Testing PAXgene, PFA, (and Z-fix) for Immunofluorescence, FISH + LCM preps and sectioning

Immunofluorescence and FISH Preparation: Testing PAXgene, PFA, (and Z-fix) for FISH + LCM preps and cryosectioning

First steps, August 2023

  1. Use paxgene-fixed and decalcified samples from April to test cryosectioning protocol, Hoescht staining, and confocal imaging of PAXgene-fixed tissues. If these results are promising, begin fixing more coral tissues in PAXgene (or PFA to save money), decalcifying, embedding, and perfect sectioning protocol and begin testing staining / LCM.

    1. (Can also try non-cryosectioning protocol (Katie Barrot Protocol): Try fixing in PFA as detailed exactly in the protocol below and go through all steps for sectioning and slide preparation, and do the same in parallel for PAXgene fixed tissues)
    2. With validated antibody or probes (talk to Hollie), try prepping some slides for FISH/Immunofluorescence and keep some to prepare the same way but for LCM.

Supplies List

Helpful protocol and video for FISH

Cryosectioning Protocol

see updated protocol here

The following protocol is the protocol written by Katie L. Barott and Martin Tresguerres, which was based in part on the protocol by Puverel et al. (2004)

Barott, K. L. and Tresguerres, M. (2015). Immunolocalization of Proteins in Corals: the V-type H+-ATPase Proton Pump. Bio-protocol 5(17): e1573. DOI: 10.21769/BioProtoc.1573.

Materials and Reagents

  1. Live coral
  2. Electron microscopy grade paraformaldehyde (16% solution in 10 ml ampules) (Electron Microscopy Sciences, catalog number: 15700 or 15710 )
  3. Ethyl alcohol (absolute, 200 proof)
  4. Ethyl alcohol (190 proof)
  5. Xylenes (any reagent ≥ 98.5% xylenes, other tissue clearing agents such as Safeclear, are also acceptable) (Safeclear, catalog number: 044-192 )
  6. Paraffin embedding medium (e.g. Paraplast X-TRA) (McCormick Scientific, catalog number: 39503002 )
  7. Triton-X-100 (Bio-Rad Laboratories, AbD Serotec®, catalog number: 1610407 or Sigma-Aldrich, catalog number: T8787 )
  8. Normal goat serum (Vector laboratories, catalog number: S-1000 )
  9. Keyhole Limpet Hemocyanin (Sigma-Aldrich, catalog number: 7017 )
  10. Phosphate buffered saline (PBS) (e.g. 10x) (Corning Incorporated, catalog number: 46-013-CM )
  11. Microscope slides and coverslips
  12. Custom, antigen-affinity purified, rabbit polyclonal anti-VHAB antibodies developed using a peptide antigen matching a conserved region of the VHAB subunit (AREEVPGRRGFPGY) (Genscript USA, Inc.). The epitope is 100% conserved among diverse animal species, and specifically recognizes VHAB from coral (Barott et al., 2015), bone-eating worms (Tresguerres et al., 2013), hagfish (Clifford, et al. 2015), and sharks (Tresguerres et al., 2010; Roa et al., 2014).
    Note: This protocol may be adapted for other proteins following validation of the primary antibodies in the coral species to be examined. However, successful localization using different primary antibodies may require optimization of the fixation procedure from that described below.
  13. Secondary antibodies: Goat anti-rabbit IgG Alexa Fluor 555 (2 mg/ml) (Life Technologies, catalog number: A21429 )
    Note: The Alexa555 fluorophore is used to avoid overlap with the coral’s endogenous green fluorescent protein (GFP), but other fluorophores outside of the GFP emission range may also work.
  14. Hoescht 33342 (Life Technologies, catalog number: H1399 )
  15. Mounting medium (e.g. Electron Microscopy Sciences, catalog number: 17895 )
    Note: Unless otherwise noted, reagents can be obtained from any general laboratory reagent manufacturer or supplier.
  16. S22 buffer (see Recipes)
  17. Ca-free S22 buffer + 0.5 M EDTA (see Recipes)
  18. Fixative solution (see Recipes)
  19. Blocking buffer (see Recipes)

Equipment

  1. Chemical fume hood
  2. Fluorescence microscope (e.g. Zeiss AxioObserver Z1, or any other microscope with the ability to detect blue (“DAPI filter”, peak excitation 350 nm; peak emission 461 nm; for Hoescht staining of nuclei) and red (“TRITC filter”, peak excitation 550 nm; peak emission 570 nm; for secondary antibodies) fluorescence.
  3. Optional
    • Coral endogenous GFP and chlorophyll from dinoflagellate symbionts (Symbiodinum) are visible using settings for “green fluorescence” (“FITC filter”, peak excitation 490 nm; peak emission 525 nm).
    • Structured illumination (Zeiss Apotome2), confocal capabilities *suggested but not required.
    • Differential Interference Contrast (DIC) (Nomarski microscopy) *suggested but not required.
  4. Microcentrifuge [any model capable of spinning 1.5 ml tubes at 3,000 x g at 4 ℃
    Note: This temperature can be achieved by placing the microcentrifuge inside of a fridge.
  5. Tissue cassettes (e.g. C&A Scientific, catalog number: EC-0109x )
  6. Tissue embedding station or any other method to pour liquid paraffin (e.g. Kedee, model: KD-BM II )
  7. Embedding base molds (metal) (e.g. C&A Scientific, catalog number: HB-07 , 15 , 24 , 30 )
  8. Embedding rings (plastic) (e.g. Biologix, catalog number: 41-3004 )
  9. Microtome (any model capable of cutting 5-30 μm paraffin sections)
  10. Slide warmer/hot plate (any model capable of warming up slides to 30 ℃)
  11. Hydrobarrier pen (Pap pen) (e.g. Vector Laboratories, catalog number: H-4000 )
  12. Rotating platform (any rotator/shaker/rocker)

Procedure

A. Tissue preparation
Note: Section A takes approximately 10-14 days when completed without stopping. There are several stopping places within Section A.

  1. Immerse a live coral fragment (~1-2 cm in length) in fixative solution [S22 buffer with 3% paraformaldehyde (PFA), see Recipes] and incubate overnight at 4 °C (Puverel et al., 2005).
    • Notes:
    • PFA stock solution should be handled in a chemical fume hood.
    • It is best to add PFA stock solution (16% or 32% PFA) to S22 buffer immediately before use. Diluted PFA can be stored at 4 °C for up to one week. Stock PFA may be stored at 4 °C for up to one month or at -20 °C for 6 months once unsealed.

  2. Decalcify by transferring fragment to Ca-free S22 buffer with 0.5 M EDTA and 0.5% paraformaldehyde and incubate at 4 °C. Replace the buffer daily until the skeleton is completely dissolved (~7 days depending on size of fragment and thickness/density of skeleton). Dissolution of the skeleton is determined visually; the coral tissue should appear transparent with no visible white calcium carbonate remaining

    Note: It is best to add PFA stock solution (16% or 32% PFA) to Ca-free S22 buffer immediately before use.

  3. Dehydrate tissue.
    Note: Incubations in steps 3.1-7 are conducted on a shaking platform.
    1. Incubate in 50% ethanol for 2-5 h.
    2. Incubate in 70% ethanol overnight at 4 °C.
      • Tissue will keep in 70% ethanol at 4 °C for months.
    3. Put tissue in mesh plastic tissue cassette for final incubation steps in glass beaker.
    4. Incubate in 95% ethanol for 20 min.
    5. Incubate in 100% ethanol for 20 min. Repeat with fresh ethanol for a total of three times.
    6. Incubate in xylene for 20 min. Repeat with fresh xylene for a total of three times.
      • Do in HOOD.
    7. Incubate in paraffin for 30 min. Repeat with fresh paraffin for a total of three times.
  4. Embed tissue in paraffin wax.
    1. Transfer tissue to base mold and position as desired.
      • Fill with wax.
    2. Place embedding ring on top of base mold, and fill with wax to the top.
    3. Let wax solidify at room temperature for 1-2 days.
    4. Embedded tissue may be stored at room temperature* for several months prior to sectioning; long-term, archival storage must be done at 4 °C (*but storage at 4 °C is recommended).
  5. Sectioning.
    1. Turn on hot plate to 40 °C.
    2. Trim wax block around the coral tissue into the shape of a trapezoid (i.e. so top and bottom are parallel, bottom edge longer, and left half square; Figure 2).
    3. Load sample onto microtome.
    4. Load blade with holder.
      • Blade stored at -20 °C.
      • Blade set at 15 degree angle (may be adjusted).
    5. Unlock stage and slide until blade almost touching sample.
    6. Start cutting sample using thick (15-30 μm) sections until surface is flat and smooth and tissue is reached.
    7. Switch to desired tissue thickness for slides (5-10 μm).
      • Should get ribbon of consecutive sections (each section is easily identifiable because the right edge of each section is not parallel to the left edge of the following section).
    8. Cut ribbon into individual sections with razor blade and place onto glass microscope slides.
      • Blade-cut side down.
      • Label slides with pencil.
    9. Float section on water, remove any bubbles (two or three consecutive sections can be placed on each slide).
    10. Stretch wax section by placing slide on hot plate (~30 sec). Surface should become smooth.
      • Remove water with Kim wipe.
    11. Let dry/affix to slide by leaving slide on hotplate overnight at 30 °C with paper towel underneath.
    12. Store at 4 °C until use.
    Deparaffinization and rehydration.
    1. Incubate in xylene bath for 10 min.
      • Repeat with fresh xylene for a total of three times.
      • Do in HOOD.
    2. Incubate in 100% ethanol bath for 10 min.
    3. Incubate in 95% ethanol bath for 10 min.
    4. Incubate in 70% ethanol bath for 10 min.
    5. Incubate in PBS-TX (1x PBS with 0.2% (v/v) Triton-X-100) bath for 10 min.

Recipes

  1. S22 buffer
    1. 450 mM NaCl
    2. 10 mM KCl
    3. 58 mM MgCl2
    4. 10 mM CaCl2
    5. 100 mM Hepes (pH 7.8)
      Note: Adjust pH of Hepes buffer with NaOH.
  2. Ca-free S22 buffer + 0.5 M EDTA
    1. 450 mM NaCl
    2. 10 mM KCl
    3. 58 mM MgCl2
    4. 0.5 M EDTA
    5. 100 mM Hepes (pH 7.8)
      Note: Adjust pH of Hepes buffer with NaOH.
  3. Fixative solution (10 ml)
    1. Break open PFA ampule (16% PFA stock solution).
    2. Add 1.9 ml PFA stock solution to 8.1 ml S22 buffer (3% PFA final concentration). Invert to mix.
      Notes:
      1. Remaining PFA stock solution can be stored at 4 °C for up to one month or at -20 °C for 6 months.
      2. Fixative solution can be stored at 4 °C for up to one week.
  4. Blocking buffer
    1. 4 ml PBS-0.2% TritonX
    2. 80 µl NGS (normal goat serum)
    3. 0.8 µl KLH (Keyhole Limpet Hemocyanin) solution

My protocol / plan:

  1. Try fixing in PFA as detailed exactly in the protocol below and go through all steps for sectioning and slide preparation, and do the same in parallel for PAXgene fixed tissues
  2. With validated antibody or probes (talk to Hollie), try prepping some slides for FISH/Immunofluorescence and keep some to prepare the same way but for LCM.
    • We may need to go the cryosectioning route instead of paraffin embedding, so an alternaitve protocol using cryosectioning is also presented below.

Supplies List: https://docs.google.com/spreadsheets/d/1KlfFGhXL9WP4ChCF0b–V7Zvu5qfZ8f7ct2V04dX2rA/edit?usp=sharing

Helpful protocol and video for FISH: https://www.jove.com/v/57853/in-situ-hybridization-techniques-for-paraffin-embedded-adult-coral

https://link.springer.com/protocol/10.1007/978-1-0716-2172-1_19

Immunofluoresence of TRP channels in nematostella: https://doi.org/10.1242/jeb.243717

Written on August 14, 2023