2024-08-28 LCM Experiment Sample Prep

Fixing and decalcifying Pocillopora for LCM Experiment

Protocol here

  1. Day 1 (8/28/24): Fixed tissue in PAXgene fixative (Pocillopora from CBLS Wetlab)
    1. Selected 5 plugs from crate 14, labelled A-E. Took photos with size standard of each.
    2. Pretty_Frags.png
    3. Frags_label_Size.png
    4. Then cut off branches and transfer gently to PAXgene fixative for 24 hours in fume hood, room temp
      1. First frag fixed at 16:42, last at 15:12 (so all within 20 minutes)
      2. 2 branches each in a 5 mL tube with 5 mL PAXgene fixative each
    5. Environmental conditions:
      1. Temp: 26.38 ºC
      2. pH: -73.2 mv (8.1)
      3. Salinity: 35.14 psu
  2. Day 2 (8/29/24): Replace fixative with stabilizer, transfer to 4 ºC
  3. Day 3 (Fri 8/30/24): Started decalcification, kept on shaker in cold room (4 ºC)
    1. First wash with ICE-COLD Rnase-free PBS to remove excess ethanol, then transfer to sterilized and RNA-se cleaned beaker or tube with ICE-COLD EDTA and put in cold room at 4 ºC.
  4. Day 4 (Sat 8/31/24): Changed EDTA solution (24 h)
  5. Day 5 (Sun 9/1/24): Still not decalcified, had to change the solution again. Moved to 6-well plate (48 h)
  6. Day 6 (Mon 9/2/24): Still not decalcified, had to change the solution again. (72 h)
  7. Day 7 (Tues 9/3/24): Morining: Transfer into 10 mL of 15% RNAse-free sucrose at 4 ºC in RNAse-free PBS until the tissue sinks
    1. Then into 30 % sucrose in RNAse-free PBS until tissue sinks –> then embed and store at -80 overnight
    2. Decalcification went from 5pm 8/30 to 8am 9/3 - about 87 hours.

Embedding Protocol here

  1. Day 7 (9/3/24): Embed on powdered dry ice and store at -80 ºC until sectioning
    1. Make sure to cool forceps on dry ice and OCT to 4ºC
    2. Make sure to fully dry tissue before embedding
    3. Let tissue sit in OCT for 2 minutes before freezing, to let it fully soak in
    4. Tightly wrapped in aluminum foil cleaned with Ethanol and RNAse away, then put in whirlpak and stored at - 80 ºC in a plastic box.
      1. Two branches per frag
    5. Cryomold, alumnium foil labelled with frag number, two wrapped molds per one labelled whirlpak per frag.

Sectioning and performing LCM on these branches, using the same steps outlined in this July post

Current timeline plan (will very likely need to change)

  1. 9/4/24: Sectioning, LCM, and extraction on Frag A (if time)
  2. 9/5/24: Sectioning on Frags B&C
  3. 9/6/24: LCM on Frags B&C; extraction if time
  4. 9/10/24: Sectioning on Frags D&E
  5. 9/11/24: LCM on Frags D&E; extraction if time

Modifying sectioning protocol to fix tissue to slides immediately after sectioning, following the PAXgene suggestions here

Modified Cresyl violet staining solution (from here)

  1. Prepare Cresyl violet staining solution at least one week prior usage
  2. Add 0.5 g Cresyl violet into 50 ml 100% ethanol
  3. Mix solution and store at 4°C sealed air-tight and dark
    1. mixed on orbital shaker overnight in cold room, with occasional vigorous shaking to resuspend powder into solution

Sectioning Procedure: 9/4/24 (Frag A), 9/5/24 (Frags B&C), 9/10/24 (Frags D&E)

  1. Section onto PEN membrane slide for LCM as well as extra slides for confocal morpholgy imaging and backup for RNA and DNA extraction
  2. All surfaces and equipment should be treated with RNAse cleaner!!!
  3. Morning of: prepare PEN slide with UV and RNAse cleaner
    1. Make sure to not damage or touch the membrane in any way
    2. Using sterilized and RNAse zap-ped forceps, dip slides in RNAse zap for 15 seconds
    3. Follow this by two 15 second rinses in DEPC water
    4. Let dry, at room temperature or at 37 ºC
    5. When visibly dry, place in UV box for 30 minutes (ideally do so immediately prior to sectioning)
    6. With clean, gloved hands, transfer to slide box for sectioning
  4. Prepare solutions:
    1. 70% Ethanol, 2 containers
    2. 100% Ethanol
  5. Sectioning procedure:
    1. Keep slide cold in the cryostat, briefly warm sections with finger to adhere
    2. In clean petri dish, pipette ice-cold 100% ethanol onto the sections and let sit for 30s.
    3. Air-dry the slide for 15 min at room temperature (15–25°C) in dessication chamber or falcon tube with dessicant.
    4. 70% Ethanol, 1 minute (some protocols say 2 min)
    5. Transfer the slide to ice-cold 100% ethanol for transport to lab
  6. In lab: Staining Procedure (modified/combined from here and here to reduce exposure to aqueous solutions of lower than 70% ethanol to reduce RNAses) (ALSO)
    1. ALL SOLUTIONS ICE COLD
      1. Place slide on petri dish
      2. Apply Cresyl violet staining solution (in 100% ethanol) directly with syringe and sterile filter to the section and incubate for 1 minute, swivel gently
      3. Dip for 5 seconds in 70% ethanol (dip several times to remove all OCT)
        1. Repeat with fresh container of 70% ethanol
        2. may need to dunk slide in RNAse-free H2O to remove all OCT. If so it needs to be very quick and ice cold and then immedately dunked in cold ethanol again.
      4. Dip for 5 seconds in 100% ethanol (dip a few times)
        1. Can fix up to 2 minutes in 100% ethanol or even for short-term storage prior to LCM
      5. Air dry sample 1-2 in drying chamber with desiccant or fume hood
        1. if possible, proceed to immediate LCM at this step!
  7. Freeze slides at -80 ºC

LCM Procedure: 9/4/24 (Frag A), 9/6/24 (Frags B&C), 9/11/24 (Frags D&E)

  1. Bring to LCM
    1. Lab notebook
    2. 70% ethanol and RNAse away
    3. Kimwipes + some paper towels
    4. Gloves
    5. p200 + tips
    6. Dry ice for tubes (& slides when done dissecting)
    7. RNAse free 0.2 mL PCR tubes (I keep one rack with clean closed tubes at RT and one in the dry ice cooler)
    8. Buffers to collect cells in, bring to LCM and fill tube caps with 40 uL of solution when loading onto scope:
      1. Zymo Proteinase K digestion buffer (95 uL 2X, 95 uL H2O) + proteinase K (10 uL)
      2. Charm DD1 frozen tissue buffer
    9. LCM slides prepared above
  2. Clean working space, and all parts of scope around the slide and sample collectors (also computer mouse and focus/stage knobs) with 70% ethanol and RNAse cleaner. Do so carefully for scope with damp kimwipe, do not spray anything onto the equipment.
  3. Unload the slide holder and clean with ethanol and RNAse away
  4. Load slide onto slide holder (thick part with label goes towards the spring holder)
    1. Slide should be loaded right-side up, with tissue on the bottom
  5. Unload the collector and clean with ethanol and RNAse away
  6. Label 4 PCR tubes and put in the collector
  7. Fill caps of each tube with appropriate buffer, recorded in lab notebook
    1. Note order carefully and know which tube is A, B, C, & D on the LCM
  8. Load collector onto scope and get crackin!
  9. Windows to pull up: Laser, microscope control
  10. Take overview image of slide at 5X
  11. Dissect cells at 20X. Find areas (typically not in the polyp where there are many sections through the tissue and oral vs. aboral is harder to tease apart) where you can clearly differentiate oral epidermis and aboral tissue. For each section, choose a tube that will collect the oral epidermis and the tube that will collect aboral epi/gastrodermis. Record this carefully! When drawing areas to dissect, select the correct tube in the bottom left and make sure the ROI drawing is the same color as the tube color. You can draw multiple ROIs for different tubes in one view, just make sure to switch tubes between ROIs. And when going from one view/dissection to the next, always confirm the tube selected for collection before dissecting.
    1. Make sure scale bar is on
    2. Take image before dissecting, with ROI in view (SAVE + L), and name the image with the tube # and the dissection #
    3. The program records the area in um^2 for the ROI drawn, and I record this in my lab notebook for each dissection number to get a general idea of how much tissue has been dissected per tube (However, the ROI area doesn’t necessarily equal tissue area!!)
    4. Keep dissecting along the section , with the appropriate tissue going into the appropriate tube (and taking pictures/writing notes for each cut) until you have enough tissue for RNA extraction (for POC, I am estimating this will be around 7-8 20X dissections)
    5. Try to work as quickly as possible. Once a section is done, unload the collector and remove the tubes with the dissections VERY CAREFULLY, closing the cap as you remove it. Spin down in the mini centrifuge, then vortex 5-10s, then spin down for at least 30s. Then place in tube rack on dry ice.
  12. After a slide is done, I place it back in the falcon tube with dessicant and transport on dry ice to -80 for storage.

Sectioning and LCM Notes, Day 1 (9/4/24)

  1. Sectioining: Today went much harder than expected. I was having issues during sectioning with the tissue coming out of the OCT, and some rolling issues as well. Sectioined two slides - labelled A1 and A2.
  2. Even though I was very careful during all staining and washes (pipetting solutions only, no dunking), and did not notice any visible tissue loss, the tissue quality did not look great on the LCM. Collected 6 samples total (2 each from 3 sections on slide A2); focusing on collecting external epidermis (possibly difficult to disentangle from gastrodermis) and internal calicodermis. I pooled dissections within one section only all into the same tube for each tissue layer/section.
  3. I was having issues with static. I performed the LCM with the tissue facing upward for better visualization, which could have contributed to the static? It could also just be an issue of cutting small dissections and them not being heavy enough to fall well.
  4. But, I got samples. It took a long time. But there are some. But this replicate may need to be redone
  5. Samples collected (all from the second slide from this morning, A2)
    1. Section 1
      1. Tube 1: Aboral tissue, 7 dissections (dissections 2 and 8 lost)
        1. 108,479 μm2
      2. Tube 2 (did not end up using for dissections)
      3. Tube 3: Oral epidermis, 6 dissections
        1. 99,914 μm2
    2. Section 2
      1. Tube 4: Aboral tissue, 9 dissections
        1. 133,602 μm2
      2. Tube 5: Oral epidermis, 6 dissections
        1. 139,106 μm2
    3. Section 3
      1. Tube 6: Oral epidermis, 4 dissections
        1. 100,191 μm2
      2. Tube 7: Aboral tissue, 4 dissections
        1. 71,444 μm2
  6. LCM lasted from 1:30-4:30. Needs to be faster.

Example photo:

TUBE3_4.jpeg TUBE3_4_POST.jpeg

LCM Protocol adjustment Day 2 (9/6/24)

  1. I am modifying the staining protocol for today. After sectioining, the slides were dried at room temp for 1 miniute, and then fixed at the cryostat with ice-cold 100% ethanol for 2 minutes. Then this was removed, sides of the slide were blotted with a kimwipe, and I transferred the slide to a falcon tube with dessicant and let this dry at room temp for 15 minutes. Then transported back to the lab on dry ice and transferred to -80 ºC for overnight.
  2. Morning of LCM: Bring slide up to room temperature, slowly to avoid formation of water condensation inside the container. Did the following:
    1. 30 minutes at -20 ºC
    2. 30 minutes at 4°C
    3. 15 minutes at room temp
    4. The performed staining with cresyl violet
      1. ALL SOLUTIONS ICE COLD
      2. Place slide on petri dish; on tube rack over dry ice (not immediately on the dry ice otherwise the 70% ethanol can freeze)
      3. Apply Cresyl violet staining solution (in 100% ethanol) directly with syringe and sterile filter to the section and incubate for 1 minute, swivel gently
      4. Rinse (pipetting) with ice-cold 75% ethanol
      5. And then place back on clean petri dish and cover slide with ice-cold 70% ethanol (ideally the ethanol just stays on the slide but if it rolls off then gently submerge the slide in 75% ethanol)
        1. Ideally this removes all the OCT. And hopefully no tissue.
      6. Be as gentle as possible and keep everything as cold as possible
      7. If successful, we should hopefully be able to skip the H2O step.
      8. Final fix, 1 minute in 100% ethanol or even for short-term storage prior to LCM
      9. Air dry sample 1-2 in drying chamber with desiccant or fume hood
        1. Proceed to LCM now.

Sectioning and LCM Notes, Day 2 (9/6/24, tissue sectioned 9/5/24)

  1. Sectioning went better than yesterday. I was able to get the brush method to work pretty well for sections at -22ºC. I got one slide per frag (B&C), with four sections each. There was definitely still a little bit of folding and rollage in some cases during section transfer, but overall I think better than yesterday.
  2. LCM went okay. The tissue looked much better than 9/4/24, so the sectioning was definitely improved. There was more intact tissue with the adjusted staining protocol, but there was also lots of OCT left on the slide, which is a problem. However, maybe having to cut through extra OCT is the price to pay for intact tissue, especially with these very delicate layers.
  3. Got one slide each for Frags B and C, and collected two sets of samples for Frag B and three for Frag C.
  4. LCM lasted about 5 hours, from 11 to 4.

Example photo:

TUBE17_3.jpg

Frag B Samples collected

  1. Section 1
    1. Tube 8: Oral epidermis, 8 dissections
      1. 157,063 μm2
    2. Tube 9: Aboral tissue, 7 dissections
      1. 99,063 μm2
    3. Tube 10: Symbionts only, 17 dissections
      1. 156,815 μm2
  2. Section 3
    1. Tube 11: Oral epidermis, 3 dissections
      1. 78,431 μm2
    2. Tube 12: Aboral tissue, 3 dissections
      1. 42,643 μm2

Frag C Samples collected

  1. Section 3
    1. Tube 13: Oral epidermis, 7 dissections
      1. 282,917 μm2
    2. Tube 14: Aboral tissue, 7 dissections
      1. 192,027 μm2
  2. Section 1
    1. Tube 15: Oral epidermis, 8 dissections
      1. 470,189 μm2
    2. Tube 16: Aboral tissue, 5 dissections
      1. 300,731 μm2
  3. Section 4
    1. Tube 17: Oral epidermis, 7 dissections
      1. 364,858 μm2
    2. Tube 18: Aboral tissue, 4 dissections
      1. 194,848 μm2

Sectioning and LCM Notes, Day 3 (9/11/24, tissue sectioned 9/10/24)

  1. Sectioning went well, used mixture of brush method and roll plate. Sectionined at -22ºC. One slide each for frags D&E.
  2. I added the RNAse-free water rinse back into the staining protocol today to remove excess OCT. All was done over dry ice and kept very cold. The cresyl violet stain took better in the tissues today than 9/6/24.
  3. LCM went much better, I flipped the orientation of my slides on the LCM (tissue on bottom instead of on top) and it seemed that the focal distance for images and ability of the laser to cut were much, much better. Barely had to re-cut any dissections or adjust the laser power, which helped with speed and likely dissection quality
  4. Got one slide each for Frags D and E, and collected two sets of samples for Frag D and four for Frag E.
  5. LCM lasted about 4.5 hours, from 11:30 to 4.

Example photo:

TUBE27_4.jpeg TUBE26_5A.jpeg

Frag D Samples collected

  1. Section 2
    1. Tube 20: Oral epidermis, 7 dissections
      1. 297,944 μm2
    2. Tube 21: Aboral tissue, 8 dissections
      1. 217,521 μm2
    3. Tube 22: Cnidocytes only, 16 dissections
      1. 103,883 μm2
  2. Section 1
    1. Tube 24: Oral epidermis, 9 dissections
      1. 313,540 μm2
    2. Tube 25: Aboral tissue, 9 dissections
      1. 292,276 μm2

Frag E Samples collected

  1. Section 2
    1. Tube 26: Oral epidermis, 10 dissections
      1. 387,947 μm2
    2. Tube 27: Aboral tissue, 11 dissections
      1. 371,129 μm2
  2. Section 4
    1. Tube 28: Oral epidermis, 13 dissections
      1. 467,440 μm2
    2. Tube 29: Aboral tissue, 15 dissections
      1. 517,157 μm2
  3. Section 3
    1. Tube 30: Oral epidermis, 5 dissections
      1. 217,426 μm2
    2. Tube 31: Aboral tissue, 5 dissections
      1. 182,599 μm2
  4. Section 1
    1. Tube 32: Oral epidermis, 7 dissections
      1. 289,071 μm2
    2. Tube 33: Aboral tissue, 7 dissections
      1. 307,806 μm2
Written on August 28, 2024