Updated In Situ Protocol

    Probe Preparation:
  1. Linearize 10 µg of DNA at the 5' end in a standard 200 µl digest. (The enzyme should generate either a 5' overhang or a blunt end, not a 3' overhang.)
  2. Run 5 µl of the digest (0.25 µg) on a gel to make sure that it is linearized completely.
  3. Meanwhile, PIC extract remaining digest with 200 µl. Spin 5'.
  4. EtOH precipitate 190 µl of the aqueous phase by adding 20 µl 3M NaOAc pH 5.2 and 440 µl EtOH. Freeze on dry ice for 5', spin 15', then wash pellet with 70% EtOH. Spin 5' and air dry pellet ~10' at RT until all liquid has evaporated. Suspend pellet in 9.5 µl RNase-free H2O. The linearized template is now at 1.0 µg/µl.
  5. If gel results are good (i.e. fully linearized template), set up transcription reaction:
    6. 13 µl RNase-free H2O
    2 µl 10X transcription buffer
    2 µl DIG-NTP mix
    1 µl linearized DNA
    1.5 µl RNA polymerase
    0.5 µl RNasin
    20 µ Total Volume

    Incubate at 37°C for 2 hours. Make sure you have a gel poured to analyze the RNA.

  6. Add 30 µl H2O and purify through RNA G-50 spin column.
  7. Run 5 µl on gel. The RNA (although usually diffuse) should be at least as strong as the vector band.
  8. Add 95 µl Hyb solution, and store probe at -20°C. Typically use 20-40 µl probe per ml of hyb solution on your slides.

Day One (Slide Processing and Hyb):

  1. NaOH treat glassware by putting everything into an autoclave tub, adding some (~50 ml) 10 N NaOH, then filling tub with dI water, being sure to cover all glassware. Don't forget to also treat the slide rack and forceps. After glassware has soaked in NaOH for more than 1 hour, rinse ~10x with tap water, then 5 times with dI water. Drain on clean diapers until ready to use.
  2. Prepare solutions as follows:

    3. 0.2M Phosphate Buffer (PB), 2.2 L - prepare fresh in 4 L graduated cylinder
    48.1 g Na2HPO4 anhydrous (141.96 MW) - 154 mM final
    15.95 g NaH2PO4·2H2O (155.99 MW) - 46 mM final
    Fresh MilliQ H2O to 2.2 L

    4% paraformaldehyde, 400 ml - prepare fresh
    200 ml H2O
    Heat to ~70°C with stir bar
    Add 16 g paraformaldehyde and 1 drop of 10 N NaOH
    As soon as para dissolves (should be immediate), place on ice
    When RT, add 200 ml 0.2 M PB
    Sterile filter and equilibrate to RT

    PBS, 4 L - prepare fresh
    To remaining 2 L of 0.2 M PB, add 35 g NaCl
    Fresh MilliQ to 4 L

    Proteinase K Buffer, 400 ml - prepare fresh
    5 ml 0.5 M EDTA pH 8.0
    20 ml 1 M Tris pH 7.5
    Fresh MilliQ to 400 ml
    28 µl Proteinase K (14.4 mg/ml) --> 1 µg/ml final (add just before use)

    Acetylation Buffer, 300 ml - prepare fresh
    295 ml fresh MilliQ
    4 ml triethanolamine (Fluka 90279)
    0.525 ml concentrated HCl
    Mix well.
    *You will add acetic anhydride after you put the slides in. See below.

    Hyb Solution, 1 L prepare in advance - aliquot into 50 ml tubes and store at -20°C
    500 ml 100% formamide (Roche 1814320) (50% final)
    250 ml 20X SSC (5X final)
    100 ml 50X Denhardts (5X final)
    0.25 g bakers yeast RNA (Sigma R6750) (250 µg/ml final)
    0.5 g herring sperm DNA (500 µg/ml final)
    150 ml sterile MilliQ H2O

    Slide Processing:

  3. Fix and mount embryos as usual. (2 h fix at 4°C in 4% para/PB or PBS, rinse several times then overnight in PBS at 4°C, 2 h at 4°C in 30% sucrose/PB, mount in OCT.)
  4. Cut ~18 µm sections onto slides from a fresh pack and air dry until you are finished a set. Label with a pencil- Sharpie pen washes off during in situ processing. Cut serial sections such that the number (i.e. 1, 2, ...) refers to the set number and the letter (i.e. a, b, ...) refers to the probe. Place slides into clean slide box when they are dry. You can process 30 slides maximum for in situs. Slides can be stored at -20°C indefinitely or used as soon as they are sufficiently dry.
  5. Place slides in NaOH treated slide rack.
  6. Submerge in 4% para/PB for 10' RT. (Don't dump the para- you will need it again!)
  7. Wash 3 times with PBS for 3' each. (*Don't be too generous with the PBS or you'll run out by the end.)
  8. Digest in proteinase K 5' RT. (Digest younger tissue for less time ~3', older tissue for more time ~10'.) Generally, 5' works fine.
  9. Fix again in the original 4% para/PB for 5'.
  10. Wash 3 times with new PBS for 3' each.
  11. Acetylation: Pour 300 ml acetylation buffer into glass dish. Place slides in dish. They will not quite be submerged. Now dribble 0.75 ml acetic anhydride back and forth over the tops of the slides. Then mix by dunking the slide rack several times. Acetylate 10'.
  12. Wash 3 times with new PBS for 3' each.

    13. Prehyb and Hyb: (Preheat water bath to 80°C, and hyb oven to 68°C!):

  13. Place slides onto slide tray and add ~ 1ml hyb. buffer to each slide. Be sure tissue sections are completely covered. Incubate 0.5 - 2 h at RT. Squirt some PBS in the bottom of the slide tray so slides don't dry out (not really an issue here).
  14. Assemble and denature necessary amount of probe:
    Use 100 µl probe per slide containing ~ 20-40 µl probe/ml hyb solution. For example, for 3 slides with the same probe at 40 µl/ml, combine 330 µl hyb solution + 13.2 µl probe in an eppendorf tube (this includes an extra 10% to account for pipetting losses).
    Denature at 80°C for 5', then place immediately on ice until ready to use.
  15. Prepare white hyb. boxes by placing a slide ~ 3 slots from the bottom, and stuffing 2 Kimwipes saturated in PBS beneath the slide. Prepare a different box for each probe you will use.
  16. One slide at a time, pour off prehyb solution and dab away excess on a paper towel. Pipette 100 mµl probe in several drops across slide, then rotate slide to help mix probe with residual prehyb. (This seems to be important for even hybridization which always seems to be a problem.) Coverslip with 60 mm coverslips from fresh box, and place in the prepared slide box which should remain vertical. Continue with the next slide. Slides from different probes should go in different slide boxes to be safe. When you are finished all slides from one probe, tightly tape around the slide box to prevent drying out and place the box in suitable tupperware container. Place the tupperware into preheated hyb oven (68°C) on top of microfuge racks (so it's not touching the metal bottom of the oven).
  17. Continue with the slides from the next probe, until all are in the hyb oven. Incubate overnight at 68°C.

Day Two: (Washing and Antibody Incubation):

  1. Prepare the following solutions:

    2. 5X SSC, 400 ml - prepare fresh, preheat to 65°C
    100 ml 20X SSC
    400 ml MilliQ

    0.2X SSC, 2L - prepare fresh, preheat 1 L to 65°C
    20 ml 20X SSC
    To 2 L with MilliQ

    B1, 1L (Prepare ahead, sterile filter and store at RT indefinitely)
    100 ml 1M Tris pH 7.5 (100 mM final)
    30 ml 5M NaCl (0.15 M final)
    870 ml H2O
  2. Equilibrate small glass jar with the 5X SSC and large glass jar with 1 L 0.2X SSC to 65°C in the large water bath.
  3. Remove slides from the hybridization boxes, and use gloved hands to place them in small slide rack. Try to move quickly.
  4. When all slides are transferred, place rack in the 5X SSC bath. Incubate ~10' until the coverslips are visibly sliding off.
  5. Using clean forceps, transfer slides individually into the 0.2X SSC bath (65°C) with a large slide rack.
  6. Wash at 65°C for 1-3 hours. Usually at least 2 hours.
  7. Transfer slide rack to fresh 0.2X SSC at RT for 5'.
  8. Put slides into slide tray, and add 2 ml B1 to each slide.
  9. Block with 2 ml B1 + 10% HINGS for 1 h at RT.
  10. Pour off B1 into the slide tray and place 0.5 ml antibody solution (B1 + 1% HINGS + 1:5000 anti-DIG Ab) on each slide. Incubate overnight in the cold room.

Day Three (Washing and Color Reaction):

  1. Wash slides 3 times 5' with 2 ml B1.
  2. Equilibrate with 2 ml B3 5':
    3. B3, 1L - prepare ahead, sterile filter and store at RT indefinitely
    0.1 M Tris pH 9.5
    0.1 M NaCl
    50 mM MgCl2
  3. Place two pieces of clean parafilm flat in the bottom of a slide tray (without pipettes!). Roll 5 kimwipes per tray and prewet with PBS. Place these along the perimeter of the tray and one down the middle between the two parafilm pieces.
  4. Pipette 100 µl B4 in a straight line across the parafilm. Pour off B3, dab slide on towels, then invert carefully into the B4 solution. When all slides are finished, place trays in a drawer (or cover with foil) and incubate at RT for 6 h to 3 days. Observe them occasionally under the dissecting scope to determine when to stop the reaction:
    5. B4, (prepare fresh)
    First make a 24 mg/ml solution of levamisole in H2O.
    Dilute in B3 (you will need 100 ml per slide):
    3.375 µl NBT/ml (from 100 mg/ml stock)
    3.5 µl BCIP/ml (from 50 mg/ml stock)
    0.24 mg/ml levamisole (1:100 from 24 mg/ml stock)
  5. Stop the reaction by transferring slides to a tray and washing once with TE. Slides can be stored in TE until all are finished. (Overnight is okay.)
  6. Slides can be mounted at this point, or counterstained with antibody.

    Antibody Counterstaining (optional):

  7. Equilibrate slides in PBS for 1 h.
  8. Block with 1 ml per slide PBS, 10% HINGS, and 0.1% TX-100. 1 h RT.
  9. Add 0.5 ml primary antibody in PBS, 1% HINGS, 0.1% TX-100 overnight at RT.
  10. Wash 3 times 10' with PBS.
  11. Add 0.5 ml biotinylated secondary antibody diluted 1:200 in PBS, 1% HINGS, 0.1% TX-100 1-3 h at RT.
  12. Wash 3 timesb10' with PBS.
  13. Prepare ABC complex (Vectastain Kit)
    1 ml PBS
    20 µl A
    20 µl B
    stand 30' on nutator at RT
    Add 10 µl 10% TX-100

  14. Add 0.5 ml ABC complex and incubate 1 h RT.
  15. Wash 3 times with PBS.
  16. DAB reaction
    5 ml H2O
    1 DAB tablet from Sigma
    1 H2O2 tablet
    Vortex.
    Syringe filter
    Place 0.5 ml of this solution on a slide (wear gloves always- this stuff is nasty!)
  17. Watch reaction under dissecting scope until the appropriate color has been reached.
  18. Stop the reaction by washing in PBS.
  19. Continue with remaining slides.
  20. Wash slides for 1 h in PBS before mounting.
  21. DAB solution should be inactivated with bleach before dumping.

    22. Mounting:

  22. Place slides in small rack and dehydrate for 3-5' each in 30%, 50%, 70%, 95%, 100% x 2 EtOH series.
  23. Incubate 2 x 20' in xylene.
  24. One at a time, remove slide and dab on paper towel to remove most, but not all of the xylene. Place 1 good drop of Permount on the right side of the slide and coverslip. Allow to dry overnight before moving slides.

Back