Fixing and decalcifying fragments from four species from CBLS wetlab

Montipora capitata, Porites compressa (both collected from HI in October 2023), Pocillopora acuta, and Seriatopora sp. (both from Ocean State Aquatics, long-term residents in CBLS wetlab)

Protocol here

1. Fixing in two PAXgene containers for good tissue:fixative ratio, fragments do not need to be separated because we are not using these for downstream molecular work, just morphology.
   • Day 1 (10/10/23): Fixed tissue in PAXgene fixative
   • Day 2 (10/11/23): Replaced fixative with stabilizer, transferred to 4 ºC
   • Day 3 (10/12/23): Started decalcification, kept on shaker in cold room (4 ºC)
   • Day 4 (10/13/23): Changed EDTA solution
   • Day 5 (10/14/23): Changed EDTA solution
   • Day 6 (10/15/23): Changed EDTA solution
   • Day 7 (10/16/23): Decalc still not done for Porites or Montipora! Changed EDTA solution for these two and removed the POC and SER branches into PBS @ 4ºC.
   • Day 8 (10/17/23): Changed EDTA solution for Porites and Montipora

   • Day 9 (10/18/23): Removed the Porites and Montipora into PBS @ 4ºC.

   • to do : Decalcification done, tissue washed (10 mins, 1X PBS) and split each tissue piece into two fragments
     • one goes to sucrose for cryoprotection and embedding for cryosectioning
     • one substantial “tunic” will be transferred to 1X PBS (or paxgene stabilizer??) for confocal imaging as seen in Yost et al. 2013
       • From Yost et al. methods: “2.2. Microscopy Sub-fragments (approximately 1–2 cm2) of each coral were fixed in 4% paraformaldehyde for 1 h and then de-calcified in 10% HCl until the tissue tunics (intact biological tissues) were skeleton-free. The tissue tunics were stored in 1× PBS at 4 ◦C in the dark. Decalcified tissue tunics were bisected with a scalpel and visualized using confocal microscopy.”

Images before decalcification:

Montipora capitata

Porites compressa

Pocillopora acuta

Seriatopora sp.

All corals in decalcification (we are not using these for RNA so we can keep them in the same container):

Images after decalcification:

Montipora capitata

decalcificaton not fully done, taken on 10/16/23:

Porites compressa

decalcificaton not fully done, taken on 10/16/23:

Pocillopora acuta

=

Seriatopora sp.

Imaged Tissue “Skirts” on the following days:

• 10/19/23: Pocillpora, did not work well
• 10/31/23: Pocillpora
• 11/3/23: Seriatopora + more Poc
  • chlorophyll A (CPhA): 639 laser, power @ 8, gain 700
  • GFP (TGFP): 488 laser, power @ 7, gain 700
  • chlorophyll B (CPhB): 405 laser, power @ 4, gain 700
    • probably redundant
• 11/10/23: Porites
  • did not work well at all

Embedded samples on 11/3/23

Attempted sectioning on 12/1/23 but failed.

• Had lots of issues with sections rolling and crumpling
• The Porites and Montipora were almost melting as I sectionined them? And the sections I did get were undefined and weird. Maybe we need to skip the sucrose step, becuase it is leading to an explosion in the porites tissue and maybe also the others that is less evident.

Microscopy methods:

• Yost et al. methods: “Samples were scanned with excitations of violet (405 nm) and green (498 nm and 543 nm) light, and emissions were collected at 450 nm to visualize host tissues and at 600 nm to visualize autofluorescence of Symbiodinium. Measurements of tissue thickness were taken in triplicate at random locations on each tissue sample to determine a colony average. Coral tissue thickness was characterized by either (i) high biomass, thick tissues anastomosing through highly perforate skeletons of perforate corals or (ii) low biomass, thin tissue ‘veneers’ of imperforate corals. Thus, tissue thickness did not appear to be altered by water absorption mechanisms (e.g., in the gastrovascular cavity) that may alter tissue thickness.”
• Other papers:
  • Liu et al 2020: “DAPI was excited using a 410‐nm laser, and the fluorescence signal was collected with a 490‐nm bandpass filter. Coral green fluorescence protein and apoptosis TUNEL assays were excited using a 488‐nm laser and collected with a 533‐nm bandpass filter. […] The autofluorescence of the chlorophyll was excited using a 410‐nm laser and collected with a 610‐nm long‐pass emission filter.”
  • Majerova et al 2021: “Host cell fluorescence was excited with the 405 nm laser and collected in emission range 454-621 nm with master gain set at 650, digital gain 1.0, laser power 28%, and pinhole 35.5. Symbiodinium fluorescence was excited with the 405 nm laser and collected in emission range 655–718 nm with master gain set at 575, digital gain 1.0, laser power 28%, and pinhole 35.5.”

Fixed extra corals on 12/14/23

Just wanted to get some P. compressa and M. capitata in fixative before break. Transferred to stabilizer on 12/15/12 at 4PM in 4 ºC.

01/26/24 - Decalcified one P. compressa and one M. capitata from the 12/14/23 fixation

• Decalcified for ~72 hours (15:30 01/26/24 - 14:30 01/29/24)
  • skeleton almost all gone on 1/28/24

• Embedded without sucrose step on 01/29/24 - washed in DNAse/RNAse free H2O, then embedded using this protocol, with LN2.

• 02/01/24 - Sectioining new tissue and attempting to extract RNA from cryosections
  • sectioning protocol
  • NOTE: These were embedded without a sucrose cryoprotection step beforehand

Sectioning Porites compressa

Sectioning Montipora capitata, cut at -19 ºC

• 02/01/24 - Extracted RNA using CHARM kit, both with RIN ~5, here

• 2/6/24 - Stained one slide each from Pcomp and Mcap according to Hoescht staining protocol here

02/19/24 - Decalcified the other P. compressa and another M. capitata from the 12/14/23 fixation

• Trying to get tissue for tissue skirt imaging (see above) - to image on afternoon of 2/23/24
• But also getting tissue for sucrose cryoprotection and then embedding, but trying a 15% sucrose step first before 30% to see if slower sucrose infiltration helps maintain the tissue integrity
  • Slightly regret not also keeping some tissue for embedding without sucrose to compare RNA quality…
• Decalcified for ~96 hours (12:00 02/19/24 - 9:30 02/23/24)
  • skeleton almost all gone on 2/22/24 but I wanted to make sure all CaCO3 was gone for successful sectioning

Didn’t get any good montipora pictures, but Porites looked great:

• On 2/23/24:
  • washed for 15 mins in RNAse free PBS at 4 ºC on shaker
  • poured into petri dish with RNAse free PBS and disected each fragment (one per species) into two parts:
    • one went into 4 mL of RNAse free PBS at 4 ºC in the dark for tissue skirt imaging
• the other went into 10 mL of 15% sucrose in RNAse free PBS until the tissue sank,
  • P. comp: sank after 40 minutes in 15 % sucrose, after ~3 hours in 30% sucrose
  • M. cap: sank after 2.5 hours in 15% sucrose, ~4 hours in 30% sucrose
• then into 30 % sucrose in RNAse free PBS until the tissue sank

• then embedded on powdered dry ice

• 2/23/24: successfully imaged the Porites tissue “skirt”, both polyp side and underside. GFP & Chlorophyll channels

Sectioned on 3/18/24

• had a lot of issues with section curling, but otherwise the tissue cut much more nicely than last time - I think all the skeleton was successfully dissolved much better this time, and the sucrose embedding may have helped as well
• still some ripping of the tissue, though, which could be a temperature issue.

• will try to stain slides on 3/19/24 for morphology imaging and test RNA and DNA quality of sucrose-incubated POR/MON ASAP

• 3/22/24 - Stained one slide each from Pcomp and Mcap according to Hoescht staining protocol here