Zebrafish Care and Experimental Techniques: In Situ Hybridization - Zebrafish Sections

-

  1. **Day 0:

    Goals: To preserve zebrafish and obtain sections using the cryostat.
    **Everything used on this day needs to be RNase free. Wear gloves and be sure to use only PBS that has been treated with DEPC.

    1. Anesthetize zebrafish

    2. Fix in freshly thawed BT fix for 3 hours at room temperature. Keep solution on the "belly dancer" so that it is constantly moving.

    3. Rinse 3 x 5 minutes in 1X PBS. Leave embryos in pbs until you are ready to section them. Embryos can be stored in the fridge until sectioning, but keep the days between fixing, sectioning and staining as short as possible.

    4. Rinse fish in 30% sucrose and PBS until embryos sink approx. 30mins.

    5. Trnsfer embryos from sucrose to plastic molds, 5 fish pre mold, with heads facing the side with writing. Absorb as much sucrose as possible.

    6. Add OCT to the mold, and swirl fish until they are oriented correctly. Freeze in -80 degree freezer.

    7. Follow instructions for sectioning using the cyrostat.



    **Day 1:

    Goals: Permeabilize zebrafish tissue, block non-specific binding, and bind specific probes (GFAP and slitrk probes)

    **Note: Everything used on this day needs to be RNAse free!!! Wear gloves and use special tips.  Keep Day 1 glassware separate from all other glassware. There are three Coplin jars labeled Day 1 – one is labeled for TEA and the other two are used for PBST rinses. Keep them labeled and cover with parafilm when not in use and only use for Day 1.

    Steps 1-9 should be performed in the hood. All wash steps are performed in Coplin jars. Each jar holds about 50 mL of liquid.

    1. Thaw sections for 15 minutes at Room Temperature

    2. Fix in BT Fix for 10 minutes in hood. You will want to take a tube of BT Fix out of the freezer and place in fix fridge to thaw the day before you start this protocol. Overlay slides with about 1 mL of fix. Might have to use pipette tip gently spread fix so that it covers the whole slide. While slides are fixing mix PBST.

    3. Wash 2 x 5 minutes in PBST to rinse off fix from previous step. While slides are rinsing mix proK.

    4. Add Proteinase K  (ProK) 10 ug/mL for 10 minutes to permeabilize the tissue. Overlay slides with about 0.5 mL ProK. The timing on this step is critical, so do not let it go over time. If you treat with ProK too long the cells can disintegrate or all the RNA could go out of them.

    5. Wash 2 x 5’ in 1X PBST to rinse off ProK
    6. Fix again in BT fix 10 minutes (in hood).  Overlay slides with about 0.5 mL. This is to preserve the tissue in the permeabilized state.

    7. Wash 2 x 5 minutes in 1X PBST. During this step, pre-warm the hybridization oven and HM (hybridization media) in dry bath incubator 50 C and mix TEA.

    8. Acetylation for 10 minutes in a Coplin jar (pre-mix by adding 125 ul acetic anhydride (located in the “Flammable” cabinet under the hood) to 50 mls 0.1 M TEA. Mix in a 50 mL conical tube in hood.) This step eliminates the charge on the superfrost plus slides.

    9. Wash 2 x 5 minutes in 1X PBST

    10. Pre-hybe with HM for 2 hours in 50 C hybridization oven to block non-specific binding. Overlay each slide with 400 μl HM.

    11. Add fresh HM + probe to the slides and keep in 50 C oven overnight (1:1000 dilution). Use the GFAP probe as a positive control and then use other slitrk probes on the remaining slides depending on the experiment. During this step the slitrk and GFAP probes are binding to the RNA and move PBST from Day 1 bottle into PBST Day 2 and # Bottle.

    Day 2:

    Goals: Wash off unbound probe from Day 1, block non-specifc binding, and bind antibody to probe that is bound to RNA.

    Steps 1-6 performed in Coplin jars.

    1. Wash 15 minutes in 2X SSC (dilute from 20X stock) at 50 C (pre-warm solution in water bath; turn water bath on about 20 minutes before needed)

    2. Wash 5 minutes in 2X SSC at Room Temperature

    3. Add 20 ug/ml RNAse A (dilute in 2X SSC) for 30 minutes at 37 C. Use the hybridization oven for this step and overlay the slides with about 0.5 mL.

    4. Wash 5 minutes in 2X SSC at Room Temperature

    5. Wash 2 x 30 minutes in 0.2X SSC (dilute from 20X stock) at 50 C (pre-warm solution by placing in water bath ahead of time)

    6. Wash 2 x 10 minutes in 1X PBST at Room Temperature

    7. Add Blocking Solution (1X PBST and 10% sheep serum) for 1 hour at Room Temperature in the humid chamber. Overlay each slide with about 1 mL of blocking solution.

    8. Add Blocking Solution + anti-DIG-AP antibody at 1:5000 overnight at 4 C. Overlay each slide with about 1 mL and keep in humid chamber. Place humid chamber in the fix fridge.

    Day 3:

    Goals: Wash off unbound antibody from Day 2. Perform reaction to reveal staining pattern in sections.

    1. Discard liquid on slides

    2. Wash 4 x 15 minutes in 1X PBST. These washes are rinsing off the unbound antibody from Day 2.

    3. Wash 2 x 5 minutes in 100 mM Tris pH=9.5 (Dilute from 1 M stock). The tablets in the next step contain tris, so this step is acclimating the tissue to that solution.

    4. During step 3 combine 5 ml Millipore H2O and ½ of a NBT-BCIP tablet in a 15 mL conical tube. Shake well to dissolve tablet. Keep in a drawer away from light.

    5. Add about 500 μl solution from step 4 to each slide in the humid chamber. Keep at Room Temperature in the dark. Easiest to store humid chamber in a cubby/cabinet with a door.

    6. Check slides under microscope at the end of the day for staining pattern. Slides can be left for up to two days before stopping the staining reaction. If you are leaving slides longer than one day, change the staining solution after 24 hours. Rinse slides in 100 mM Tris for 5 minutes then add fresh staining solution to each slide.

    7. To stop the staining reaction, wash slides 2 x 5 minutes in 1X PBS. (Insert pictures of successful staining we have seen so far.)

    8. Add mounting medium and coverslip. Store in a slide box at 4 C (fix fridge).
  2.  
    See the drop box files for staining worksheet (with recipes). Use a staining worksheet each time you complete a stain, making notes about that particular trial. Put the worksheet in the " In Situ Zebra Fish Protocol Worksheet" binder. If you uncover something significant that is helpful or requires a change in protocol, make note of it on the worksheet in an obvious way.

Comments

Post a Comment

Popular posts from this blog

Prepare 6X Laemmli buffer

-
Laemmli is a sample buffer to use in western blot. 6X sample buffer is added to each protein sample and is boiled or heated for 5-10 minutes. Recipe to prepare 10 ml: - 1.2gr SDS (sodium dodecyl sulfate) - 6mg bromophenol blue - 4.7ml glycerol - 1.2ml Tris 0.5M pH6.8 - 2.1ml dwater warm it a little bit and shake it till everything is dissolved. It takes time. - add 0.93gr DTT. Wait until it is completely dissolved. Aliquote and keep frozen at-20ºC.
Read more

0.1 M Triethanolamine

-
0.1 M TEA: TEA = Triethanolamine You will dilute TEA 1:10 in Millipore water: Pour 50 mL TEA into a 50 mL conical tube (TEA is very viscuous, so best to measure it in something that is disposable). Pour TEA from conical tube into 450 mL Millipore water with stirring. pH to 8.0
Read more

100% Steinberg’s Solution

-
100% Steinberg’s Solution (AKA 5xMR) (makes 2 liters) 40 ml NaCl stock solution (17% w/v NaCl) 20 ml KCl stock solution (0.5% w/v KCl) 20 ml CaCl2 stock solution (0.5% w/v CaCl2) 20 ml MgSO4 stock solution (1.025% w/v MgSO4) 1 ml gentamycin stock solution (4% w/v gentamycin) 2.4 g HEPES 1899 ml ddH2O Adjust pH to 7.4 - 7.6 and store in refrigerator. Usually, you will make 2-4 liters at a time. To make the required stock solutions,
Read more