Whole Mount Insitu
from Koops April 97

 

 


SYNTHESIS OF PROBE

Mix these reagents in the following order at room temp.
sterile DEPC distilled water 9.5ul
5x transcription buffer 4ul
0.2M DTT 1ul
nucleotide mix (pH8.0)(lOx DIG RNA labelling mix) 2ul
linearised plasmid (~1ug/ul.) 1ul
placental ribonuclease Inhibitor (40U/ul) 1.5ul
Sp6,T3 or T7 RNA pol. (20U/ul) 1ul
Total vol 20ul

2.Incubate at 37 0C for 1 hour (40 oC for Sp6), then add another 20U of RNA polymerase. (This depends on how much RNA you want)
3.Incubate for a further hour at 37 0C (or 40 0C).
4.Remove a 1ul, aliquot and run on a 1% agarose TAE gel to estimate the amount synthesised. An RNA band approximately 10 fold more intense than the plasmid band indicates that ~l0ug of probe has been synthesised.
5.Add 2ul of RNase free DNase1(10U/ul). Incubate at 37 0C for 15 minutes.
6Pack a Sephadex G50 column equilibrated with 0.1% SDS, 50mM Tris-Cl (pH7.5), 0.5mM EDTA (TES) by loading a 1ml syringe barrel and spinning at 1500rpm for 90sec.
Dilute probe reaction to100ul with TES and load onto packed G50 column. Collect fraction and load another 100ul to the column and spin again (do not pool the fractions at this stage as all of the probe may be in the first fraction).

 

7Add 1/10 volume NaOAc, 2 volumes 100% EtOH and incubate at -20 oC for 30 minutes.
8Spin in a refrigerated microfuge for 10 minutes, wash pellet with ice-cold 80% EtOH Airdry pellet.
9Redissolve pellet in DEPC H2O at ~0.1ug/ul and store at -20 0C. (You may be better to redisolve in formamide).


l0x transcription buffer. 4OOmM Tris.Cl (pH 8.25), 60mM MgCl2, 2OmM spemidine.
nucleotide mix: 10mM GTP, 10mM ATP, 10mM CTP, 6.5mM UTP, 3.5mM DIG-UTP.

PRETREATMENT OF EMBRYOS

 

1Dissect embryos out in ice-cold PBS. Try to remove as much of the extraembryonic membranes as is possible. Be sure to puncture the amnion and in embryos of 10dpc or older, puncturing the head with a syringe needle helps to avoid trapping of reagents in the lumen.
2Fix in 4% parafonnaldehyde (PPA) in PBS at 4 0C overnight.
3Wash twice with PBTX for 10 minutes each at 4 0C.
4Wash with 25%, 50%, 75% MeOH/PBTX, then twice with 100% MeOH for 20 minutes each. (Can store the embyros at 4 0C at this point, although some people believe this increases background)
5Rehydrate by taking the embryos back through the MeOH/PBTX series in reverse.
6Wash three times with PBTX for 10 minutes each.
7Treat with 10ug/ul Proteinase K in PBTX for 5-10 minutes at room temperature. This treatment should be sufficient for mouse embryos up to 9.5dpc however for older embryos, add another 5 minutes to this incubation time for each frrther day of development. This is intended as a rough guide, and a titration experiment may be necessary for each batch of Proteinase K as well as for each stage of development.
8Wash twice with PBTX for 5 minutes each. Be careful! The embryos are fragile!
9Refix with 0.2% glutaraldehyde/4% PFA in PBTX for 20 minutes.
10Wash twice with PBTX for 10 minutes.
11Addition of prehybridisation mix, allow the embryos to sink, then transfer to a 2ml eppendoph tube.
12Incubate at 55 oC to 65 oC overnight. The embryos can be stored in this solution at -20 oC.
13Add hybridisation mix including ~0.2-1.0 ug/ml DIG labelled RNA probe.
14Incubate at 55 oC (or 65 oC if there are background problems) overnight.


PBTX: PBS with 0.1% Triton X-100
prehybridisation mix: 50% formamide, 5xSSC, 2% Boehnnger blocking powder (cat. no. 1096176, dissolve directly into the mix), 0.1% Triton X-100, 0.5% CHAPS (Sigma C-3023), 1mg/ml yeast RNA (sigma R-6625), 5mM EDTA, 50ug/ml heparin.
For hybridisation add probe to 0.2-1 ug/ml.

Steps 1-10 are carried out in 100 tubes. Enough liquid must be used to ensure that all embryos are completely covered during each step. All steps are carried out with sufficient rocking to agitate (but not destroy) the embryos, and unless otherwise stated, at room temperature. 4% PFA must be prepared ahead of time by dissolving the powder at 65 oC, and then cooled, A stock of 25% glutaraldehyde is stored at -20 oC and an aliquot is thawed just prior to use.

POST-HYBRDISATON WASHES

 

1Wash with the following for 5 minutes each at 55 oC (or 65 oC)

100% Solution 1
75% Solution 1:25%, 2XSSC
50% Solution 1:50%, 2XSSC
25% Solution 1:75%, 2XSSC
(During these washes, start preabsorbing the antibody as described below).
2 Wash with 2xSSC, 0.1% CHAPS twice for 30 minutes at 55 oC (or 65 oC).
3Wash with OixSSC, 0.1% CRAPS twice for 30 minutes at 55 oC(or 65 oC).
4 Wash with TBTX, twice for 10 minutes at room temperature.
5Preblock the embryos with 10% sheep serum, 2% BSA in TBTX for 2-3 hours at room temperarure.
6Remove the 10% sheep serum, 2% ESA from the embryos and replace with the preabsorbed antibody (See below). Rock overnight at 4 oC.

ALL washes are with about 1.ml in a 2ml eppendoff tube, and with rocking. For the washes at 55 oC or 65 oC,it is convenient to use a heater block placed on its side on a rocking platform.

Solution 1: 50% formamide, 5xSSC, 0.1% Triton X-100, 0.5% CHAPS
TBTX:50mM Tris.Cl pH 7.5), 150mM NaCl, 0.1% Triton X-100


PREABSORPTION OF ANTIBODY



The antibody must be preabsorbed with embryo powder (the longer the better). and despite the manufacturers recommendation is quite stable after dilution, and actually can be used up to four times following dilution.

1During the washing of the embryos (step 1 above), weigh out 3mg of embryo powder into a microtube, add 0.5ml of 10% sheep serum, 2% BSA in TBTX and 1ul anti-DIG antibody (Boehringer 1093274).
2Rock gendy at 4 oC for 3 hours or longer.
3Spin in a microfuge for 10 minutes at 4 oC
4Dilute the supernatant to 2ml using 10% sheep serum. 2% BSA in TBTX
5 Storeat4 oC until use.

Mouse embryo powder is prepared as follows: Homogenise -12.5-14.5dpc mouse embryos in a minimum volume of PBS. Add 4 volumes of ice cold acetone, mix and incubate on ice for 30 minutes. Spin at 10K x g for 10 minutes and remove supernatant. Wash the pellet with ice-cold acetone and spin again. Spread the pellet out and grind into a fine powder on a sheet of filterpaper and allow it to airdry. Store in an air-tight tube at 4 oC.

POST-ANTIBODY WASHES AND HISTOCHEMISTRY

 

1Wash at least five times with TBTX containing 0.1% BSA for 1 hour (the antibody solution can be kept at 4 oC and recycled up to 4 times)
2Wash overnight at 4 oC with TBTX containing 0.1% BSA.
3Wash twice with TBTX for 30 minutes.
4Wash three times with NTMT for 10 minutes.
5Incubate with NTMT includin g 6.6ul NBT and 3.3ul BCIP (X-phosphate) per ml. Rock for the first 20 minutes then transfer the embryos to a glass embryo dish (avoid using a plastic petri dish as crystals tend to form). Keep in the dark as much as possible and be careful not to let the colour reaction proceed too far as background can increase dramatically in a short time. If its time to go home and the colour reaction hasn't proceeded to the desired extent, Its best to wash in NTMT, then TBTX and store the embryos in TBTX at 4 oC overnight. The colour reaction can be started again in the morning.
6When the colour has proceeded to the desired extent, wash with NTMT then with PBTX.
7Wash several times in PBS with 1% Triton X-100. This can be done for several days and will decease any background.
5Fix the stain by incubating the ee'hryos in 4% PFA in PBTX overnight at 4 oC.
9If the embryos are to be stored for extended peroids or transported, take them trough a PBTX/glycerol series into 100% glycerol.


NTMT: l00mM NaCl, l00mM Tris.Cl (pH 9.5), 50mM MgCl2, 0.1% Tween-20. This must be made fresh on the day of use as pH decreases during storage due to the absorption of CO2.
NBT: 75mg/ml in dimethylformimide (store at -20 oC)
BCIP: 50 mg/ml. in dimethylformimide (store at -20 oC)
It is best to let these reagents warm to room temperature before use as it may decrease the amount of crystal formation in the colour reaction.

EMBEDDING AND SECTIONING OF DIG WHOLE MOUNTS

 

1if the embryos have been stored in glycerol, take the embryos back through a PBTX/glycerol series and wash several times with PBTX.
2Transfer to glass scintiflation vials (tetrahydmnapthalene dissolves plastic!)
3Replace with MeOH for 5 minutes, then with isopropanol for 10 minutes
4Replace with tetrahydronaptalene (Aldrich) for 15 minutes, and then with fresh tetrahydronapihalene. This must be carried out in a fume hood!
5Add an equal volume of molten paraffin wax at 60 oC and incubate for 20 minutes with occasional shaking.
6Replace with three washes of molten paraffin wax, each for 20 minutes at 60 oC.
7Transfer to an embryo dish (at 60 oC), transfer to room temperature, orientate using a heated needle and let the wax set. To remove the wax pellet from the dish, place the dish at -20 oC until it is possible to push the wax pellet ouc by applying pressure to one cornen using a straight blade, cut a block around the embryo and then affix this wax block to a wood or plastic chuck, for use in the microtome, using mounting wax.
8Cut sections using a microtome ( 14um sections), mount on subbed slides arid dry at 37 oC overnight.



SUBBING OF SLIDES

 

1 Dip slides in 10% HCl/ 70% EtOH.
2 Rinse using distilled water.
3 Dip in95% EtOH.
4Dry slides at 150 oC for 5 minutes and allow to cool. NB. The first 4 steps are not required if using precleaned slides.
5Dip slides in 2% TESPA (3-arninopropyltriethoxysilane) in acetone for 10 seconds.
6 Wash twice in acetone, then with distilled water.
7 Dry overnight at 37 oC.

COUNERSTAINING AND MOUNTING


1 Dewax slides in histoclear for 5 minutes with occasional shaking.
2 Wash with histoclear for a few seconds.
3Take the slides quickly through the following EtOH Series (a few seconds in each) 100% EtOH 100% EtOH, 100% EtOH, 95% EtOH, 70% EtOH, distilled water.
4Dip the slides into 0.5% eosin in 25% EtOH for 15 seconds. Eosin acts as a progressive stain so it's best not to leave the slides in eosin for more than 15 seconds.
5Take the slides through the following EtOH series: 70% EtOH, 95% EtOH, 100% EtOH, 100% EtOH, 100% EtOH. These washes remove excess eosin, only do each wash for a few seconds asthe stain will be stripped off.
7Wash twice with histoclear, do not dry. (The slides can be left in histoclear for several hours if convenient)
8Clean a coverslip using a lint-free tissue and then apply DPX using a glass rod.
9Gently lower the wet slide down onto the coverslip. Be careful so as to not entrap any air bubbles. Turn the slide over and let the DPX 'settle' for about 6 hours at room temperature.
10 Dry the mounted slides in a 37 oC incubator over two days.

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