sect13
For a good account of the theoretical background and scope of in situ hybridisation refer to the pink Course Manual for the University of Leicester In Situ Hybridisation Course, by Paul Senior et al. (September, 1989).
Decide which radiolabelled UTP you will use.
32P is cheap, available every week (HFI) and gives good signal rapidly but with only fair resolution at the tissue level. Losses and contamination can be easily monitored during the transcription/hybridisation process. A good nucleotide to start with.
35S and 33P give better resolution but take twice as long to give a result on autoradiography. 33P apparently gives lower background than 35S (Bev. Faulkner-Jones, WEHI). I have never tried 33P but I have found relatively high backgrounds with 35S for my particular probes.
The importance of positive controls.
In the early stages of optimising this technique, it is extremely useful to have tissue that will act as a positive control, i.e. a known site of target mRNA where you can rely on obtaining signal to check that the experiment has worked in a technical sense. I include positive controls with every experiment to gauge variation between experiments.
Refer to the Promega Protocols and Applications Guide for further information, pp 58 - 64.
N.B. All tips/reagents etc. should be prepared as for RNA work, except do not use DEPC-treated water in the transcription reaction. Trace amounts of DEPC apparently inhibit RNA polymerase activity.
A1. Template preparation
The DNA which will act as a template must be subcloned into a vector containing promoters for RNA polymerase, e.g. Bluescript has T3 and T7 promoters. Ideally, the insert DNA should be cloned into Bluescript in both orientations so you can use the same RNA polymerase to generate both the sense and anti-sense strands (and eliminate variability between the T7 and T3 RNA polymerases - one way to try to match sense against anti-sense).
The template must be linearised when undertaking in vitro transcription. First, determine the orientation of your template and which strand will be sense and which anti-sense. This diagram shows the multi-cloning site region of Bluescript II KS:
If a DNA insert designed to be the template is cloned into Bluescript as shown, then transcription with the T3 RNA polymerase will generate sense mRNA. However, to prevent transcription of Bluescript sequences the template needs to be linearised in this case with BamHI or XbaI. Similarly, if the template is linearised with HindIII and transcribed with T7 RNA polymerase then anti-sense mRNA will be generated - this will give the positive result on tissue section. It is important to avoid linearising the template with restriction enzymes which generate a 3' overhang. Always try to use an enzyme that leaves a 5' protruding end. If there is no alternative then you can use the 3'-->5' exonuclease activity of Klenow DNA polymerase to convert the 3' overhang to a blunt end (see page 58 of the Promega Protocols and Applications Guide).
To linearise the template:
1. Cut a generous amount (e.g. 15 ug) with the appropriate enzyme overnight in 100ul.
2. Make up to 400ul with sterile water.
3. Phenol/chloroform extract once.
4. Chloroform extract once.
5. Precipitate by adding 40ul of 3M NaAcetate pH 5.3 and 1ml of absolute ethanol. Spin for at least 15 minutes at 13,000g
6. Wash in 70% ethanol and spin for a further 5 minutes.
7. Carefully take off the supernatant and dry in the speedivac.
8. Resuspend in sterile water to give a concentration of 1mg/ml (assume losses of about 30%). This template can be stored at -20¼C for years.
9. Run 0.5ul of the template on an agarose gel to check that it is fully linearised. Run some un-cut plasmid in an adjacent lane as a control.
B. In vitro transcription reaction
Decide how much cold UTP to use in the reaction. The concentration of any nucleotide must be at least 1.5 mM (and ideally much higher than this). UTP will be the limiting nucleotide because only a certain concentration can be achieved when the nucleotide is radiolabelled. With 32P, if 10ul (100 mCi) of a 20ul reaction is hot UTP32, then the final concentration will be 1.67mM. I usually add an equal concentration of cold UTP (to give a final concntration of 3.33 mM) when transcribing a 750bp probe. Full-length transcripts are what is desirable and these are more likely to be produced at higher nucleotide concentrations. When transcribing from a longer template you could try 3:1 cold:hot UTP, i.e. add cold UTP to a final concentration of 5mM. The down side of adding cold UTP is that the specific activity of the resultant probe decreases.
Transcription reaction:
| For 1 probe: | ul | Final conc | |
| 5 x Transcription buffer (Promega) | 4 | ||
| 250 mM DTT | 0.5 | ||
| RNAsin | 0.5 | ||
| 10mM ATP | 0.8 | 333 mM | |
| 10mM CTP | 0.8 | 333 mM | |
| 10mM GTP | 0.8 | 333 mM | |
| 50 mM UTP (cold) | 0.67 | 1.67 mM | |
| hot UTP32 (100 mCi) | 10 | 1.67 mM | |
| linearised template DNA (1 mg/ml) | 1 | ||
| RNA polymerase | 1 | ||
| Total | 20.4 | ||
I usually make up a cocktail for the total number of probes I am transcribing then aliquot this and add the appropriately line arised template and RNA polymerase to each one.
[Promega 5 x Transcription buffer is:
| 200 mM | Tris-HCl, pH 7.5 |
| 30 mM | MgCl2 |
| 10 mM | spermidine |
| 50 mM | NaCl |
It is recommended not to chill this buffer on ice after adding the template DNA because the spermidine can precipitate the DNA.]. The in vitro transcription reaction is incubated at 37¼C for about 40 minutes, then a further 1ul of RNA polymerase is added and the reaction incubated for a further 35 minutes.
C. Post-transcription steps
The next step is to digest the DNA template with DNase.
To each tube, add:
2ul of tRNA (i.e. Boehringer E.Coli tRNA, molecular biology grade, 10mg/ml)
2ul of RNAsin
0.5ul of RNase-free DNase I and incubate at 37¼C for 10 minutes.
(N.B. if using 35S UTP it is important to add 1ul of 1M DTT at this stage and to ensure that all aqueous solutions that the probe is dissolved in from this point onwards contain DTT to a final concentration of 10 mM -- some people say 100 mM).
Add 75ul of sterile water to make the volume up to 100ul. At this point I remove 1ul and spot it onto DE81 paper for Cerenkov counting (to enable calculation of probe yield later) and I aliquot 1ul into an eppendorf tube for running on a polyacrylamide gel to assess the quality of transcription. This piece of paper is designated the "initial" count.
Next the probe needs to be phenol/chloroform extracted and precipitated:
1. Add 300ul of sterile water.
2. Add 400ul of phenol/chloroform and do extraction.
3. Discard the phenol/chloroform phase after checking that it is not hot (if very hot, e.g. >> 500 cps on mini-monitor, then your losses are too high).
4. Add 40ul 3M Na Acetate and 1 ml of absolute ethanol and put the tube on dry ice for > 10 minutes.
5. Spin at 13,000g for at least 15 minutes.
6. Pour supernatant into 50ul tube and check that your losses are not high.
7. Spin tube for further 2 minutes and remove remaining supernatant.
8. Resuspend in sterile water. The volume used depends on whether you are going to hydrolyse your probe (see next section). If so, then resupend in 100ul of water. If not then resuspend in 20ul of water and proceed to the section, "Calculation of riboprobe yields".
D. Hydrolysis of the probe
Hydrolysis of the probe is desirable if the probe length is greater than 300 to 400 bp. Alkaline hydrolysis is carried out to reduce the average probe length to about 150 to 200 bp to improve access of the probe to target mRNA in the tissue section (see page 45 of the Leicester manual). The formula for calculating hydrolysis time (T) is T = Lo -Lf / 0.11.Lo.Lf where Lo is the original length in kb and Lf is the desired length in kb.
To the 100ul of probe from the section above add 100ul of hydrolysis buffer and incubate at 60 ¼C for the appropriate time (for a 750 bp probe this is approximately 48 minutes). After that time has elapsed add 200ul of stop buffer.
The probe is then precipitated by adding:
1ul of tRNA
40ul of 3M NaAcetate
1 ml of absolute ethanol
and left on dry ice as above before spinning for 15 minutes at 13,000g.
The probe is resuspended in 20ul of sterile water.
Hydrolysis buffer:
| 80 mM | NaHCO3 |
| 120 mM | Na2CO3 |
| 20 mM | b-mercaptoethanol |
Stop buffer:
| 0.2 M | NaAc pH 6.0 |
| 1.0% | glacial acetic acid |
| 10 mM | DTT |
E. Calculation of riboprobe yields
I remove 1ul from my final 20ul reaction volume and add 4ul of sterile water to this. Of this 5ul, I spot 1ul onto a piece of DE81 paper and keep the rest for loading onto a polyacrylamide gel. This piece of paper will give the "final" count.
With 32P, I count my papers in the tritium channel of the scintillation counter.
The yield of each probe is calculated using the following formula:
yield (in ng) ="final" count divided by the "initial"
count
x amount of UTP in mCi
x MW of UTP
x 4.13 (a constant to allow for the differing weights of the other
nucleotides)
divided by the specific activity of the radiolabelled nucleotide
x the total of the ratio of cold to hot UTP added
i.e. for UTP32 in the above protocol using 1:1 cold:hot UTP:
yield (in ng) = "final" count/"initial" count x 100 x 500 x 4.13 x 1/3000 x 2
= "final" count/"initial" count x 137.7
Therefore the maximum theoretic yield is 137.7 ng of probe.
[N.B. To be strictly correct the MW of UMP, not UTP, should be used in the above calculation, i.e. 333 not 500. However, I have always used 500 and, because everything is relative, it really doesn't matter so long as you are consistent between experiments. The amount of probe that I use in the hybridisation step below is based on the yield determined using the above calculation with 500 as the MW].
F. Checking the transcription product on a polyacrylamide gel
I cast a 4% polyacrylamide gel and run the pre- and post-hydrolysis probes on it to assess the quality of the reaction product and to check that all has gone well with the hydrolysis step. The most important aspect is the amount of full-length transcript present.
I cast a narrow, long gel and use the shark's tooth combs.
For 50ml of 4% acrylamide solution (with running buffer of 1% TBE):
2.5ml of 20X TBE
5ml of 40% acrylamide
Make up to 50ml with 7 M urea solution.
Add 120ul of 25% ammonium persulphate and 45ul of TEMED and let set for 45 to 60 minutes.
To the 1ul aliquots of probe, add 4ul of sequencing stop buffer (containing xylene cyanol and bromophenol blue). Heat the tubes to 80¼C for 2 minutes and then load 2ul per well. I leave an empty well between samples.
Run the gel till the bromophenol blue is about to run off. The gel will be quite radioactive so take due care. It can be placed against XAR-5 film as is, sandwiched between glad wrap, or dried on the gel drier and then placed against film (the latter is preferable). An exposure of 1 hour is usually adequate.
The following protocol uses paraffin sections as opposed to frozen ones. In general, paraffin sections result in better morphology, allow the collection of serial sections and produce far better results for "watery" tissues like embryos. I have not observed any diminution in signal when comparing results obtained with paraffin as opposed to frozen sections.
A. Preparation of paraffin-embedded tissue and cutting of sections
The tissue is collected and placed immediately in 4% paraformaldehyde (PFA) dissolved in PBS (phosphate buffered saline). It is generally left immersed in the fixative overnight. The pieces of tissue should not be too large - about 5 mm x 5 mm x 5 mm is large enough. It is important for the fixative to gain access to any compartments or spaces within the tissue as easily as possible. For example, any embryos older than E14.5 should have their head and body cut apart with a razor blade and their peritonel cavity opened. The amniotic sac should be opened for any embryo older than E9 5. Larger pieces of tissue should be transected to allow ready exposure to fixative. This can be achieved with the least disruption when the tissue has already been immersed in 4% PFA for several hours - the tissue goes quite hard allowing easier dissection.
After 12 to 16 hours of fixation, the tissue is transferred to a 0.5M solution of sucrose dissolved in PBS. It is stored at 4¼C until processing through to paraffin. As a general rule this should be done as soon as is possible, although I have successfully used blocks of tissues stored at 4¼C for several weeks.
Processing through to paraffin consisits of taking the tissue through a series of graded alcohols to a clearing solution and then paraffin. It is performed overnight on an automatic processor located in the Histology section of the Department of Anatomy (phone 344-5752 for bookings, Helen Makin is in charge). Either short (1 hour per step) or long (2 hours per step) cycles are possible. Larger pieces of tissue require the longer infiltration time. After processing, I generally embed my own tissue into blocks of paraffin. In this way I can orientate the tissue according to how I want to section it. This is important because orientation can be crucial in navigating one's way through a tissue block on the microtome.
Tissue sections are cut on a microtome in the H.F.I. histology facility. I generally cut 5 to 10 mm sections. Thicker sections will improve signal per section but possibly at the expense of resolution. A dark field microscope on hand next to the microtome can be essential in checking one's progress through the block. Sections are mounted on glass slides coated in aminoalkylsilane, an agent which assists adhesion of the section to the slide (see below for protocol of how to "sub" slides). To mount each section, first delicately transfer the section (using two paint brushes) from the cutting stage of the micotome to float it in a container of distilled water mixed with a few mls of absolute ethanol. Then use a plain glass slide to transfer the section to a water bath maintained at about 50¼C. The traces of alcohol on the section cause a surface tension effect which immediately removes all wrinkles and after 10 seconds or so the section is ready to be manouevred into position onto a "subbed" slide and then dried on a heating rack overnight at about 45¼C. Once thoroughly dried, wax sections can be stored in a safe location at room temperature and used successfully for hybridisation histochemistry years later.
[Protocol for coating slides in aminoalkylsilane:
1. Place glass slides in racks and immerse them in a tub of hot water containing a small amount of "Decon" detergent overnight.
2. Rinse the racks in running water for 3 hours.
3. Sterilize the slides in an oven set at 180¼C overnight.
4. Allow slides to cool.
5. In a fume hood, coat slides in a freshly prepared 2% solution of 3-aminopropyltriethoxysilane (Sigma A3648) in dry acetone for about 5 seconds.
6. Rinse once in dry acetone for a few seconds.
7. Rinse twice in distilled water, 5 minutes each.
8. Remove from the fume hood and dry the slides overnight at 42¼C.
9. Store the slides at room temperature in a dust free environment.]
B. Pre-treatment of sections
Prior to hybridisation paraffin slides need to be pre-treated with a gentle protease digestion to improve access of the riboprobe to the target mRNA. The protease used is pronase E (Sigma P5147). Pronase E is autodigested and aliquotted prior to use. To do this, dissolve 40 mg/ml of pronase E in distilled water and allow it to autodigest at 37¼C for 4 hours to destroy contaminating nucleases. Aliquot the solution into sterile eppendorf tubes at 125ul/tube and dry it down in the speedivax. Store at -20¼C. When using coplin jars, these contain ioslides in 40ml of appropriate solution.
Pre-treatment protocol:
1. De-wax slides:
-Heat in 65¼C oven for 60 mins
-Leave slides in Histoclear for at least 60 mins
(Meanwhile make up 4% paraformaldehyde in phosphate buffer & set up 37¼C water bath)
2. Hydrate slides:
-1 minute each in 100% x2, 95%, 90%, 70%, 50% ethanols, then water
3. Rinse in P buffer (50 mM Tris-HCl, pH 7.5/5 mM EDTA, pH 8.0).
4. Digest in pronase E for 10 mins:
- Measure 40 mls of pre-warmed P buffer into Coplin jars. Use 1 ml of this P buffer to resuspend an aliquot of pronase E. The final concentration of pronase E in P buffer is 125ug/ml.
5. Rinse in phosphate buffer x2.
6. Post-fix in 4% paraformaldehyde for 10 mins.
7. Rinse in phosphate buffer x2.
8. Transfer slides to racks and dehydrate:
-Rinse in water, then 50%, 70% and 100% ethanol
9. Dry slides in dessicator ready for hybridization. This is a good time to mark the position of the section on the underside of each slide using a marker pen. Slides pre-treated in this way should be hybridised as soon as possible to avoid degradation of the tissue mRNA. If required, they can be stored for short periods.in ethanol vapour at 4¼C.
C. Hybridisation
First it is necessary to determine how many slides you will hybridise. This often depends on the yield of your probes. For practical purposes, 50 slides is the upper limit for one run. For the riboprobes that I have used, I find a probe concentration of 75 to 100 ng of probe per ml of hybridisation buffer is optimal. This appears to work for other people's probes as well.
The amount of hybridisation buffer used per section depends on the size of each section but I generally use an average of 50ul per section. Therefore if you have a yield of 75ng of probe you will have enough to hybridise 20 sections with that probe.
The basic recipe for 1 ml of hybridisation buffer is:
100ul of 10 x salts
500ul of deionized formamide
35ul of tRNA (20 mg/ml)
200ul of 50% dextran
165ul of Q water (including probe)
My approach is to calculate the total amount of hyb buffer required (for all probes, including "losses"), make this up excluding the water, aliquot this out and then add the appropriate amount of water (and probe) for each probe separately. This is done as follows:
1. Determine the number of sections to be hybridized and the volume of hybridization buffer to be used for each slide:
-e.g. 50ul of hyb buffer per section for 50 sections
-i.e. 2.5ml of hyb buffer + 20% losses = approx. 3ml required in total.
Losses are high because the dextran is very viscous and a lot of hyb buffer is left coating the eppendorf tubes and pipette tips.
2. Determine the concentration of each probe to be used:
-e.g. 75ng of probe per ml of hyb buffer
-i.e. 3.75ng of probe per section if using 50 ml of hyb buffer per section
3. Make up hyb buffer minus probe and water.
e.g. 3 ml of hyb buffer are required, thus we need 3 tubes of 1 ml each.
In each tube:
100ul of 10 x salts
500ul of deionized formamide
35ul of tRNA (20 mg/ml)
200ul of 50% dextran
[N.B.
a) preheat dextran to 80¼C to make it tractable. I pipette out 200ul of water in a second eppendorf tube and compare this against the amount of dextran to be aliquotted out. Use a syringe or a pipette tip with the end cut off-- it is very viscous! To make the 50% dextran solution initially,dissolve 50 g of dextran sulphate in sterile water at 80¼C, make this up to 100 ml and store in aliquots at -20¼C.
b) 10 x salts is: 3 M NaCl
100 mM Na2HPO4, pH 6.8
100 mM Tris-HCl, pH 7.5
50 mM EDTA, pH 8.0
0.2% BSA (Sigma A7638)
0.2% Ficoll 400 (Sigma F9378)
0.2% Polyvinyl pyrolidone (Sigma P6755)]
4. For each probe determine the:
a) number of sections
b) total volume of hyb buffer required
c) actual volume of hyb buffer minus water required
d) volume of probe+water to be added to c)
e) ng of probe required for that number of sections
f) volume of probe to be added (based on probe concentration, i.e. amount of probe required in ng divided by concentration of probe)
g) volume of water to be added
5. For each probe, aliquot out the required amount of hyb buffer minus water.
6. For each probe in turn, add the required amount of water and probe to the aliquotted hyb buffer, heat the mixture at 80¼C for 1-2 mins then vortex, pulse spin and apply to sections before cover-slipping.
7. Carefully place the slides with cover slips in position into a humidified chamber containing 50 ml of wash buffer (see below) and then allow them to incubate at 50¼C for 16 to 20 hours.
Steps 4 to 6 can be complicated so let's consider an example:
Assume we are hybridising 30 slides with 2 probes (15 slides each). For the anti-sense probe, the yield was, say, 65 ng and this is dissolved in 20ul of sterile water (as per section II.D above, after probe hydrolysis and precipitation).
From step 4, for the anti-sense probe:
a) = 15
b) = 15 x 50ul= 750ul
c) = 0.835 x 750ul = 626ul
d) = 750 ml - 626 ml = 124ul
e) 3.75 ng per section x 15 sections = 56.3 ng
f) With the anti-sense probe, we have 65 ng in 20ul, i.e. 3.25 ng/ml and we require 65 ng. Therefore we need to add 56.3 / 3.25 = 17.3ul of probe to the total of 124ul
g) = 124ul - 17.3ul = 106.7ul of water to be added.
Step 5: 626ul of hyb buffer (minus water and probe) is aliquotted out for the anti-sense probe.
Step 6: 106.7ul of water is added to this and then 17.3ul of anti-sense probe is added. The tube is heated at 80¼C, vortexed, then pulse spun in the microfuge before applying about 45ul to each section (to cover losses) and placing coverslips completely over the hyb buffer and section.
[Comment on temperature of hybridisation:
It is said that the hybridisation of riboprobes to mRNA in tissue sections cannot be adequately predicted by the Tm equations used for nucleic acids bound to solid supports. Thus the optimal temperature of hybridisation needs to be determined empirically. For my probes, and also for the PTHrP probe, the optimal temperature seems to be 50¼C. However, for an IGF II riboprobe used by Felix Beck's group, 60¼C appears to be optimum. The other aspect to this is that the temperature of the post-hybridisation washes appears to be far less significant in terms of the signal achieved, compared to the temperature of hybridisation].
D. Post-hybridisation washing and ribonuclease A treatment
I follow the protocol of post-hybridisation washes as detailed in the pink Leceister in situ manual. In brief, this is as follows:
1. Formamide washes
The aim of the formamide washing step is to remove all excess probe and particularly the viscous dextran sulphate from the sections. Three washes (with agitation) at 50 to 55¼C, each of 45 to 60 minutes duration, are performed in formamide washing buffer. Each wash uses 500 ml of buffer in a plastic tray which can fit 2 slide racks each containing 25 slides. One litre of formamide washing buffer consists of:
100 ml 10 x salts
400 ml water
500 ml formamide (not necessary to de-ionise this)
The cover-slips are first dislodged by sequentially immersing each slide in a beaker containing approximately 200 ml of this buffer. (This is the same buffer is as that placed on the bottom of the hybridisation chamber to provide humidification during hybridisation (see above). Do not use water or 2 x SSC for this purpose - it must be a solution containing 50% formamide and the appropriate concentration of salts).
2. Rinse in RNase buffer
I make up 3 litres of this buffer and use 500 ml for the subsequent RNase A digestion step. The remaining 2.5 l is used to thoroughly rinse the slides and their racks to remove all traces of formamide washing buffer. 3 litres contains:
87.6 g NaCl
30 ml 1 M Tris-HCl, pH 7.5
6 ml 0.5 M EDTA, pH 8.0
water to 3 litres
3. RNase A digestion
75 mg of RNase A (Sigma R5503) is dissolved in 1 ml of RNase buffer and boiled for 2 minutes before adding it to the 500 ml of pre-warmed buffer (final RNase concentration = 150ug/ml). The slides are incubated with agitation in this solution for 60 minutes at 37¼C. It is important that the slide racks and trays used for this and subsequent steps are reserved for this purpose only and designated as "RNase contaminated". If any RNase used in this part of the protocol find its way into the pre-hybridisation treatment steps (section II B. above) of a subsequent experiment, the result could be a complete loss of target mRNA from these sections and a hopelessly flawed experiment.
4. Wash in 2 x SSC
I perform this wash at 55 to 60¼C for 15 minutes with one change of 2 x SSC and a further 15 minute wash. After this, the slides are rinsed in sterile water and then dehydrated through 70% ethanol and two changes of 100% ethanol (1 minute each) before being dried in the dessicator. The slides are now ready for autoradiography.
E. Autoradiography
1. Autoradiography with X-ray film
For slides hybridised with probes labelled with 32P, 35S or 33P, Kodak XAR-5 film is appropriate for autoradiography. The slides are taped face up to a piece of blotting paper and the X-ray film laid over the top. A 16 hour period of exposure to the film is usually adequate for 32P-labelled probes and should show a good signal in positive control sections. The aim of this step is to assess the success of hybridisation and to gauge the anticipated period of autoradiography with liquid emulsion. A reasonable signal seen on X-ray film after 16 hours exposure with a 32P-labelled riboprobe suggests that a 5 to 10 day period of exposure to liquid emulsion is necessary.
2. Autoradiography with liquid emulsion
a) Dipping the slides
I use the Ilofrd K5 emulsion. To coat 50 slides, I weigh out 10 g of emulsion in the dark and dilute this with 10 ml of glycerol water (i.e. 1 ml of glycerol dissolved in 59 ml of water and autoclaved). The emulsion is placed in a light-proof box and melted at 45 to 50¼C for 1.5 to 2 hours with occasional swirling to mix the solution. Once melted, the emulsion is poured into a dipping chamber placed in a 45¼C waterbath.
The slides are dipped with a smooth, gentle in/out action and the emulsion wiped from the back surface of the slide. The slides are placed vertically in a light-proof can to dry. After about 6 hours, dessicant is added to the bottom of the can and the slides left for the appropriate period of time. It is important to include a number of test slides which can be developed at periodic intervals to judge the best time to develop the main batch.
b) Developing the liquid emulsion
I develop my slides by immersing them in filtered Kodak D19 developer at 20¼C for 2 minutes, rinsing in filtered water and then fixing them for 2 minutes in filtered Ilford Hypam fixer diluted 1:5. With the lights now on, the slides are rinsed in water, immersed in 4% buffered formaldehyde for 10 minutes to harden the emulsion, then rinsed again. I prefer to counterstain my sections with haematoxylin only (for 30 seconds) and then dehydrate the slides for 3 minutes each through 70%, 90% and absolute ethanol (three changes) to histoclear (2 changes) and finally xylene before mounting in DePeX. It is important that the emulsion-coated slides do not come into contact with any acid-containing solutions, as this will potentially lead to dissolution of the silver grains.
The slides are now ready to be examined under bright- and dark-field illumination.