Recently, it has become possible to do non-radioactive in situs; (described elsewhere in this manual). These have several major advantages, the main one being the ability to localise the expression with single-cell resolution. However, there are cases where radioactive in situs ;might still be the method of choice: they may be a bit more sensitive, and tissues deep within the embryo seem a bit less accessible to digoxygenin-labelled probes than to 35S labelled probes. I would try a non-radioactive whole-mount in situ first, and if that was unsuccessful, resort to the old-fashioned way. In time, as the non-radioactive protocol is refined, this one may become obsolescent.
This protocol is very largely that of Phil Ingham.
Fixing the embryos:
1) Use as many embryos as it is practical to collect. Less than about 500-1000 is
probably much harder to work with than a gram or two.
2) Wash embryos from collection plate into a sieve with a fine enough mesh to retain the embryos. Rinse them with water to remove excess yeast.
3) Dechorionate in 50% household bleach for about 2 minutes.
4) Rinse the dechorionated embryos thoroughly with water containing 0.1% Triton X-100.
5) Transfer the embryos to a tube with a tightly fitting lid that contains a 1:1 mixture of 4% paraformaldehyde ;in PBS, and heptane. For about a gram of embryos I usually use a 50ml tube, with 20ml each of formaldehyde and heptane. For fewer embryos, scale down the sizes appropriately.
6) Shake the tube vigorously for 15 minutes. Then remove as much as possible of the aqueous (bottom) layer with a pipette, being careful not to remove the bulk of the embryos that will be at the interface.
7) Add a volume of methanol equal to the heptane layer. Shake the tube very hard for about 30 seconds, and then allow the two phases to separate. The devitellinised embryos will sink to the bottom of the tube, and the rest will remain at the interface with the empty vitelline membranes. The yield at this stage is variable - anything over 50-60% is fine; if there are fewer than that, try shaking harder and longer.
8) Remove as much liquid as possible and all the interface, leaving the embryos at the bottom of the tube.
9) The rest of the treatments are carried out in the same tube as the embryos are now in; each liquid is added to the tube, the embryos allowed to settle, and the liquid removed with an aspirator.
10) Rinse the embryos in methanol for 2 minutes.
11) Rehydrate them as follows:
2 minutes in 9:1 methanol:4%paraformaldehyde
2 minutes in 7:3 " "
2 minutes in 5:5 " "
2 minutes in 3:7 " "
12) Post-fix the embryos in 4% paraformaldehyde for 20 minutes.
13) Wash once in PBS.
14) Dehydrate in an ethanol series for 5 minutes each: 30%, 50%, 70%.
15) The embryos can be stored in 70% ethanol at -20° for several weeks.
Embedding the embryos:
1) Fully dehydrate the embryos as follows:
90% ethanol, 5 minutes, twice
95% ethanol, 5 minutes, twice
100% ethanol, 3 minutes, four times
2) Transfer the embryos to a glass test tube, and incubate them in xylenes, twice for ten minutes each.
3) Incubate in a 1:1 mixture of Paraplast Plus ;(Polysciences):Xylenes at 61° for ten minutes, twice. Keep the tubes in a hot block for the rest of the procedure.
4) Incubate in Paraplast Plus at 61° for ten minutes, twice.
5) Set up the molds on a hot plate set just hot enough to ensure that the wax does not set when a little is put in the bottom of the mold.
6) Quickly dispense some of the embryos in wax into the bottom of the mold. Use a pasteur pipette with the end broken off for this operation. I warm it briefly in a bunsen flame, bring it to the temperature of the wax sitting in the water bath, and then pipette the embryos. You want to put in enough wax/embryos to cover the base of the mold in a thin layer. The number of embryos should be enough to form a monolayer in the mold: fewer will mean that there are fewer embryos in each section; more will stop the embryos from settling horizontally in the wax.
7) Allow the embryos to settle for about one minute, and then try to make them move into the centre of the mold. I do this by blowing the surface of the wax with a pasteur pipette, and thus pushing the embryos away from the sides. This takes a little time to perfect - it does not really matter if the embryos are spread around the whole base.
8) Fill the mold with wax by slowly and gently, running more wax down the side; try not to disturb the embryos. The wax should fill the mold to about two thirds of its volume. Don't let the wax solidify before adding more, as this will form a weak region, and the block may break.
9) Carefully remove the mold from the hotplate and allow it to set undisturbed for about 15 minutes.
10) The blocks can now be stored at 4° for several months before cutting sections.
Subbing slides;:
1) The microscope slides are subbed in poly-lysine to make the the sections stick to them
well. The slides can be subbed in large batches, and stored in a dust-free
environment.
2) Wash the slides by soaking them in a large container of hot water with some detergent. Gently agitate them. Rinse thoroughly (e.g. running water for 30 minutes), and finally rinse in distilled water.
3) Load the slides into racks, and dry in an oven.
4) Soak each rack of slides in a solution of 50µg/ml poly-D-Lysine in 10mM Tris pH8 for about 10 minutes.
5) Air dry in a dust-free place; this can take 24 hours.
6) Store the slides in boxes at room temperature.
Cutting sections:
1) The exact procedure used will depend on the microtome that you use. The following
notes should provide a guideline.
2) Push the wax block out of the mold, and mount it onto a wooden microtome block, using molten Paraplast Plus to stick it.
3) Trim the block with a razor blade so that the face to be cut is a trapezoid shape.
4) Mount the block onto the microtome, and cut 6 micron sections. The number that you can fit onto each slide depends on the size of the sections and the size of the coverslip that you intend to use for the hybridisations. I use 22mm2 coverslips, and use about four sections per slide. I usually cut individual sections rather than ribbons, but both are OK.
5) Place a subbed slide onto a hot-plate set at 45°. Put a drop of clean water onto the slide, and carefully place the section onto the water drop. It is important to put the newly cut (shiny) side of the section downwards. The section will spread flat over the water, and will be lowered slowly on to the slide as the water evaporates. After you have put enough sections onto a given slide, leave it on the hot-plate, undisturbed.
6) Leave the slides on the hot-plate overnight (or in a 45° oven).
7) The sections can be stored at room temperature in a dry box for several weeks prior
to hybridisation.
Making the 35S riboprobe:
1) The template DNA should be linearised downstream of the insert. In order to remove
any contaminating RNAses, treat the digest with 200µg/ml proteinase K for 30
minutes at 37°, phenol extract once, and chloroform extract. Ethanol precipitate and
resuspend at 1µg/µl.
2) I use the Stratagene transcription kit;, and follow their protocol:
1µg DNA
1µl 10mM rATP
1µl 10mM rCTP
1µl 10mM rGTP
1µl RNAse Block
1µl 0.75M DTT
4µl 5X transcription buffer (Stratagene)
5µl 35S UTP 40µCi/µl, 800Ci/mmole (SP6/T7 grade)
1µl of a 1:5 dilution of T3 or T7 polymerase in 1X transcription buffer
Final volume = 20µl
3) Incubate at 30° for 1 hour.
4) Add 10X MS to 1X, and 1µl of DNAse (use RNAse-free DNAse). Incubate at 37° for 30 minutes.
5) TCA precipitate a tiny aliquot, to discover the percentage incorporation of label, which allows to you calculate the amount of probe transcribed.
6) Add tRNA (phenol extracted, ethanol precipitated, and resuspended in DEPC treated water). Since you are transcribing new RNA, the total amount made depends on the amount of the hot (limiting) nucleotide in the reaction, and the incorporation. To get the right probe and tRNA concentration for the hybridisation, use the following example as a guide. For 75% incorporation, using 200µCi of UTP in the reaction, add 500µg tRNA. For different amounts of transcripts, add an amount that will give the same relative amount (i.e. for 50% incorporation of the same amount of label, add 333µg).
7) Phenol extract once; back extract with an equal volume of 10mM DTT; chloroform extract once; add an equal volume of 4M ammonium acetate (DEPC treated), and 2.5 volumes of ethanol.
8) Chill, spin, rinse pellet, and resuspend in 50 µl of 10mM DTT.
9) Add 50µl 2X carbonate buffer, and incubate at 60° for 140 minutes.
10) Add equal volume of 0.2M sodium acetate, 1% acetic acid; add 2.5 volumes of ethanol, chill, spin, rinse pellet, and resuspend in 50% formamide. Aim for 5 x 105 cpm/µl in the formamide. (For the example above - 75% incorporation of 200µCi - use 96µl of formamide).
-The probe is now 5X concentrated. Store it at -20° till use.
Prehybridisation Treatment of the Slides:
1) Load slides into a rack.
2) The following treatments should be carried out in glass dishes: the dishes I use hold 400ml.
3) Dewax in xylenes, twice, for 10 minutes each.
4) Rehydrate in ethanol series: 100%, 95%, 80%, 60%, 30%, 2 minutes each.
5) Incubate in 0.2M HCl for 20 minutes at room temperature.
6) Rinse in H20, 5 minutes.
7) Incubate in 2X SSC, 30 minutes, 70°.
8) Rinse in H2O, 5 minutes.
9) Digest with 0.125mg/ml protease in P buffer at room temperature for 10 minutes.
10) Incubate in 0.2% glycine in PBS for 1 minute.
11) Rinse in PBS, twice, 1 minute each.
12) Post-fix in 4% paraformaldehyde in PBS, 20 minutes, room temperature.
13) Rinse once in PBS, 1 minute.
14) Acetylate the sections in 0.5% acetic anhydride in 0.1M triethanolamine, pH8.0 (I use 3ml in 600ml), 10 minutes, room temperature, in the hood, with vigorous stirring. Acetic anhydride is very unstable in water - add it to the triethanolamine at the same time as the slides.
15) Rinse in PBS, 2 minutes.
16) Dehydrate in ethanol series: 30%, 60%, 80%, 95%, 100%, two minutes each.
17) Air dry.
-The hybridisations should be set up soon after this pretreatment - the sections are probably not very stable at this stage.
Hybridisation:
1) The probe concentration during the hybridisation should be around 100,000cpm/µl.
2) Boil the probe (which is in 50% formamide, 5mg/ml tRNA) for 2 minutes (3µl for each slide to be hybridised).
3) Chill on ice and add 4 volumes of hybridisation buffer. Mix well, and spin for 30 seconds.
4) Place 15µl on to each slide, near the edge of the tissue sections.
5) Carefully lower a 22mm2 clean (but not necessarily siliconised) coverslip over the liquid in such a way as to make the probe cover the sections. Try not to allow too many bubbles.
6) Seal the edge of the coverslip with rubber cement; use plenty. I find it easiest to pour some into a 5ml syringe and use it from the syringe.
7) Hybridise overnight in a humid chamber at 50° (not neccessary, but not a bad idea).
Washing:
1) Carefully peel off the rubber cement, using a sharp pair of forceps; try not to dislodge
the coverslip while doing this.
2) Place the slide in a rack in a tank of wash buffer at 50°. Do this, and all subsequent washes in the hood - the wash buffer is very smelly. Suspend the rack so that there is space for the coverslips to slide off, which should happen within a few minutes. If needed, stir the wash buffer gently.
3) When the coverslips are off, move the rack into a fresh tank of wash buffer. Wash for about 4 hours at 50°, with hourly changes of buffer.
4) Wash the slides in 1X NTE at 37°, for 5 minutes.
5) Incubate in 20µg/ml RNAse A (the stock solution is 10mg/ml, boiled for 5 minutes, allowed to cool slowly, and stored at -20°) in NTE at 37° for 30 minutes. This step removes single stranded RNA, and reduces background.
6) Wash in NTE at 37° for one hour with 4 changes.
7) Dehydrate through ethanol series: 30%, 60%, 80%, 95%, 100%. Dilute the ethanol with 0.3M ammonium acetate instead of water.
8) Air dry.
Autoradiography:
1) I use Kodak NTB2 emulsion, diluted 1:1 with water, and aliquotted into 5ml
scintillation vials, which are stored in absolutely light tight conditions at 4°. This is
the right quantity to fill the dipping chamber that I use, which is made of perspex and
is just larger than a slide: any vessel that is deep enough to allow the slide to be
dipped so that the emulsion easily covers the sections is suitable, but adjust the
aliquot size accordingly.
2) Set up a water bath at 45° in the dark room and melt an aliquot of emulsion. I put the vial inside a film developing tank which has some water in it, and put the whole thing in the water bath. This is then light tight, and you can come and go safely. Alternatively a dark room with a properly sealed rotating door can be used. The emulsion takes about 15 minutes to melt.
3) While dipping the slides and drying them, they must be kept out of all light other than weak safelight of the appropriate kind for the emulsion used (Kodak no. 2 for NTB2). The way that you organise the dark room depends on your set up, just make sure that it is really light-tight.
4) Pour the melted emulsion into the dipping chamber, which should be sitting in the water bath, to keep it warm, and thus to stop the emulsion from solidifying. Dip each slide briefly into the dipping chamber and remove it without rubbing off the layer of liquid emulsion. Place the slide into a rack so that it stands vertically with the end that you held while dipping, downwards. This allows excess emulsion to drain away from the sections, and causes there to be a thin, uniform layer over them.
5) Allow the slides to dry for at least two hours before putting them in light tight boxes, which should have some drying gel (e.g.. Drierite) in them. Tape the boxes shut, and wrap them extensively in aluminium foil. Expose at 4°, away from any radioactivity.
6) It is convenient to have three or four slides for each probe used, and to put one of each type into each box. This way when you want to develop them, you need only open one box, and you need not re-wrap it.
7) Try developing the first set of slides after about 4 or 5 days - moderate to strong
signals should be exposed enough by then. Judge how much longer to leave the other
slides from the amount of signal seen at this time.
Developing the slides:
1) It is crucial that all the solutions used in the developing and staining of the slides are
at the same temperature, which should be below 20°. If they are not, the emulsion,
which is very delicate when wet, may come off the slides during their treatment,
thereby ruining the whole experiment. To avoid this I make up the solutions the day
before using them, and let them all equilibrate in an 18° room. When they are to be
used they can then be taken into the dark room, and it is OK if they all warm up
slowly together. The solutions are:
Kodak D-19 developer
2% acetic acid
1:3 dilution of Kodafix fixer
3 litres distilled water
10mM sodium phosphate buffer, pH6.8
2) Allow the box of slides to come to room temperature while still sealed (about 1 hour).
3) In a dark room with appropriate safelight, load the slides into a rack.
4) Immerse them in developer for 2 minutes.
5) Transfer them to 2% acetic acid for 30 seconds (this acts to stop the developer, and is probably not crucial).
6) Incubate the slides in fix for 5 minutes.
7) Rinse them in distilled water for 15 minutes with two changes. They are safe in the light from this point onwards.
8) The final process is to stain the slides with Giemsa;. The degree of staining wanted varies with the intensity of the signal: if the signal is strong, stain for about 15 or 20 minutes, but if it is weak, stain for only 30 seconds to 1 minute. I usually stain the first batch to be developed weakly, and then decide how to treat the next batch.
9) Make up a 5% Giemsa solution in 10mM NaPO4 immediately before staining the slides.
10) Incubate the slides in the Giemsa solution for the desired amount of time.
11) To remove the stain, gently pour water into the container until the film that forms at the surface of the Giemsa solution is poured away. This is important so that the film does not coat your slides.
12) Rinse the slides two or three times in water for about 30 seconds. While wet, they can be viewed at low power, and if they appear overstained, they can be rinsed more extensively: normally this is unnecessary.
13) Air dry the slides, and then mount them in DPX mounting medium under a coverslip.
Solutions:
P buffer:
50mM Tris 7.5, 5mM EDTA.
5X Transcription Buffer:
200mM TrisHCl, pH8, 40mM MgCl2, 10mM spermidine, 250mM NaCl.
10X MS:
100mM Tris pH7.5, 100mM MgCl2, 500mM NaCl.
2X Carbonate Buffer:
80mM NaHCO3, 120mM Na2CO3, which should be pH10.2.
Hybridisation Buffer:
The hybridisation mix can be made in a large volume, and stored at -20°. The recipe
for 1.25X buffer is:
50% formamide
12.5% dextran sulphate
0.375M NaCl
12.5mM Tris pH7.5
12.5mM sodium phosphate buffer pH6.8
6.25mM EDTA
1.25X Denhardt's
12.5mM DTT
Wash buffer:
1X salts, 50% formamide, 14mM mercaptoethanol.
10X salts:
3M NaCl, 0.1M Tris, 0.1M NaPO4, 50mM EDTA, pH6.8.
Quantities for 1 litre: 175.3g NaCl
14.04g Tris HCl
1.34g Tris base
6.78g NaH2PO4.H2O
7.1g Na2HPO4
100ml 0.5M EDTA
10X NTE:
5M NaCl, 100mM Tris pH7.5, 10mM EDTA.
References
-Cox et al., Dev Biol, 101, 485 - 502 (1984)
-Ingham et al., Nature 318, 439 - 445 (1985)