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).

• 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.

• 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.

• 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.

• 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.

• 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.

• 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.

• 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.

• Lyophilize 25 ml (=25 mCi) each of 3H-dATP, dCTP, dGTP and TTP in an Eppendorf tube that has been rinsed with EtOH. Minimize the drying time (¾ 1h in Speed Vac) by dividing into smaller aliquots, if necessary.
• To the tube of lyophilized 3H-dNTPs*, add:
  +2.5ml 10X NTB
  +0.5mg DNA
  +H2O to 24ml
  Resuspend by vortexing.
  * If not all nucleotides are tritiated, add cold nucleotides to 25mM
• Add 0.5-2.0 ml diluted DNase I (1:400 dilution of 1mg/ml stock). If the precise amount of DNase to use is not known for a specific DNA fragment or batch of the enzyme, set up several reactions (0.5, 1.0, 2.0 ml) in parallel.
• Add 10U Pol I.
• Incubate 45' @ room temperature.
• Stop the reaction by adding 25ml 50mM EDTA (pH 8), on ice.
• Determine the % incorporation by TCA precipitation:
  -Add 1ml reaction mixture to 100ml 0.5 mg/ml carrier DNA in a 7ml glass tube
  -Spot 10ml on a GF/C filter disk
  -To the remaining 90ml, add 5ml cold 10% TCA; incubate 10' on ice
  -Filter onto GF/C, wash with 10% TCA, wash with 95% EtOH and dry
  -20-50% incorporation is good and corresponds to a S.A. ‰ 1-2 x 108 dpm/ug
  -probe with incorporation ¾ 10% should not be used
• Unincorporated nucleotides may be removed on a Stratagene Push Column.
• Probe size may be checked on a 8%acrylamide/7M Urea sequencing gel. - Prerun 200V, 30' - Load: 105 cpm/sample to 10ml formamide, boil 1', chill and add 2.5ml 5X loading buffer (0.25% BPB, 0.35% XC, 15% Ficoll in 5X TBE) - End - labeled DNA fragments (e.g., pBR322/Hinf I) may be used as size markers - Run 200V, ‰ 4h (BPB about 2/3 down the gel) - Soak 20' in distilled water - Impregnate 15 - 30' in Fluorohance (RPI; or comprable enhancer) - Dry and autoradiograph - Optimal size 75-150 bp
• Preparation of probe for hybridization (use at 2-20ng/ml) - Precipitate 125mg carrier DNA per 0.5mg 3H-probe (dry ice, spin 10', 70% EtOH wash, air dry in vacuo 2') - Resuspend in 10ml ddH2O - Denature probe by boiling for 2', chill and quick spin - Add 300ml hybridization buffer and store on ice before use. Probe (in hyb) may be stored several months @ -20 degrees C. Before using, boil 2' and chill on ice.

• Plasmid templates should be RNase-free CsCl-banded DNA. Linearize DNA with an enzyme that cleaves downstream of the insert. Digest with 200mg/ml Proteinase K (30' @ 37 degrees C), phenol-Sevag extract (2X) and NaOAc/EtOH ppt. Resuspend in TE @ 1mg/ml. Check on gel for completeness of digestion.
• Transcription reaction (25ml):
  9.5ml DEP-H2O
  5.0ml 5X TB
  1.0ml linearized DNA template
  1.0ml 10mM rATP
  1.0ml 10mM rCTP
  1.0ml 10mM rGTP
  1.0ml 0.5mM rUTP
  1.0ml 0.75M DTT
  1.0ml RNase-Block (25U/ml)
  2.5ml 35S-rUTP (100mCi)
  1.0ml T3 or T7 polymerase (10U)
  Vortex, spin down and incubate 1-2h @ 30 degrees C. Incorporation should be 70-90%.
• Determine % incorporation by TCA precipitation: - Add 1ml reaction mixture to 100ml 0.5mg/ml carrier DNA
  - Spot 10ml on a GF/C filter
  - To the remaining 90ml, add 5ml cold 10% TCA and incubate 10' on ice
  - Filter onto GF/C paper, wash with 10% TCA, rinse with 95% EtOH and dry
  - Fraction incorporated = cpm (TCA)/cpm (spot) x 10 - Calculate the mass of probe synthesized:
  ng probe = fraction incorporated x input 35S (in Ci)/Specific activity of UTP (Ci/mmol) x 340 x106ng/mmol x 4
• Add 1ml RNase-free DNase (1U/ml) to the 24ml reaction mixture and incubate 30' @ 37 degrees C.
• Stop the reaction:
  +60ml 10mM EDTA, 10mM Tris-HCl (pH 7.4), 0.2% SDS
  +1.67ml 5M NaCl
  +1.0ml 1M DTT
  Incubate 10-15' on ice
• Hydrolyze the probe (ideal fragment length = 75-150 NTs) by adding +2ml 10N NaOH. Incubate 45' on ice.
• Neutralize the reaction: +20ml 2M HEPES (unpH'd).
• Phenol-Sevag extract.
• Remove unincorporated nucleotides over a Stratagene Push Column.
  - Equilibrate column with 70ml STE + 20mM DTT
  - Load sample (‰ 110ml), push through
  - Wash with 70ml STE+DTT
• Count 1ml.
• Check length of hydrolyzed and unhydrolyzed probe on a 6% acrylamide/7M Urea gel. End-labeled DNA fragments may be used as markers.
• Probes are stored as 5X concentrates. Precipitate with the appropriate amount of tRNA as determined below. The amount of probe used per slide is 0.3-1.0ng/ml/kb (where kb refers to the length of the unhydrolyzed probe). Resuspend the probe in 20mM DTT/50 % Formamide (1/5th volume of the final hyb solution) and store @ -70 degrees C. mg tRNA = (nmol label) (fraction incorporated) (350ng/nmol) (4) (0.3ng/ml)(kb)

HYBRIDIZATION AND WASHES

Hybridization

• Pretreat the slides.
• Mix probe with appropriate buffer.
• Place hyb mixture onto the slides: Lay slides horizontally on a black surface. Apply 10-15ml probe solution along- side one edge of the tissue. Using forceps, carefully lower an 18 x 18mm coverslip from one side, evenly distributing the hyb so as to cover the entire tissue section. Avoid air bubbles!
• To avoid evaporation, seal the edges of the coverslip with rubber cement using a broken off Pasteur pipet or syringe. Allow the cement to dry.
• Incubate slides in a humid chamber 18-24h at 37 degrees C (DNA probe) or 50 degrees C (Riboprobe).

• For every rack of slides, prepare 2L DNA Wash Buffer (for 5 washes).
• After hybridization, gently peel off the rubber cement seal with a pair of forceps. Stand the racked slides in the first wash for 5' @ room temperature (RT). Carefully pick up each slide (the coverslip should fall right off) and dip a couple of times in the wash.
• Immediatlely transfer the slide to a rack in the second wash and incubate 10-15' @ RT. Do NOT let the slide dry out!
• Transfer the slides in the third wash into a 37 degrees C water bath for 18-24h. During this time, change the wash buffer two additional times.
• Dehydrate the slides through EtOH (@ RT): 70% (2 x 5') and 95% (2 x 5').
• Air dry.

• For every rack of slides, prepare 2L each of WDTT and NTE wash buffers.
• Remove coverslips as described above for DNA probes but use WDTT wash buffer which has been preheated to 50 degrees C.
• Wash 4-6h in WDTT @ 50 degrees C.
• RNase treat: Rinse slides 2 x 5' in NTE @ 37 degrees C
  Incubate 30' in 20mg/ml RNase A in NTE @ 37 degrees C
  Rinse 2 x 5' in NTE @ 37 degrees C
  Wash 1h in NTE @ 37 degrees C
• Wash in WDTT 14-18h @ 50 degrees C. Change the wash buffer once during this time.
• Dehydrate the slides through EtOH: 70% (2 x 5') and 95% (2 x 5').
• Air dry.

• Prepare the emulsion by placing an aliquot in a 45 degrees C water bath until melted (no longer than 10-15'). Slowly pour the emulsion into a prewarmed dipper. Add an equal volume of 45C water and gently invert the dipper to mix the emulsion.
• Place a metal or glass plate onto a tray containing ice to promote quick solidification of the emulsion on the slide.
• Dip the slides individually into the emulsion (@ 45 degrees C). Wipe the back of the slide with tissue paper and drain excess emulsion by holding the slide vertically on a paper towel.
• Lay the slide on the precooled surface and allow the emulsion to set for ‰10'.
• Transfer the slides to a vertical position in a drying rack.
• Allow the slides to dry in a light-proof place for „ 2h. Put the slides into a lightproof box and expose, dessicated, @ 4 degrees C for the appropriate amount of time. I use a slide box containing a drying agent (Drierite) sealed with tape and wrapped in aluminum foil.

• Remove the slides from the refrigerator and allow them to come to RT.
• Set up three staining dishes containing developer, water and fixer. The emulsion is very fragile when wet. It is important that all the solutions be at the same temperature during the developing process. Precool all the solutions to 15 degrees C in a water bath or on ice. Place the solutions at RT for developing and washing (allowed to warm slowly at RT).
• Place the slides in a rack and immerse into 15 degrees C Kodak D-19 Developer for 4'.
• Transfer rack to a stop bath (15 degrees C water or 2% acetic acid) for 30''.
• Fix the slides in Kodak Fixer (15 degrees C) for 5'.
• The lights may now be turned on.
• Place the slides back in the water and let stand until they reach RT (at least 20 degrees C).
• Rinse 1-2 times in ddH2O.
• Stain the slides in Giemsa 1-3' (DNA probes) or 3-5' (Riboprobes). Giemsa displays a lot of batch variation, so test one slide before staining them all. Rinse by dipping into water and wipe stain off the backs of the slides (EtoH helps).
• Air dry.
• Mount the slides with 1-2 drops of Polymount or Immersion oil.

Acetic anhydride (SIGMA A6404)

Bleach (any commercial product)

Bovine Serum Albumin (SIGMA A7906, Fraction V)

Carrier DNA (SIGMA D1626, Type III from Salmon testes): Prepare a 10mg/ml stock in TE. Shear to an average size of 50-500 bp by autoclaving in 100ml aliquots for 20'. Carrier RNA (SIGMA R1753, E. Coli Strain W, Type XX): Prepare a 10mg/ml stock in TE. Phenol extract (2X) and NaOAc/EtOH ppt. to clean up.

100X Denhardts : 2% of each Bovine Serum Albumin (SIGMA A7906, Fraction V), Ficoll 400 (SIGMA F4375) and Polyvinylpyrrolidene (SIGMA PVP360). Store @ -20 degrees C.

50% (w/v) Dextran Sulfate (Parmacia 17-0340-02): Mix 110g dextran sulfate in 130 ml sterile water (takes a while to go into solution). Store @ 4 degrees C.

DMSO (SIGMA D5879)

DNA Hybridization Buffer
50% deionized formamide
0.6M NaCl
1X Denhardts
10mM Tris-HCl, pH 7.5
1mM EDTA
20% Dextran sulfate
2-20ng/ml probe

DNA Polymerase I (BRL, NEB, etc.)

DNA Wash Buffer - For 1L:
0.6M NaCl = 120ml 5M NaCl
20mM Tris-HCl, pH 7.5 = 20ml 1M Tris-HCl, pH 7.5
1mM EDTA = 2ml 0.5M EDTA
50% Formamide = 500ml Formamide

DNase I (Worthington/Calbiochem DPFF): Prepare a 1mg/ml stock in 10mM Tris-HCl, pH 7.5/10mM MgCl2. Freeze in 20ml aliquots and thaw only once. Each batch of DNase will probably need to be titrated along with the amount required for optimal radiolabeling of each specific DNA fragment.

1M DTT (SIGMA D0362, FW 154.2)

0.5M EDTA, pH 8 (FW 372.2)

50mM EGTA, pH 8 (FW 380.4)

Emulsion [Kodak NTB-2/Cat 165 4433(3H) or Kodak NTB/Cat 165 4425(35S)]: The emulsion is very sensitive to light and should only be handled in the dark under red safe light (Kodak Safe Light Filter No. 2/Cat 152 1525, 15V light bulb) conditions. The emulsion is gelatin-based and is a solid at room temperature. Dispense in 15ml aliquots since repeated freeaing/thawing leads to elevated background levels. I store the emulsion in 50ml polypropylene tubes which I subsequently use as the dipping chamber. Wrap the tube in aluminum foil and store at 4 degrees C (do NOT freeze) in a light tight container.

Ethanol

Formamide
Low grade (98%) for washes (e.g., Mallinkrodt 3797)
High grade for hybs (e.g., BRL 5515UA): Deionize my stirring with 5-15% (w/v)
mixed bed resin (e.g., BIORAD AG501-X8, 20-50 mesh) for 1h @ RT.

Giemsa (SIGMA GS-500): Use at a 1:20 dilution in 10mM NaHPO4, pH 6.8.

2mg/ml Glycine (SIGMA G7126) in 1X PBS

0.2N HCl

2M HEPES (SIGMA H3375, FW 238.3)