In Situ HYBRIDIZATION TO TISSUE SECTIONS

<center> <font><font><strong>In Situ HYBRIDIZATION TO TISSUE SECTIONS</strong><br /> (Kornberg et al., Cell 40:45, 1985))</font> </font></center>

PARAFFIN SECTIONS
Preparation of embryos (¾10 hr) for paraffin sections

• Wash embryos off collection plates onto a nylon screen with 0.4% NaCl/0.3% Triton X-100. Rinse well to remove all food and yeast. Blot well to remove excess liquid.
• Dechorionate embryos in freshly diluted 50% chlorox for 3' at room temperature with gentle agitation. Rinse with NaCl/Triton; blot well.
• Transfer dechorionated embryos to either buffer A or B (depending on their age):
  • Buffer A (0-2, 1-3 hr embryos)
  • 2.5 ml 8% paraformaldehyde (fresh)
  • 2.0 ml 1X PBS
  • 0.5 ml DMSO
  - or -
  • Buffer B (2-4, 3-5 hr, etc.)
  • 2.5 ml 8% paraformaldehyde
  • 0.5 ml 1X PBS
  • 2.0 ml DMSO
• Add 5 ml heptane and shake vigorously for 20' (Buffer A) or 30' (Buffer B).
• Transfer to 10 ml 90% MeOH/EGTA (9ml MeOH + 1 ml 50mM EGTA, pH 8) +10 ml heptane
  • The MeOH/EGTA/heptane mix should be prechilled to -70 degrees C on dry ice. It takes awhile to come down to temp. so place it on dry ice about 1 hr earlier.
  • One way to transfer the embryos to the cold mix is to aspirate off the heptane (top) and paraformaldehyde (bottom) phases (the embryos will be at the interface), pour the embryos out onto a screen and harvest the embryos with a paint brush.
  • Alternatively, remove both phases completely from the original tube and then pour the cold mix onto the embryos.
  Shake vigorously on dry ice for: 10' (0-2, 1-3, 3-5, 4-6 hr embryos) 15' (5-7, 6-8 and 8-10 hr embryos)
• Rapidly warm the flask by swirling under hot running tap water. The devittellinized embryos will sink to the bottom. Transfer the embryos to a vial with a wide bore pipet tip. Wash 3X with 90% MeOH/EGTA.
• Rehydrate embryos (0-2 and 1-3 hr embryos need not be rehydrated through paraformaldehyde, just leave them in the appropriate MeOH/PBS solutions), 10' in each solution:
  MeOH/EGTA : 1X PBS : 4% Paraformaldehyde 9 : 1 : 0
  9 : 0 : 1
  7 : 3 : 0
  7 : 0 : 3
  5 : 5 : 0
  5 : 0 : 5
  3 : 7 : 0
  3 : 0 : 7
  
• Fix an additional 15' in 4% paraformaldehyde without agitation. At this point, embryos may be stained with DAPI or Fluorochromes. For sectioning, rinse thoroughly with PBS.
• Dehydrate through 30%, 50% and 70% EtOH; 10' in each solution (embryos may be stored at this point at -20 degrees C for at least a week).

Embedding embryos in paraffin

• Complete the dehydration:
  90% EtOH, 10'
  95% EtOH, 10'
  100% EtOH, 10'
  100% EtOH, 10'
  Xylenes, 10'
  Xylenes, 10'
  Transfer the embryos to a sieve and pat dry. The sieve may be put in a beaker with xylene (because once the embyos are in xylene they are hard to see).
• Transfer to Xylene/paraffin (1:1) @ 58 degrees C and allow them to sit in this mixture for 20' (embryos ¾ 6 hr) or 30' (embryos > 6 hr). Be careful to not let the temp. drop below 56 degrees C or the wax will harden rapidly. Do not let the temp. go above 62 degrees C or the wax will not polymerize correctly!
• Place embryos in the mold (Scientific Prod. 7275-1; peel-away disposable, 8x8mm):
  • Cut off the end of a Pasteur pipette and heat it briefly in a flame. Plunge into melted wax to lower the temperature of the pipette.
  • Draw up as many embryos as possible in as small a volume as possible using the warmed pipette.
  • Expel the embryos quickly and carefully onto the base of a preheated plastic embedding mold. Keep the drop in the center of the mold. If it touches the edges (and it will!), swirl the mold so that the embryos will settle in its center.
  • Allow the embryos to settle for about 10'. When they are almost longitudinal, remove the mold from 58 degrees C.
  • Watch the wax set. When a skin develops on the top, quickly pour melted wax down the side of the mold until it is at least half full. Let it harden. If you pour the wax in too soon, the embryos will be unsettled (and therefore no longer in the cen- ter nor longitudinally oriented). If you pour the wax in too late, the layer containing the embryos will not stick to the rest of the block, making sectioning impossible.
• Blocks may be stored at 4 degrees C for at least 6 months and probably longer.

Cutting paraffin sections

• Trim the block to a trapezoidal shape with a razor blade (try to cut off as little as possi- ble). The trapezoidal shape facilitates later separation of sections, one from another.
• Mount the block onto a wooden specimen holder using melted wax (just melt the shavings from the trimmed block with a hot spatula and quickly press the block into the pool of melted wax). Let it harden about 5'.
• Mount the specimen holder onto the microtome with the smaller edge on top.
• Cut sections. It is possible to make sections 4 microns thick, but they tear easily. Six micron sections are easily managed. The knife should be thoroughly cleaned with xylene in between blocks. Doing this will prevent many problems
  Troubleshooting
  
  • Possible problem: Sections tear as they are sliced.
    Potential remedies: There may be a piece of paraffin dust on the knife. Remove it with a brush. Or there may be a nick on the blade. Change the position of the knife.
  • Possible problem: Sections stick to the top of the block as the microtome returns to its starting position.
    Potential remedies: The blade may be dirty. Carefully (with upward strokes) clean it with xylene. Or the knife may be at a bad angle to the block. Adjust it until the problem is solved or until it is obvious the knife angle isn't the problem. Or the knife may be too warm. Suspend a chunk of dry ice over the block (ring stand).
  • Possible problem: Sections are deformed as they're sliced.
    Potential remedies: Slicing too fast, slow down the stroke. Or knife and/or block are too warm, cool with dry ice.
  • Possible problem: Sections curl up as they are sliced.
    Potential remedy: Check the angle of the knife. Or knife and/or block are too cold.

Mount the sections:

• Put a small drop of ddH2O on a polylysine-subbed slide. The drop should be large enough that water is visible around the periphery of the section.
• Place the section on top of the drop.
• Let the slides dry on a 45 degrees C slide warmer for 6 hr to overnight. The slide warmer should be in a dust-free area. As slides dry, wrinkles in sections should flatten out.

Pretreatment of slides for subsequent hybridization
Sections fall off slides easily during the pretreatment.
Handle the slides gently throughout the process.

• Dewax sections in xylene, 2 X 10'.
• Rehydrate through 100%, 95%, 80%, 60% and 30% EtOH; 1' in each solution.
• Incubate in 0.2N HCl, 20'.
• Rinse in ddH2O, 30".
• Incubate @ 70 degrees C in 2X SSC (preheated to 70 degrees C), 30'.
• Rinse in ddH2O.
• Pronase treat:
  -Remove the slides from the rack and place horizontally in a tray.
  -With a Pasteur pipet, carefully drip ‰0.25mg/ml pronase solution onto the sections and incubate 10' (thhe incubation time starts with treatment of the first slide).
  -Drain off pronase and replace slides in the rack.
• Stop the pronase reaction by inhibition with 2 mg/ml glycine in PBS, 30" then 1X PBS, 2X 30"
• Fix in 4% paraformaldehyde, 20'.
• Rinse in 1X PBS, 2X 4'.
• Acetylation (do under the hood):
  -Place racked slides in 500 ml 0.1M Triethanolamine, pH 8.0 (fresh; titrate with HCl).
  -While stirring rapidly, add 1.25 ml acetic anhydride drop by drop to ensure proper mixing. Incubate 10' during which time the stirring may be slowed down.
• Rinse in 1X PBS, 2X 3'.
• Dehydrate through 30%, 60%, 80%, 95% and 100%, EtOH; 2' in each solution.
• Air dry.

FROZEN SECTIONS
Preparation of embryos (>10 hr) and larvae for frozen sections

• Collect and dechorionate embryos as described for paraffin sections.
• Embed unfixed embryos or larvae immediately in O.C.T.
• Place a drop of O.C.T. directly on the chuck.
• Immerse the chuck in liquid nitrogen but do not let the nitrogen cover the top or touch the O.C.T.
• Remove the chuck when the O.C.T. starts to turn white but before it's completely frozen.
• Place a second, smaller, drop of O.C.T. on top of the unfrozen portion of the first.
• Quickly place the embryos in the center of the drop. With a paint brush, gently move the embryos back and forth until they are evenly distributed over the middle portion of the drop.
• Place the chuck back into liquid nitrogen until the second drop is completely frozen as before (again, do not let the nitrogen touch the O.C.T.).
• Put the chuck with embryos in the cryostat so they can come up to temperature.

Cutting frozen sections

• Take the chuck (plus embryos!) out of the cryostat (they should be at cryostat tempera- ture, -14 to -19 degrees C).
• Quickly trim the block into a rectangle with a razor blade.
• Put the trimmed block back into the cryostat. If the block falls off the chuck, glue it back on with some fresh O.C.T.
• Place the chuck into the microtome head.
• Slice. Sections are mounted onto slides as they are sliced. Pick up section(s) directly off the knife onto a gelatin subbed slide. You won't have to touch the sections, they will "jump" off the knife onto the slide. Picking up sections will take a bit of practice.
  - 6-8 micron sections are fairly easily managed
  - put 1-3 sections on each subbed slide
• Air dry the slides for at least 20'.
• Fix with:
  4% paraformaldehyde 20'
  3X PBS 5'
  1X PBS 5'
  1X PBS 5'
  30% EtOH 2'
  60% EtOH 2'
  80% EtOH 5'
  95% EtOH 2'
  100% EtOH 2'
• Sections may be stored at RT for at least a month.

  Troubleshooting
  Problems and their solutions are essentially the same as for paraffin sections. However, the knife should not be cleaned with xylenes. Changes in temperature should be achieved by regulating the cryostat temperature. In addition, if the guide-plate is in a bad position you may get torn or mangled sections, sections that curl up and don't come down the knife or sections which stick to the guide plate.

Pretreatment of slides for subsequent hybridization
Basically the same as for paraffin sections but dewaxing is not necessary.

• Incubate in 0.2N HCl, 20'.
• Rinse in dd H2O, 30".
• Incubate @ 70 degrees C in 2X SSC (preheated to 70 degrees C), 30'.
• Rinse in ddH2O.
• Pronase treat:
  -Remove the slides from the rack and place horizontally in a tray.
  -With a Pasteur pipet, carefully drip 0.25mg/ml pronase solution onto the sections.
  -The incubation time starts with treatment of the first slide.
  -Drain off pronase and replace slides in rack.
• Stop the pronase reaction by inhibition with 2 mg/ml glycine in PBS, 30", then 1X PBS, 2X 30".
• Fix in 4% paraformaldehyde, 20'.
• Rinse in 1X PBS, 2X 4'.
• Acetylate (do in fume hood):
  -Place racked slides in 500 ml 0.1M Triethanolamine, pH 8 (fresh; titrate with HCl)
  -While stirring rapidly, add 1.25 ml acetic anhydride drop by drop to ensure proper mixing. Incubate 10' during which time the stirring may be slowed down.
  -If gelatin slides are used, acetylation is unecessary and step 19 may be omitted.
• Rinse in 1X PBS, 2X 3'.
• Dehydrate through 30%, 60%, 80%, 95% and 100% EtOH; 2' in each solution.
• Air dry.

 

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