI
https://dx.doi.org/10.17504/protocols.io.eq2ly4qyrlx9/v1
  • 1Arizona State University;
  • 2School of Molecular Sciences, Arizona State University
  • Marine Biochem Research Roger Lab
Saborni Chowdhury
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DOI: https://dx.doi.org/10.17504/protocols.io.eq2ly4qyrlx9/v1
Protocol Citation: Liza M Roger, Saborni Chowdhury 2026. Coral cell dissociation and cell sampling protocol . protocols.io https://dx.doi.org/10.17504/protocols.io.eq2ly4qyrlx9/v1
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: October 20, 2025
Last Modified: June 08, 2026
Protocol  Integer ID: 230323
Keywords: Coral cell dissociation, coral cell culture, heat-stress on coral cells, cultured coral cell, stress response in cultured coral cell, coral cell dissociation, coral species, free artificial seawater to detach cell, branching coral species, cells from pocillopora acuta, coral, coral fragment, cells in the culture media, cellular heat, algae, cell, detach cell, obtaining cell, cell sampling protocol
Abstract
The protocol describes the coral cell dissociation and cellular heat-stress experimental technique for obtaining cells from Pocillopora acuta, and studying heat-stress response in cultured coral cells. It is optimized for the branching coral species and uses Ca-Mg-free artificial seawater to detach cells from the coral fragment. Compared with previous tissue extraction protocols, such as airpicking and homogenization, this protocol allows us to isolate live host (coral) and symbiont (algae) cells to study the real–time molecular mechanisms underlying heat–stress response and bleaching. The procedure requires approximately 3 hours and yields 1 x 10 cells mL-1 (from ~ 1 cm nubbin) with a viability > 80%. Users should note that the cells in the culture media can maintain viability for 24–48 hours.
Guidelines
Health and safety precautions
All work should be conducted in a biosafety cabinet, wearing standard personal protection equipment (lab coat, nitrile gloves, safety glasses).

Materials
  1. Medical clippers 
  2. Tweezers 
  3. 70% Isopropanol
  4. Kimwipes
  5. Sterile filtered artificial seawater  
  6. Sterile filtered Ca-Mg-free artificial seawater
  7. Crystallizing dish
  8. Sterile plastic petri dish (small) 
  9. Clean glass petri dish 
  10. Conical tube (50 mL and 15 mL) 
  11. Centrifuge (can reach 1200 rpm and 8000 rpm) 
  12. Pipettes and tips (sterile) 
  13. Serological pipettes ( 5 mL and 10 mL)
  14. Microcentrifuge tube (MCT)
  15. Multi–well plate (24 wells)
  16. Neubauer Hemacytometer, Disposable (iNCYTO C–Chip, DHCN015)
  17. Coral dip (Coral Rx coral dip) 
  18. Antibiotics-antimycotics solution (anti-anti) (Thermo Scientific Catalog no.15240062)
  19. Gentamicin (Thermo Scientific Catalog no. 15710064)
  20. Dulbecco’s Modified Eagle Medium (DMEM), high glucose, HEPES, and no phenol red (Thermo Scientific Catalog no. 21063029)
  21. Fetal Bovine Serum (FBS) (Thermo Scientific, Catalog no. A5670701)
  22. Hoechst Nucleic Acid Stain (Thermo Scientific Catalog no 62249)
  23. SYTOX Orange Nucleic Acid Stain (Thermo Scientific Catalog no S11368)
  24. Sodium chloride
  25. Sodium sulfate
  26. Sodium bicarbonate
  27. Potassium chloride
Safety warnings
  • Avoid multiple freeze-thaw cycles of the reagents. Always thaw the reagents at 25ºC.
  • For using a prepped medium stored at 4ºC, please make sure to pre-warm the medium to 25ºC before adding it to the cells. Temperature shock can lead to cell death.
Before start

  • Turn ON the water bath and set the temperature to 25ºC.
  • Once the temperature is set to 25ºC, thaw the following reagents in the water bath: DMEM, FBS, and Anti-Anti solution.
  • Sterilise the medical clippers and tweezers with 70% Isopropanol and Kimwipes.
  • Turn ON the UV light in the biosafety cabinet for 5 min. After 5 min, wipe down the inside surface with 70% Isopropanol to ensure proper sterilization of the biosafety cabinet.
Ca-Mg free artificial seawater preparation (CMF-ASW)
Mix the following reagents in 1 L of deionized water.
  • 23 g Sodium chloride (NaCl)
  • 0.763 g of Potassium chloride (KCl)
  • 3 g of Sodium sulfate (NaSO4)
  • 0.25 g of Sodium bicarbonate (NaHCO3).
Autoclave and sterile filter (0.22 µm)
Coral cell culture media preparation
Note: This media should be prepared in the biosafety cabinet (sterile conditions).
Prepare a 50 mL sterile conical tube. Using a 10 mL serological pipette, add the following reagents:
  • 7.5 mL of DMEM (15% final concentration)
  • 5 mL of FBS (10% final concentration )
  • 250 µL of anti-anti (0.5% final concentration)
  • 250 µL of gentamicin (0.5% final concentration)
  • Add 37 mL of FASW to adjust the total volume of the cell culture media to 50 mL.
  • Gently mix the solution.
Isolation of nubbins from Pocillopora acuta fragment
40m
Using sterile clippers, cut a single piece of coral fragment (nubbin) (~ 1 cm length) and transfer it to a crystallizing dish containing artificial seawater (ASW).
10m
Add ~ 2.64 mL of Coral Rx coral dip disinfectant solution to 500 mL of ASW in the crystallizing dish. Incubate the fragment in this solution for 10 minutes with constant aeration.
10m
After 10 minutes, return the coral fragment back into the aquarium tank. Transfer the nubbin to a sterile glass petri dish and image it with a scale.
In the biosafety cabinet, use a pair of sterile tweezers to transfer the nubbin into a sterile plastic petri dish. Add 4 mL of filtered sterile artificial seawater (FASW) (autoclaved and sterile-filtered, 0.22 µm) to the petri dish. (Use a 5 mL serological pipette to avoid contamination.)
Using a 1 mL pipette tip, rinse the nubbin with FASW (5 to 10 times on both sides of the nubbin) to wash away the mucus on the surface of the nubbin. Aspirate the solution and add another 4 mL of fresh FASW to the petridish. Repeat the washing steps twice. Altogether, rinse the nubbin three times.
20m
Dissociation of cells from Pocillopora acuta nubbin
8m
Prepare a 15 mL sterile conical tube with 3 mL of calcium-magnesium-free artificial seawater (CMF–ASW) (autoclaved and sterile filtered, 0.22 µm). Use a pair of sterile tweezers to transfer the rinsed nubbin into the conical tube containing CMF–ASW. Incubate the nubbin in CMF–ASW for 1 hour in the biosafety cabinet under ambient light.
1h
After incubation, gently wash the nubbin (gentle pipetting) with CMF–ASW to detach cells from the surface of the nubbin. As the cells detach from the surface of the nubbin, the nubbin will turn pale to white in color, exposing the underlying skeleton. The solution containing the coral cell suspension will turn turbid.
5m
Transfer the cell suspension to a fresh sterile conical tube. Do not transfer the nubbin! Centrifuge the cell suspension at 1200 rpm (204 RCF) for 3 minutes at 25ºC.
3m
Coral cell culture preparation
3m
After centrifugation, remove the supernatant and add 1 mL of coral cell culture media. Gently resuspend the cells in the media.
Coral cell count and cell viability calculation
50m
To 1 mL of cell suspension, add 10 µL of SYTOX Orange (0.2 µM final concentration) and 2 µL of Hoechst 33342 (40 µM final concentration). Note: Hoechst 33342 and SYTOX Orange are both DNA-binding dyes; however, Hoechst 33342 is membrane permeable, whereas SYTOX Orange is not. In other words, Hoechst 33342 stains all cells, while SYTOX Orange stains only cells with damaged membranes 
30m
Incubate in the dark for 30 min. While the cells are incubating, start the plate reader.
After incubation, centrifuge for 3 min at 1200 rpm at 25°C, remove the staining solution, and replace it with coral culture medium.
Add 10 µL of the stained cells to the hemocytometer chamber. Use the Brightfield (BF), DAPI, TRITC, GFP, and CY5 channels to image the cells at 4x and 20x magnification. Calculate the cell solution concentration using the formula: C = c/d/A*f
  • C = concentration (cells/mL)
  • c = average number of cells counted (cells)
  • d = depth of hemocytometer chamber ((0.1 mm Neubauer, 0.2 mm F-Rosenthal)
  • A = area of hemocytometer counted (mm2)
  • f = conversion factor (1000 mm3/mL)
20m
  • DAPI channel = Total no. of cells
  • TRITC channel =No. dead cells
  • Viability(%) = ([No. of cells in DAPI]-[No. of cells in TRITC])/ [No. of cells in DAPI]* 100.
  • CY5 channel = Symbiont (Algae) cell count
  • GFP channel = Host cell the count
Heat-stress experiment setup
50m
Obtain two 24-well plates. Plate 1 mL of cell suspension (~ 1 x 106 cells mL-1) in wells designated for each time-point. Add media to the wells around the edges to prevent evaporation. Incubate one plate at 25ºC and the other plate at 30ºC.
Cell sample collection
15m
After dissociation, collect the T0 (0 h) cell sample for both temperatures in two separate microcentrifuge tubes (MCT), one tube for 25ºC and the other tube for 30ºC. Centrifuge the cell suspension at 8000 rpm (6010 RCF) at 4 ºC for 10 mins. Store cell pellets at – 20ºC until protein extraction. Sample T6, T12, T18 and T24 accordingly.
15m
Protocol references
Roger LM, Reich HG, Lawrence E, Li S, Vizgaudis W, Brenner N, et al. (2021). Applying model approaches in non-model systems: A review and case study on coral cell culture. PLoS ONE 16(4): e0248953. https://doi.org/10.1371/journal.pone.0248953