In situ hybridizations

Basic Blue in situ Hybridizations for Drosophila melanogaster embryos

Materials

› Formaldehyde-fixed embryos (dechorionated embryos fixed in 4% FA in 1:1 PBS:Heptane for 20 minutes, washed in 100% methanol and frozen)

› Glass Pasteur Pipettes (Short) and Bulbs 

› Nutator

› Cold Room

› 55° Water Bath (or 65°C Water Bath)

› Floating rack for tubes

› Permeabilization Mix:
› This is a complex solution that should be made well ahead of time.

› For a 100 ml stock solution
› 100 ul 100% Triton X-100
› 50 ul Igepal CA-630
› 50 mg Sodium Deoxycholate 

› 50 mg Saponin

› 200 mg BSA Fraction V 

› Water to 100 ml

› Phosphate Buffered Saline

› Paraformaldehyde (20%)

› PTx: PBS+0.1% Triton X-100. For one Liter, mix 100 ml 10x PBS, 1 ml Triton X-100 in a final volume of 1 Liter (+ ~900 ml ddH2O).

› Hybridization Buffer
› This is a complex solution that should be made well ahead of time.

› For 100 ml stock solution:
› 50 ml Deionized Formamide (Invitrogen / Thermo AM9342) 

› 25 ml 20x SSC (Invitrogen/Thermo AM9770)
› 2 ml 50x Denharts Solution (Invitrogen / Thermo 750018)
› 1 g Boehringer Blocking Solution (Sigma 11096176001)
› 200 ul Heparin 50 mg/ml stock (Sigma H-3393)

› 10 ml Yeast RNA (10 mg/ml) (Sigma 10109223001) › 100 ul Tween-20
› 0.1 g CHAPS (Sigma C3023-1G)
› Water to 100 ml

› Digoxigenin Labeled Riboprobe
 

› Western Blocking Reagent (Sigma 11921673001)
› This is a 10x solution that gets diluted in PTx.

› anti-Digoxygenin-AP FAb Fragments (Sigma 11093274910)

› PTw: PBS + 0.1% Tween-20.

› Alkaline Phosphatase Buffer:
› This should not be made up in advance, but at the time of use:

› For 10 ml:
› 1 ml Tris-HCL pH 9 (1 M)
› 200 ul NaCl (5 M)
› 500 ul MgCl2 (1 M)
› 100 ul Levamisozle (10 mM) -- 2 drops from the Vector Labs stock... › 10 ul Tween 20
› Water to 10 ml

› NBT (50 mg/ml)

› BCIP (50 mg/ml)

› 100% Ethanol

› Glycerol

Procedure

Permeabilization and Post-fixation

1) Gather formaldehyde fixed embryos from the -20° freezer.
2) Remove methanol solution and rinse embryos 3x in methanol just to ensure any heptane is gone.
3) Replace final methanol rinse with 1 ml Permeabilization Buffer. Incubate for 2 hours on a nutator at 4°C.

In preparation for the next step, prepare sufficient 4% Paraformaldehyde in PBS. You will be using 0.5 ml per tube (100 ul PFA + 400 ul PBS per tube). Make it right at the end of the 2 hour incubation.

4) Remove Permeabilization Buffer and post-fix embryos in 500 ul of 4% Paraformaldehyde in 1x PBS for 20 minutes on a nutator at room temperature.

5) Do 5 x 5-minute washes in PTx.

At this point, it is important that the embryos aren't all clumped up together. If this is the case, try briefly vortexing embryos (while they are in one of the PTx washes) to disperse clumps. You run the risk of tearing up some of the embryos, but if the sample is clumpy, then the following steps will not occur uniformly and will lead to confusing results. It is easier to break up clumps now than it is once you start PreHyb.

Pre Hybridization

Note: the standard protocol is to use a 55°C water bath for pre-hyb and hybridization. If you find that your in situs have lots of background, we have found that incubating at 65°C can substantially reduce background without affecting the specific signal.

6) Add 500 ul of a 1 : 1 mix of Hybridization Buffer : PTx, incubate for 10 minutes on a nutator at room temperature.

7) Remove HyB Buffer from previous step. Add 250 ul 100% Hybridization Buffer and incubate for 1 hour in a floating rack in the 55° water bath.

Hybridization

8) Prepare probe/hybridization mix: final 250 ul per tube, 1 ng/ul Digoxigenin labeled probe diluted in Hybridization buffer. Note that this mix can be re-used several times. If you have done this before, you might already have some probe solution in the freezer.

9) Remove and discard prehybridization solution, replace with the Probe/Hybridization mix, flick tube gently to ensure mixing (but be careful not to spray embryos all over the place). Incubate overnight at 55°C in a floating rack.

Post-Hybridization Washes

10) Remove and save the Probe/Hybridization mix (label well and store at -20°C).

11) Rinse embryos once quickly in 100 ul Hybridization Buffer.

12) Remove Hyb Buffer rinse, and replace with 200 ul fresh Hybridization buffer. Incubate for 1 hour in a floating rack at 55°C.

13) Remove Hyb Buffer wash, and replace with 200 ul of a 1 : 1 mixture of Hybridization Buffer : PTx. Incubate for 15 minutes on a nutator at room temperature.

14) Wash 5 x 5 minutes with PTx. 

Antibody Incubation

15) Block for 1 hour in 1 ml Blocking Buffer (100 ul Western Blocking Reagent + 900 ul PTx) on a nutator at room temperature.

16) Incubate in the Antibody Solution (Blocking Buffer as above + 1 : 1000 dilution of anti-Digoxigenin FAb fragments) for 2 hours on a nutator at room temperature (or overnight at 4°).

17) Wash 5 x 10 minutes in PTw. Detection with NBT/BCIP

18) Wash 3 x 5 minutes in freshly prepared AP Buffer.
19) Incubate in 1 ml AP Buffer + 7 ul NBT (50 mg/ml) + 3.5 ul BCIP (50 mg/ml).

This part requires attention. This is the step when the color reaction will take place. It can either be done in tubes for a fixed amount of time, or all the embryos can be pipetted out into a 12-well plate and monitored under a microscope. It is not totally necessary to do the color reaction in a plate, provided that the embryos can be observed in the tubes. When doing this for the first time, perhaps consider taking a few embryos out to a fresh tube to determine whether in-tube development is feasible.

Color Development will occur between 15 minutes and overnight. Be vigilant. Note how long the total color reaction took to complete. It will take practice to know how long is long enough to let the reaction proceed.

20) Rinse embryos several times in PTw.

21) To stop the reaction, step into ethanol:

A) Remove all but 500 ul of the last PTw rinse, add 500 ul 100% EtOH, incubate 5 minutes on the nutator at room temp. (50%)
B) Remove 500 ul, add 500 ul 100% EtOH, incubate 5 minutes on the nutator at room temp (~75%)
C) Remove 500 ul, add 500 ul 100% EtOH, incubate 5 minutes on the nutator at room temp (~88%)

C) Remove 500 ul, add 500 ul 100% EtOH, incubate for 1 hour at room temperature or overnight at 4°C, on a nutator. (~94%)

Rehydration and Stepping into Glycerol for Mounting

22) Step the embryos back into PTw:

A) Remove 500 ul, add 500 ul PTw, incubate 5 minutes on the nutator at room temp. (~88%)
B) Remove 500 ul, add 500 ul PTw, incubate 5 minutes on the nutator at room temp. (~75%)
C) Remove 500 ul, add 500 ul PTw, incubate 5 minutes on the nutator at room temp. (~50%)
D) Remove 500 ul, add 500 ul PTw, incubate 5 minutes on the nutator at room temp. (~25%)
E) Remove all solution and perform three quick rinses with PTw, ending with about 1 ml PTw in the vial.

23) Step the embryos into Glycerol:

A) Remove 500 ul of the PTw, add two drops of Glycerol (using a transfer pipette), incubate on the nutator for ~5 minutes at room temperature.
B) Add two more drops of Glycerol, incubate for ~5 minutes on a nutator at room temperature.
C-X) Repeat step "B" until the volume of solution in the tube reaches ~1 ml.

Y) Remove half of the solution (500 ul ~50% glycerol) and add 500 ul 100% Glycerol by transfer pipette, incubate on the nutator at room temperature for 10-20 minutes (until the solution is completely mixed). Note, you might have to invert the tube manually a few times to make sure the very bottom of the tube is fully incorporated...
Z) Remove half of the solution (500 ul ~75% glycerol) and add 500 ul 100% Glycerol by transfer pipette, incubate on the nutator at room temperature for 10-20 minutes (until the solution is completely mixed). Once this is done, the embryos are ready for imaging, or for long-term storage.

Mounting for Imaging

24) Build a "chamber" for mounting embryos by gluing two pairs of coverslips on either end of a standard microscope slide. You are building spacers here, in-between which a drop of embryos will be placed, and upon which a coverslip will be placed.

25) Place a small drop of embryos in ~88% glycerol in the middle of the space between the coverslip spacers. Check under a dissecting scope that there are sufficient embryos of the proper stage present in this sample. You want the drop to be small enough that it won't run off the sides of the slide when you put a coverslip on top of it. If there is too much glycerine, wick some off with a kimwipe or paper towel fragment. It can also be helpful to corral the "good" embryos to a central spot, and push any off-stage embryos to the side.

.  Drop a coverslip on top of the drop of glycerine. Don't glue this one down. You can 'push' the coverslip around in some cases to re-orient embryos while you image them.

.  Image embryos under suitable conditions (a properly set-up DIC scope). Take lots of pictures.