Double in situ hybridization
Nigel Pringle (
There are at least two chromogenic agents that can be used with good sensitivity (blue and magenta) plus four fluorescent dyes (Fluorescein, Rhodamine, Cyanine 3 and Cyanine 5). Fast Red is another chromogenic option but this seems to work well only for very abundant transcripts - unlike most of the probes we deal with. Below I have described the double in situ protocols as using either two chromogenic reagents or two fluorescent reagents; however, combinations of chromogenic and fluorescent agents work fine although it would be usual to develop the fluorescence second as the chromogenic reaction product will obscure the fluorescence if both probes co-localize. The methods described below are extremely sensitive and have worked well for most cDNAs we have tried, including some corresponding to rare mRNAs.
The magenta colour is alcohol soluble and can be removed from your sections at any stage of the process if required (important if showing doubles with co-localization).
You need two differentially labelled probes - usually one labelled with DIG and the other FITC (or biotin, which we have yet to try). The FITC labelled probe is considered to be the least stable and is usually recommended for the probe that is to be developed first. However, in our experience this does not seem to be crucial. The overriding factor seems to be how highly each transcript is expressed in your tissue of interest.
Occasionally the template is not visible (too little added) and sometimes you get double bands of probe, possibly caused by incomplete digest of cDNA, but the probes still seem to work fine. The example on the left is of a DIG labeled probe. If you have labeled with FITC then the unincorporated nucleotides are highly visible (as they fluoresce on the UV light box).
Use chemicals from Roche - all are lithium salts,
Final concentrations of labeling mixture:
Roche catalogue number
10mM ATP 1 140 965 7.1ml
10mM CTP 1 140 922 7.1ml
10mM GTP 1 140 957 7.1ml
6.5mM UTP 1 140 949 4.6ml
3.5mM DIG-11-UTP 1 209 256 25ml
3.5mM FITC-12-UTP 1 427 857 25ml
RNase-free water 20.5ml
final volume 71.4ml
There are various methods of tissue pre-treatment. We prefer to fix tissue in 4% paraformaldehyde in PBS. Usually overnight at 4oC but for small pieces of tissue (e.g. young chick embryos ~E4) we fix 2-4 hours at room temp. In both cases this is followed by cryoprotection in 20% sucrose in PBS, at least 24 hours at 4oC. N.B It is very important that this sucrose solution is RNase free; we add diethylpyrocarbonate (DEPC) (1ml per litre), shake vigorously until all the DEPC globules have disappeared then autoclave to degrade the DEPC (breaks down into CO2 and ethanol) and to sterilize the sucrose solution (turns yellow on autoclaving). After cryoprotection, the tissue is then removed from sucrose and placed on a paper towel to remove excess sucrose, before immersing in Tissue-Tec, usually in small foil boats made by wrapping aluminium foil around the bottom of a suitable tube (we use disposable spectrophotometer cuvettes). The foil boat is filled with Tissue-Tec and the tissue placed in a suitable orientation for subsequent sectioning. Make a note of orientation of specimen before freezing, either by cutting a small notch in the top rim of the boat, or marking with a felt tip pen (Tissue-Tec turns opaque-white when frozen). Rapidly freeze your tissue by placing foil boat directly on to dry ice. N.B. if using a silicon mould (such as sold for EM embedding) place some aluminium foil directly in contact with the surface of the Tissue-Tec and place dry ice directly in contact with the foil – to ensure rapid heat conduction.
An alternative method is to cut frozen sections of unfixed "fresh frozen" sections and to fix them after cutting sections. It is said that this is more sensitive, but we have little experience of this method so cannot comment. However, fixing in 4% paraformaldehyde after cutting sections will require a proteinase digestion (see below) to allow access of your probe to the RNA, so it is more work.
One could write a book about cutting good frozen sections. Careful preparation and set-up of the knife and cryostat is crucial - anti-roll bar parallel to top of knife, good sharp knife (check under microscope for scratches) etc. We re-sharpen knives ourselves with a soapstone and soapy water (holding the knife in a commercially-available “sharpening back”). Sharpen one side of the knife-edge more than the other so as to prevent creating excessive bevel on both sides.
The knife angle (angle of the knife relative to the cut face of the block) can make a significant difference to the quality of the sections. We use 12.5o (Bright cryostat) but the optimal angle varies from cryostat to cryostat.
Temperature is very important and the temperature you choose will depend on many things, in particular the tissue you are cutting. We routinely use -25oC for the chamber temperature and -18 to -20oC for specimen temperature (embryonic CNS tissue). In general, if you are getting fracture lines along your sections you are cutting too cold. If the front of your block looks wet you are too warm. Play around until you find the optimum conditions for your tissue. Also note that small adjustments to the height of the anti-roll bar can have a significant effect on the quality of the sections so play around with this first, before altering temperatures etc. When you change temperature give the specimen about 15 minutes to equilibrate to the new temperature before going back to cutting sections.
Sections can be cut as thin as 5mm, but for serial sections I usually cut around 15mm. The optimal thickness will depend on the nature of your tissue. Block faces can make a big difference to section cutting and there are many thoughts on design of block face. I prefer this shape \ / where the top and bottom of block are parallel to the knife-edge. Other shapes used are circles, squares and diamond shapes. If you have uneven edges on your block you will often struggle to cut decent sections.
After cutting sections and collecting them on silicon-coated glass slides, let them air dry at room temperature. If there are any bubbles trapped underneath the section (often happens on large sections) they can be "burst" with the point of a needle, but do this before the section dries out completely.
We do not perform any further treatments on our sections. Under our conditions we have found that proteinase K treatment of sections does not increase sensitivity. However, if the sections are post-fixed with 4% paraformaldehyde then proteinase treatment is essential (this also applies to whole mounts and cell lines).
Hybridization with digoxygenin-labelled probes
The two probes - one DIG labelled the other FITC labelled - are both diluted (usually 1/1000) immediately before use in hybridization buffer (see below), denatured at 75oC for 5 minutes and an appropriate volume (usually 150ml per slide) of diluted probe placed on each slide. The slides are coverslipped (with glass coverslips, either oven baked at 200oC or straight from the box) and hybridized overnight at 65oC.
Slides are placed inside a sealed container with Whatman filter paper soaked in 2x SSC plus 50% formamide. It is important to include formamide otherwise the formamide in the hybridization buffer evaporates during incubation.
· 1x "salts"
· 50% formamide
· O.1mg/ml yeast tRNA (phenol/chloroform extract the stock solution before storing)
· 10% (w/v) dextran sulphate
· 1x Denhardt's.
Make up a large volume (100 ml will do around 1000 slides) with highest quality reagents and water (DEPC-treated) and store in aliquots at -20o.
· 2M NaCl
· 50mM EDTA
· 100mM Tris-HCl pH 7.5
· 50mM NaH2PO4.2H2O
· 50mM Na2HPO4
After overnight hybridization, slides are incubated in MABT until the coverslips slide off (in glass Coplin jars or similar). Washed a further three times (usually five minutes each) in MABT before placing in wash buffer (1x SSC, 50% formamide, 0.1% Tween-20) at 65oC for 2 x 30 minutes (can be longer doesn’t seem to matter too much). The slides are then incubated 2 x 30 minutes in MABT (100mM maleic acid pH7.5, 150mM NaCl, 0.1% (v/v) Tween-20).
Post-hybridization “blocking” of sections
The slides are transferred to a humidified chamber and incubated in blocking solution (MABT containing 2% blocking reagent -Roche, catalogue number 1 096 176) and 10% heat-inactivated sheep serum) for 1 hour at room temperature without a coverslip.
Your choice here will be determined by your choice of chromogenic vs fluorescent probes (or a mixture) but will probably involve using Alkaline Phosphatase (AP) conjugated anti‑FITC Fab fragments from sheep (Roche, catalogue number 1 207 741) followed by AP conjugated anti-DIG Fab fragments from sheep (Roche, catalogue number 1 093 274).
Chromogenic probes In the example below we first develop the FITC-labelled probe to a blue colour, then develop the DIG-labelled probe to magenta. Remember to include appropriate controls such as omitting labelled probe from some slides, or omitting the anti-AP antibody from others. This controls for specificity and is required to show that killing the activity of the AP enzyme before adding the secondary antibodies was successful (see below).
Replace the blocking solution with AP-conjugated anti-FITC antibodies (Fab fragments; Roche catalogue number 1-426-338) diluted 1:1500 in blocking solution, and continue the incubation overnight at 4oC (or four hours at room temperature for strong signals).
Post-antibody washes and colour reaction.
1. The slides are transferred to Coplin jars and washed 3 x 5 minutes in MABT, then 2 x 10 minutes in pre-staining buffer (100mM Tris-HCl pH9, 100mM NaCl, 50mMgCl2).
2. The pre-staining buffer is replaced with staining buffer (100mM Tris-HCl pH9, 100mM NaCl, 50mM MgCl2, 5% (w/v) polyvinyl alcohol (av. Mw ~100k) from BHD or FLUKA (we have had problems with some batches from Sigma), 0.2mM 5-bromo-4-chloro-3-indolyl-phosphate (BCIP, Boehringer), 0.2mM nitroblue tetrazolium salt (NBT), both from Roche. These can be bought as liquids, 3 ml of NBT catalogue number 1-383 221 and BCIP catalogue number 1 383 213, although we routinely buy in BCIP (1 585 002) and NBT (1 585 029) as powder (for economy) and make up as specified below.
3. Incubate in the dark at 37oC until the signal reaches a satisfactory intensity (usually a few hours to overnight, although exceptionally they can be left over the weekend).
NBT stock is 100mg/ml in 70% dimethylformamide- dilute 50ml per 50 ml.
BCIP stock is 50 mg/ml in 100% dimethylformamide- dilute as above.
N.B Crucially important is the inclusion of PVA. This prevents the reaction product from diffusing from the reaction site and pushes sensitivity up by approx 5-10 fold. We would not consider developing the colour reaction without PVA present; it really makes a big difference. However it is difficult to dissolve. To make a 10% solution in water place in a water bath or oven at 80oC, keep shaking/ inverting the bottle for approx 24 hours until you have a very viscous, clear solution.
We usually make up a solution of 2x staining buffer, add the BCIP and NBT to this then add an equal volume of 10% PVA in H2O. Mix well before adding to slides and develop in the dark at 37oC for as long as required. Depending on probes this can be between 4 to 8 hours. Occasionally slides need to be left overnight.
When the blue colouration has developed to your satisfaction wash slides in water.
Note, this blue reaction product is stable in alcohol and xylene, but it does tend to fade over a period of several months.
If you are only performing single in situ’s then NBT/BCIP is the best colour reagent to use, as it provides the best sensitivity. After development of the colour reaction, dehydrate through an ascending alcohol series (30,50,70,90,100% EtOh) for 30 seconds each followed by 2 x 2 minutes in xylene. Slides can then be permanently mounted using by dropping some commercial xylene mountant such as Xam (from BDH) onto the slides and covering with a glass coverslip. However if you wish to perform double in situ hybridization do not permanently mount but proceed as follows.
Killing the first AP enzyme
There are two methods of killing AP - heating at 65oC or acid treatment. I use a combination of both to be really sure that the first AP is killed.
1. Heat at 65oC in MABT for 30 minutes (or longer)
2. Wash 2 x in room temp MABT
3. Incubate in 0.1 M glycine-HCl pH 2.2 for 30 minutes
4. Wash in MABT
Slides are now ready for the next antibody.
1. Incubate sections for 1 hour in blocking solution.
2. Incubate overnight at 4oC this time in AP-conjugated anti-DIG Fab fragments (Roche, catalogue number 1 093 274) diluted 1/1500 in blocking solution. Or 4 hours at room temp. In this case dilute anti-DIG Fab fragments 1/1000.
3. Wash extensively in MABT.
4. Wash in pre-staining buffer.
5. The pre-staining buffer is replaced with staining buffer (100mM Tris-HCl pH 9, 100mM NaCl, 50mM MgCl2, 5% (w/v) polyvinyl with the addition of INT/BCIP (Roche catalogue number 1-681-4600 for 3 mls) diluted 75ml in 10mls. I make my own buying in INT (p- Iodonitrotetrazolium violet) from Sigma (catalogue number I 8377) and BCIP p-toluidine salt from Sigma (catalogue number B 8503) and dissolving both of them at 33mg/ml in dimethyl sulphoxide.
6. Place at 37oC and allow the sections to develop until a dark magenta colour forms. This usually takes from a few hours up to 24 hours depending on the signal strength. If leaving the reaction product to develop overnight the stain often turns into a mass of crystals, which can be annoying for taking photographs at high magnification. Unfortunately I’ve not found a way round this problem, but changing the staining solution before leaving overnight seems to help in some, but not all, cases.
7. Wash slides before mounting with any non-alcohol/xylene mountant (e.g 50% glycerol) and sealing the coverslip with nail varnish.
NOTE: This magenta colour reaction is alcohol soluble so be careful. This is an advantage for doing double in situ’s where the signals might overlap and you might wish to photograph the first layer before removing it and developing the next (If this is what you want then do the INT/BCIP colour reaction first, remove stain by placing in ascending alcohol series (50,70,90,100% EtOH).
The example shown is of platelet derived growth factor alpha (PDGFRa: blue, arrows) and PAX6 (magenta) in a mouse E12.5 spinal cord section.
Fluorescent in situ hybridization using tyramide signal amplification (TSA).
The TSA plus fluorescent systems use horseradish peroxidase (HRP) to catalyze the deposition of a fluorophore labeled tyramine amplification reagent onto tissue sections or cells. The reaction is quick (less than ten minutes….) and is almost as sensitive as NBT/BCIP.
The kits are available from PerkinElmer Life Sciences and comprise amplification buffer plus small tubes containing the fluorophore of choice that has to be dissolved in dimethysulphoxide (DMSO) before being diluting 1/50 in the amplification buffer just before use.
Killing HRP enzyme
Four different TSA Plus kits are currently available from PerkinElmer Life Sciences INC currently (June 2001) they cost £193 for 50-150 slides.
Fluorescein catalogue number NEL741 (50-150 slides) NEL741B (250-750 slides).
Rhodamine catalogue number NEL742 (50-150 slides) NEL742B (250-750 slides).
Cyanine 3 catalogue number NEL744 (50-150 slides) NEL744B (250-750 slides).
Cyanine 5 catalogue number NEL745 (50-150 slides) NEL745B (250-750 slides).
For an example see Pringle et al (2002) Development 130, 93-102 (Pdgfra and Fgfr3). Note that we find the fluorescent probes give a rather “spotty” image that is less suitable for high-magnification (subcellular resolution) micrography.