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Published in: Stern, C.D. (1998). Curr. Top. Dev. Biol. 36, 223-243.

Detection of multiple gene products simultaneously by in situ hybridization and immunohistochemistry in whole mounts of avian embryos.

Claudio D. Stern

Department of Genetics and Development, College of Physicians and Surgeons of Columbia University, 701 West 168th Street - #1602, New York, NY 10032, U.S.A.

TABLE OF CONTENTS

Introduction

Synthesis of digoxigenin- and fluorescein-labelled riboprobes by in vitro transcription

Preparation of embryos

Pre-hybridization, hybridization and post-hybridization washes

Antibody incubations

Detection

Detection of Alkaline Phosphatase with NBT/BCIP (blue precipitate)

Detection of Alkaline Phosphatase with Fast Red (red precipitate and fluorescence)

Detection of Alkaline Phosphatase with ELF⁷ (yellow-green fluorescence)

Detection of Alkaline Phosphatase with other chromogens (magenta, pink, green and brick-red precipitates)

Detection of Horseradish Peroxidase with Diaminobenzidine-H2O2 (brown precipitate)

Combining the colours

Time-table for multiple colour detection

Photography

Histological sectioning of whole-mounts

Designing appropriate controls

Trouble-shooting

Embryos disintegrated or folded

Background problems

Unincorporated hapten in the riboprobe

"Trapping" within cavities

Cross-reactivity with other mRNAs

Non-specific binding of antibodies

Endogenous peroxidase or phosphatase activity

No signal or low sensitivity

Persistent alkaline phosphatase activity from the first antibody

Acknowledgements

References

Introduction

Recent research on the avian embryo has now finally succeeded in successfully combining experimental embryology, for which avian embryos are unrivalled, with the availability of many antibodies and cDNA probes, allowing for the first time the objective identification of tissues and cell types and greatly increasing the level of sophistication possible in the design of experiments. At the same time, methods for interfering with the levels of expression of specific genes, including the use of retroviral vectors, have been made available for avian embryos, which expand the range of experiments that can be performed, targeted at understanding developmental mechanisms at the cellular and molecular levels.

In these experiments, it is often desirable to study the distribution of different mRNAs and proteins at a given stage of development within one embryo. For example, it may be of interest to detect the ectopic expression of a gene introduced with a retroviral vector, at the same time as studying the expression of one or more of its putative target genes. For this, a technique allowing the simultaneous visualization, with single-cell resolution, of two or more different gene products is essential. Good methods have become available only recently; this chapter reviews our present state of knowledge, with detailed protocols and instructions for designing appropriate controls and tips for trouble-shooting different problems. It should be borne in mind, however, that most of these techniques are still under development, as improvements are made continuously.

Synthesis of digoxigenin- and fluorescein-labelled riboprobes by in vitro transcription

When conducting in situ hybridization with multiple probes, it is particularly important to remove any unincorporated digoxigenin and fluorescein (especially in the case of the latter, which is "sticky"), which appear to be the main cause of high backgrounds. There are several ways to do this. The most efficient methods include: (a) the use of a Sephadex G25 spin-column made in an Eppendorf tube and (b) repeating the LiCl/ethanol precipitation step. The following protocol includes the latter method. All solutions and equipment for this part of the protocol should be RNAase-free, and gloves should be worn. Consult Sambrook et al. (1989) for instructions on how to do this. When making up solutions that will contain enzymes (e.g. RNA polymerases, DNase) it is best to use commercial RNase-free water rather than to treat water with diethylpyrocarbonate (DEPC) oneself, because traces of DEPC can inhibit enzyme reactions. If DEPC-treated water is to be used, it helps to autoclave the bottle after treatment for at least 1 hour, with the cap loosely fitted.

First, linearize 3-10μg of the vector (the most common vector is pBlueScript [Stratagene], which contains promoters for RNA polymerases T3 and T7 at each end of the polylinker) with the appropriate enzyme (Fig. 1A) for 4-5 hours or overnight (after cutting, check the efficiency of cutting in an agarose gel using 1/20th. of cutting reaction). Then add an equal volume of Phenol (equilibrated to pH 7.2-8.0) : Chloroform (1:1), vortex hard, and spin in a microfuge for 1-2 min. Take the upper layer into a clean Eppendorf and add 10μl 3M sodium acetate for every 100μl and 2.5x its volume of absolute alcohol. Vortex briefly and put at -20EC for 2 hours to overnight.

Spin at 4EC (15,000 rpm) for 15 min, take off alcohol with a sterile Pasteur pipette pulled to a fine tip. If the DNA is clean, the pellet will probably be invisible but it is important not to touch it with the pipette. Wash the pellet with 150μl 70% ethanol, vortex, and spin again 5 min. Take off the alcohol carefully and dry the pellet at 37EC. Add clean (preferably commercial Ultrapure) RNAase-free water to give a final concentration of about 1mg/ml. Allow to dissolve at 37EC for at least 15 min, with occasional vortexing or flicking the tube.

Transcribe with appropriate enzyme (T3, T7 or SP6; see Fig. 1) at 37EC (40EC in the case of SP6) for 2 hours. Proportions of different reagents are as follows (add in the order shown in Table 1):

After transcription, digest the template with RNase-free DNase (1-5 units per μg DNA): first make up a stock of 1:10 DNase in transcription buffer with 10% DTT (e.g.: 6 μl water, 1μl DTT, 2 μl transcription buffer, 1μl DNase I), then add this to the tube. It is important that the concentration of glycerol (contained in the polymerase, RNAsin and DNase solutions) should not exceed 10% of the total volume of either the transcription or DNase digestion mix, or activity of the enzymes will be impaired. Incubate for 30 minutes at 37EC. At this point, take an aliquot (e.g. 1/20th of sample) and run an agarose gel to check the probe. The yield of these transcription reactions can be as much as 10x the number of microgrammes of DNA template used.

Bring the volume up to 82μl with water, and add 8μl EDTA to stop the DNAse. Add 10μl 4M LiCl and 250μl absolute EtOH. Vortex briefly and put at -20EC as above. Precipitate 2 hours to overnight. Spin at 4EC 15 min. Take off alcohol (this time the pellet should be clearly visible, particularly when labelling with FITC-nucleotide). Wash with 300 μl 70% EtOH. Vortex very well. The pellet should be dislodged and should break up into many little pieces. Spin again 5 min. Take off alcohol. Rinse carefully with 30μl absolute EtOH and allow to dry at 37EC. Dissolve the pellet in water at 1mg/ml approx. (transcription should yield about 8-10x the original weight of the DNA). Leave at 37EC for at least 15 min, vortexing occasionally.

When making FITC-labelled riboprobes, repeating the LiCl precipitation helps to remove unincorporated FITC. Bring the volume of riboprobe solution up to 90 μl, add 10μl LiCl and 250μl EtOH and repeat the precipitation step above. After washing the pellet and dissolving again at 0.1-1mg/ml in water, check the probe in an agarose gel. In the case of fluorescein labelled probes, any unincorporated fluorescein will be visible in the transilluminator at the bottom of the gel as a green spot even in unstained gels (Fig. 1B). If some of this is still present, repeat the LiCl/EtOH precipitation once more or pass through a Sephadex G25 spin column.

Spin briefly, then add $10x its volume of hybridization buffer (see below) for storage (or make up to about 100-500 ng/ml in hybridization buffer). Store at -20EC.

For in situ hybridization with two probes, one probe is transcribed exactly as above, and the second probe is transcribed by the same method except that FITC-labelled UTP (Boehringer-Mannheim) mixed with unlabelled nucleotides replace the DIG-nucleotide mix (the proportion of FITC- to unlabelled nucleotides is described in the sheet supplied by the manufacturers).

Probes synthesized in this way and dissolved in hybridization buffer are stable at -20EC for many months. The heparin and RNA and formamide in the hybridization buffer act as very powerful RNAase inhibitors and degradation of the probe is virtually completely eliminated. However, fluorescein-labelled probes are probably a little less stable than digoxigenin-labelled ones. In our experience, however, even fluorescein-labelled probes can be stored in this way for at least 9 months and re-used at least 8 times (which also contributes to reduce background).

Preparation of embryos

When changing solutions for all the procedures involving embryos described below, particularly those involving embryos of 2 days and younger, the integrity of the embryos will critically depend on the care used in pipetting the solutions. Always use a Pasteur pipette, not a Gilson pipetman. Tilt the vial and turn it around to remove solutions with the pipette, avoiding sucking the embryos into it. At each step, remove all of the solution from the vial until embryos stay in position attached to the wall, but then add new fluid quickly so they don't dry. When adding new liquid, turn the vial so that the fluid runs down the opposite wall of the vial to that containing the embryos.

Collect embryos in calcium-magnesium-free PBS (CMF). Clean off any adhering yolk, and remove from the vitelline membrane. It is highly desirable to fix the embryos in a flat, stretched out state. Young embryos up to 1 day's incubation can be laid flat on the lid of a plastic dish, in a small drop of CMF. Just before fixing, carefully remove the CMF and start fixing by adding fixative directly onto the embryo gently using a Pasteur pipette. Older embryos (2-5 days' incubation) are best pinned out on a silicon rubber- or wax-coated dish using fine insect pins. Once arranged appropriately, the CMF can be sucked out and replaced with fixative. After a few minutes, the pins can be removed and the embryo transferred to a small (5ml or 20ml, depending on the number of embryos) glass scintillation vial. The fixative is freshly made 4% formaldehyde/CMF/EGTA (4% w/v paraformaldehyde powder added to CMF preheated at 65EC, stirring continuously. Adjust pH to about 7.5 with 5N NaOH. Allow to cool, then add EGTA to final concentration of 2mM). Leave in this 1 hour to overnight at 4EC. Transfer embryos to absolute methanol, and store in this for up to 1 week at -20EC.

On the first day of the in situ procedure, start by rehydrating the embryos through 75%, 50% and 25% methanol in PTW (PTW=CMF with 0.1% Tween-20), allowing embryos to settle between changes. Wash twice with PTW, 10 min each. For embryos older than about 2 days, bleach for 1 hour in 6% H2O2 (1ml H2O2 + 4ml PTW from 30% stock). Wash 3x with PTW, 10 min each. For the last wash, measure the volume of PTW (use 2 or 5 ml, depending on size of tube). Add Proteinase K (1:1000; final concentration 10μg/ml). Incubate at room temp. for 30 min regardless of stage of embryos, but reduce this to 15 min for very young (less than stage 4-5) or embryos that have been cultured by the New (1955) method, which become very thin. During incubation, gently roll the tube every few minutes to make sure the sides and top of vial get wet with Proteinase K (this will remove RNAase from the tube and cap).

Take off Proteinase K and rinse briefly and very carefully with a very small volume of PTW. Replace PTW with 4% formaldehyde in PTW (made as above but doesn't need to be fresh), containing 0.1% glutaraldehyde. Postfix 20 minutes.

Pre-hybridization, hybridization and post-hybridization washes

The following protocol is a modification of a method devised by Drs. David Ish-Horowicz, Domingos Henrique and Phil Ingham (unpublished). It is based on hybridization conducted under stringent conditions at low pH and low salt concentration, at a high temperature. This simplifies the post-hybridization washes, where the same solution and conditions are used. RNase treatment is not only unnecessary, but actually leads to loss of signal with most probes.

Remove postfixing solution and wash twice briefly with PTW. Remove PTW, and replace with 1 ml hybridization solution (Table 2).

Table 2 here.

Remove hybridization mix, and replace with another 1-2 ml (5 ml if using large vials) of the same solution. Place tube upright in a beaker in a water bath at 68-71EC. Incubate 2-6 hours. Remove hybridization mix, and replace with probe in hybridization mix (see above "Synthesis of labelled riboprobes"). For probes #400 nucleotides, use a lower hybridization temperature (e.g. 62EC for 150-250 nt, 65EC for 250-400 nt.). It is advantageous to prehybridize at the higher temperature, then to lower the temperature when adding the short probe. For in situ hybridization with two or more probes, add both probes simultaneously (provided that the probes are labelled differently: e.g. one with digoxigenin-UTP, one with fluorescein-UTP and the third with biotin-UTP). Allow to hybridize overnight in the water bath.

The next morning, remove the probe (keep at -20E for re-use up to 6-10 times). Rinse carefully three times with a small volume (1ml) of prewarmed hybridization solution. Wash twice with 1.5 ml (4 ml if using a large vial) of prewarmed hybridization solution, 30-45 min in water bath. Wash 20 min with prewarmed 1:1 hybridization solution : TBST (1:10 from the stock in Table 3).

Table 3 here

Rinse three times with TBST at room temperature. Wash three times 30-60 min with TBST. Rinses can be made with a very small volume. Washes are best done by filling the vial to the top, replacing the cap and then laying the vial horizontally on a tilting table.

Antibody incubations

After TBST washes, embryos are first blocked with a protein solution to lower non-specific binding of the antibody and then placed in antibody solution overnight. For whole mount staining to detect two or more labels, each antibody incubation is followed by detection of the appropriate enzyme. Then the enzyme activity is inactivated by one of several possible methods (described in more detail later) before proceeding with incubation in the next antibody. This allows the same enzyme (e.g. alkaline phosphatase) to be detected by a different chromogenic reaction.

Remove the embryos from TBST and place them in 1 ml of blocking solution: 5% heat inactivated (at 55EC for 30 min) sheep serum in TBST with 1 mg/ml BSA for 3 hours at room temperature. During this time, preabsorb the antibody (Table 4).

Table 4 here

Dilute the preabsorbed antibody (Table 4) into the blocking buffer with the embryos so that final concentration of antibody on the embryos is 1:5,000. Incubate overnight at 4EC on a rocking platform.

The next morning, remove the antibody from the embryos and keep it for re-use (up to 8-10 times; this also helps to reduce background). Rinse three times with TBST (Table 4). Wash 3x 1 hour with TBST, rocking (fill vial right up to the top and place horizontally on rocker). Embryos older than 2 days should be washed more extensively (5x 1 hour, and one overnight wash in TBST).

Detection

As mentioned above, each antibody is detected sequentially, which allows several antibodies coupled with the same enzyme (alkaline phosphatase) to be detected in different colours. Procedures for inactivating the enzyme are given later. The paragraphs below describe the chromogenic reactions.

Detection of Alkaline Phosphatase with NBT/BCIP (blue precipitate)

Remove TBST and wash 2x 10 min with NTMT (Table 5). Incubate in NTMT containing 4.5 μl NBT (75mg/ml in 70% dimethylformamide) and 3.5 μl BCIP (50mg/ml in 100% dimethylformamide) per 1.5 ml, rocking, protected from light, at room temperature. The reaction may take anything between 30 minutes and several days at room temperature. It develops faster at 37EC, but this can speed up the reaction too much and greatly increase background. If colour hasn't quite developed after about 3-4 hours, it is advisable to leave the vials at 4EC room overnight to slow down the process and to take them out to room temperature again the next day if necessary. If colour has still not developed after the whole of the next day at room temperature, there is no need to leave them at 4EC again the next night. After colour has developed as desired, stop by washing 3x 10 min in PBS. The reduction in pH from these washes will intensify the blue colour.

Table 5 here.

Detection of Alkaline Phosphatase with Fast Red (red precipitate and fluorescence)

Fast Red TR produces a red precipitate which also fluoresces intensely when viewed with rhodamine optics (green excitation, red emission). Fluorescent detection appears to give higher sensitivity and if this is to be used, less intense red staining is needed. There are several commercial sources of Fast Red, which have different properties. Vector labs supply a kit called Vector Red which contains three solutions. Two drops of each solution are added to 5ml of Tris-HCl buffer at pH 8.2 to make the staining mixture. Boehringer-Mannheim supply Fast Red tablets - one tablet is dissolved in Tris-HCl (pH 8.2) by vortexing, and then filtered through Whatman No. 2 paper before use. Sigma supply Sigma FastJ tablets for the substrate as well as the buffer. With all these sources of Fast Red, the background can be quite yellow-orange with chick embryos, particularly in the yolky cells of the extraembryonic regions. In our experience, the background problem is worst with the Boehringer-Mannheim tablets. The Vector kit gives the lowest backgrounds, provided that the kit is fairly new. Older kits ($ 1 year after purchase) reduce the signal and increase background considerably. It is best to use the Fast Red substrate to detect only the most abundant transcripts (mRNAs encoding transcription factors are usually good for this).

First remove the TBST from the embryos. Then rinse 2x 10 min in Tris-HCl (pH 8.2). Finally add the substrate as specified by the manufacturer. With the Vector kit, colour should develop within 0.5-3 hours. With the Boehringer tablets, it may take several days. After staining, stop the reaction by washing several times with PBS. The reduction in pH slighly improves the signal-to-background relationship, but in our experience the background is too high with this chromogen (from whichever source) for it to be useful in two- or three-colour detection protocols.

Detection of Alkaline Phosphatase with ELF⁷ (yellow-green fluorescence)

Recently, Molecular Probes Inc. has introduced this fluorescent substrate for alkaline phosphatase, which they supply in kit form. Two kits are available: E-6604 allows direct detection of alkaline phosphatase activity, while E-6605 also contains streptavidin-alkaline phosphatase conjugate allowing detection of biotinylated molecules. We find that avian embryos contain too much endogenous biotin and avidin and therefore backgrounds can be too high for use of the biotin-streptavidin system. However, the E-6604 kit can be used successfully and has also been used for mouse (Bueno et al., 1996) and zebrafish (Jowett and Yan, 1996) embryos. Kit E-6604 contains wash buffer (10x), blocking buffer, developing buffer, substrate and two additives, Hoechst 33342 dye (as a counterstain if desired), mounting medium and some plastic coverslips.

To stain whole-mounts of embryos with this kit, begin by washing the embryos three times for 10 min in PBS containing 0.5% Triton-X100 (since Tween-20 appears to cause the ELF crystals to be too large; see Jowett and Yan, 1996), and then follow the manufacturer's instructions supplied with the kit. Just prior to developing, place one or more embryos in a cavity slide (a glass slide with a concave depression or a home-made chamber made by attaching several layers of electrical tape to a glass slide and cutting out a hollow using a scalpel) in the pre-reaction solution. Then remove this solution and replace with 200-500 μl substrate working solution. Do not cover with a coverslip, since this will prevent convection and mixing of the substrate components. During incubation (at room temperature), check under a fluorescence microscope with UV illumination (350-380 nm excitation; >500 nm emission. DAPI/Hoechst filters are suitable) until the desired level of signal is attained. Molecular Probes state that signal should develop within 10 min to 1 hour, and recommend not exceeding a 2 hour incubation. However, weak signals may take up to 4-6 hours to develop. If no signal is visible after 2 hours at room temperature, it appears to be advantageous to place the embryos in a humid chamber at 37EC and check occasionally under fluorescence (it is best not to check too frequently, since continuous illumination may lead to photobleaching of the fluorochrome). After the desired incubation, stop the reaction by washing in PBS containing 25 mM EDTA and 0.05% Triton-X100 (pH 7.2). Then post-fix the embryos in 2% paraformaldehyde in PBS for 20 min and mount them with the materials supplied with the kit. Always perform ELF⁷ detection as the last step in a series.

Detection of Alkaline Phosphatase with other chromogens (magenta, pink, green and brick-red precipitates)

An analog of BCIP, 5-bromo-6-chloro-3-indolylphosphate (B8409 from Molecular Probes or MagentaPhos from Biosynth AG; Avivi et al., 1994) produces a magenta/mauve precipitate when reacted with alkaline phosphatase. The resulting colour can be fairly similar to that produced by NBT/BCIP, so it is not ideal for two-probe detection where the expression patterns overlap. It is also less sensitive than most of the other methods described here, so it can only be used to detect very abundant molecules. However, we have found that using Tetrazolium Red (Sigma) as a co-precipitant together with MagentaPhos makes the colour less violet and more red and slightly improves the sensitivity of the reaction. To use, simply replace MagentaPhos for BCIP and Tetrazolium Red for NBT in the protocol given above for the blue (NBT/BCIP) substrate.

A pink precipitate can be obtained using 6-chloro-3-indolyl phosphate (C8413 from Molecular Probes or SalmonPhos from Biosynth AG; Avivi et al., 1994). It is possible that Tetrazolium Red can enhance this substrate by acting as a co-precipitant but we have not experimented with this.

Another variation is to omit the NBT in the NBT/BCIP reaction, which gives a more greenish reaction product but there is significant loss of sensitivity. This can therefore only be used to detect very strongly expressed molecules, but its colour provides better contrast than NBT/BCIP when using either MagentaPhos or Fast Red to detect the other probe. Finally, INT (2-(4-iodophenyl)-3-(4-nitrophenyl)-5-phenyltetrazolium chloride; I6496 from Molecular Probes) can be used instead of NBT together with BCIP to give a brick-red precipitate.

Detection of Horseradish Peroxidase with Diaminobenzidine-H2O2 (brown precipitate)

We have never succeeded at detecting anti-DIG antibodies coupled to horseradish peroxidase in whole mounts, for reasons that remain mysterious. We therefore reserve this reaction for detection of embryonic antigens with antibodies. It is important to remember that it is not usually possible to combine in situ hybridization with whole mount detection of embryonic antigens if the latter are integral membrane proteins or anchored to the cell membrane, because the strong detergent washes and proteinase treatments used for detection of mRNAs lead to loss of these embryonic antigens. However, most intracellular antigens survive and retain antigenicity after the in situ procedure. When combining in situ hybridization with immunocytochemistry for embryonic antigens, the hybridization must be performed first because antibody solutions generally contain RNases that degrade single-stranded (but not hybridized, double stranded) RNA. However, following hybridization, it is possible to detect embryonic antigens and hapten-labelled probes in any order. It appears advantageous to detect embryonic antigens immediately following hybridization and before detecting the haptens, for the following reasons: (a) embryonic antigens are probably more labile and may degrade after several enzyme reactions and fixations, and can be affected by the acid glycine treatments used to inhibit alkaline phosphatase; (b) the diaminobenzidine (DAB) precipitate, once developed, is completely stable and is not removed by solvents or detergents, unlike many of the precipitates generated by alkaline phosphatase reactions.

To stain embryonic antigens in whole mounts in combination with in situ hybridization, take the embryos up to the post-hybridization washes and block them in blocking solution as described above. Then place them in an appropriate dilution of the primary (anti-embryonic antigen of choice) antibody in blocking buffer at 4EC, rocking, for 3-4 days (see Stern, 1993). After this, rinse 3 times and wash 3-4 times for 1 hour in TBST (Table 3) and place in appropriate dilution of the correct secondary antibody (e.g. anti-mouse IgG, coupled to peroxidase) overnight at 4EC, rocking. It is generally advantageous to preabsorb this secondary antibody against chick embryo acetone powder as described in Table 4. The next day, rinse 3 times and wash 3-4 times for 1 hour in TBST and then place embryos into a measured volume of 500 μg/ml diaminobenzidine (DAB[1][1]) in Tris-HCl (0.1 M, pH 7.4) for 1 hour in the dark. Then add H2O2 to a final dilution of 1:10,000 from a 100 VOL (30%) solution (first make a 1:100 dilution of the peroxide in Tris buffer, and then dilute this 1:100 in the DAB solution containing the embryos). The brown colour should begin to develop within 5-10 minutes and the reaction should be complete in less than an hour. At the end of this, rinse several times with distilled water or TBST and postfix for 20 min in 4% paraformaldehyde in PBS.

Antibody staining of cell surface or other labile antigens is possible before the in situ hybridisation procedure, provided that RNase activity is minimised. One way to do this is to include 1M LiCl in the initial fixation, blocking, antibody and washing solutions before staining with DAB/H2O2. We are just starting to experiment with this with some encouraging results but further development is needed.

Combining the colours

When performing detection of several molecules, the combination of colours chosen is very important. One criterion for choosing the colours is that they should contrast with each other so that they are easily distinguished. Another criterion, particularly when it is important to distinguish regions of expression that overlap with one another, is that the colours should not obscure one another. Clearly, these two criteria are partly contradictory. The best solution is to use fluorescent detection with fluorochromes emitting at two quite different wavelengths (see Jowett and Yan, 1996), because it is then easy to examine each of the two signals separately as well as to photograph them together.

In our experience, the blue product produced by the NBT/BCIP reaction is not sufficiently different from the magenta product generated by MagentaPhos to allow regions of overlap to be resolved, but this combination is good when expression is in adjacent regions (Fig. 2 A, C). The greeen product produced by BCIP alone is sufficiently different from MagentaPhos when the latter is used with Tetrazolium Red (see above), but both substrates lack the sensitivity to detect weakly expressed molecules. NBT/BCIP is sufficiently different from Fast Red to allow the simultaneous use of both of these substrates in ordinary light microscopy (however, the backgrounds produced by Fast Red can be quite orange-yellow and distracting; see Fig. 2B). At present, the best choice for detecting regions of overlap appears to be either a combination of MagentaPhos/Tetrazolium Red for the first probe with NBT/BCIP for the second probe for the light microscope (Fig. 2 A, C), or a combination of Fast Red as a fluorescent substrate with either NBT/BCIP (viewed by conventional light microscopy), if the second signal is weak, or ELF⁷ (viewed by fluorescence microscopy with UV excitation), provided that the signal is strong.

Time-table for multiple colour detection

Table 6 gives a time-table for performing in situ hybridization with two probes (using digoxigenin- and fluorescein-labelled riboprobes and alkaline phosphatase enzyme reactions with Fast Red and ELF fluorescence) and immunocytochemical detection of one intracellular antigen (using horseradish peroxidase) in the same embryos, as a guide to designing other experiments of this type. This table is only a guide, and it should be remembered that when using chromogenic reactions for the light microscope (e.g. NBT/BCIP, BCIP alone, Magenta Phos, etc.) the reactions may take several hours or even several days to develop. The time-table shown here uses HRP for the embryonic antigen, but this can be replaced by a secondary antibody coupled to Cascade Blue⁷ (Molecular Probes, Inc.) which gives a bright blue signal that is clearly separated from both the ELF and Fast Red emission wavelengths, so that three-colour fluorescence detection can be done. In this case, do not use the Hoechst counterstain provided with the ELF kit because this is not clearly separable from Cascade Blue.

Table 6 here

Photography

Whole-mounts that have been stained with chromogens suitable for light microscopy often benefit from being photographed in a dissecting microscope fitted with dark-field optics (Fig. 2 B-D). If such a microscope is not available, the embryos can be mounted in a cavity slide under a coverslip, placed over a matt black surface (such as a piece of black paper) and illuminated obliquely with a fibre optics source (taking care to avoid reflections from the coverslip). Other specimens benefit from being photographed in a conventional compound microscope, either with normal transmitted light (Köhler illumination; see Stern 1993c) or with Nomarski differential interference contrast optics. It is a good idea to experiment with different microscopes and illumination techniques for each combination of specimens, probes and chromogens. In our experience, the best film to use is Fuji 64T (tungsten balanced), but it is important to adjust the illumination so that the colour temperature of the light sources matches the sensitivity of the film to avoid excessively bluish, greenish or orange backgrounds (see Stern 1993c).

Specimens stained using fluorescent chromogens, or with a combination of fluorescent and visible light substrates, are best photographed on more sensitive film (e.g. Fuji Provia 1600) by taking multiple exposures onto the same frame. These may combine different fluorescence excitation filter sets for each fluorescent chromogen and transmitted light. However, it is important to take into account that in older microscopes, which do not have infinity-corrected optics, the path length of the light through different optical filter sets may differ, which affects the alignment of the images. A possible solution for this in some microscopes is to photograph the bright field image through a fairly colourless dichroic mirror and filter sets (see Stern 1993c).

Histological sectioning of whole-mounts

Of the chromogens used for light microscopy, only one (DAB) is completely resistant to organic solvents and can therefore tolerate dehydration, clearing, wax embedding and sectioning by conventional methods. Most of the other chromogens (NBT/BCIP, BCIP alone, MagentaPhos and Fast Red) are somewhat sensitive to organic solvents and therefore benefit from an accelerated dehydration and clearing process before embedding. After post-fixing the embryos, incubate for 5 min in absolute methanol (this intensifies some of these chromogens) and 10 min in propan-2-ol (2-isopropanol), followed by clearing in absolute tetrahydronaphthalene for 30 min. Then infiltrate in 1:1 tetrahydronaphthalene:paraffin wax for 30 min at 60EC and follow with 3-4 changes (30 min) of pure wax at 60EC before placing in a mould to set. After sectioning, the sections can be de-waxed in xylene or Histoclear as normal and mounted in Canada Balsam, Permount or DePeX.

By contrast, some other substrates (notably ELF⁷ and some other substrates marketed by Vector Labs) are very soluble in organic solvents and therefore cannot be wax embedded and sectioned. There are two possible solutions to this problem: (a) embed the stained embryos in gelatin and cut frozen sections in a cryostat or vibratome (see Stern 1993a), and mount these in an aqueous medium; (b) perform all staining with other chromogens first, wax embed and section as described in the previous paragraph, and then stain the sections with the ELF⁷ kit as described by Bueno et al. (1996). The first of these solutions has the advantage that embryos can first be photographed as whole mounts and then sectioned for further analysis.

Designing appropriate controls

With all in situ detection methods, it is important to be certain that the methods used are specific and sensitive enough to give a true image of the distribution of the molecules being studied. This is particularly important and difficult in the case of simultaneous detection of multiple molecules. Sense probes are often used as controls for in situ hybridization with riboprobes, but the nucleotide sequence of these is so different from that of the corresponding antisense probe that they are not really adequate controls. For example, sense probes do not allow the investigator to determine whether the sense probe is revealing overlapping expression between different members of a family of genes. Some workers check the specificity of the probe in Northern blots, but again this is not ideal because of the different hybridization and other conditions used in situ and in the blots. When studying the distribution of a new molecule in the embryo, it is best to start with a single probe and a well characterized and sensitive detection method (NBT/BCIP is probably the best). Ideally, two or more different antisense riboprobes should be synthesized, covering different portions of the mRNA, and these detected in separate batches of embryos to check that they give an identical distribution. If they don't, this could indicate cross-reaction with another gene product and/or the possibility of alternatively spliced transcripts (which should be visible in Northern blots).

When performing in situ detection of several molecules simultaneously, it is important to check that each detection method gives the same result as when the same molecules are detected singly. In particular, persistent alkaline phosphatase activity from detection of the first molecule could account for a false appearance of regions of overlap between two gene products (see Trouble-shooting below). To avoid this, it is valuable to do the following: (1) in some embryos, omit the antibody against the second probe - this should not produce the second colour; (2) perform reciprocal experiments with all combinations of colours (e.g. Fast Red and ELF, to detect a FITC and a DIG labelled probe, respectively, and then vice-versa) and labelling haptens (fluorescein, DIG, biotin) (see Jowett and Yan, 1996 for more details).

Trouble-shooting

Several problems may arise from the protocols given above. The following sections give some guidelines on how to diagnose and solve those most commonly encountered.

Embryos disintegrated or folded

The former is most likely to be due to rough pipetting when changing solutions. Avoid pipetting solutions directly onto the embryos, do not use a Gilson Pipetman but rather a Pasteur pipette with a good rubber teat, and avoid sucking up the embryos into the pipette. If this fails, reduce the Proteinase-K treatment time but do not lengthen the post-fixation after this step since this leads to some reduction of signal.

Folding of very young embryos (2 days or less) occurs if they are not initially fixed flat. Follow the instructions above and in Stern (1993a) for fixing embryos. If this is not the problem, it is most likely that they have been allowed to dry out too much between removing a solution from the vial and filling the vial again with the next solution. Change the solutions one vial at a time.

Background problems

It is often difficult to diagnose these accurately because there are many possible causes. Some of the most common are listed below.

Unincorporated hapten in the riboprobe

The most common source of background signals in in situ hybridization experiments is the presence of unincorporated hapten (fluorescein, digoxigenin or biotin) in the probe solution. These haptens appear to be "sticky" and can therefore cause significant background problems. Several ways to reduce the amount of free hapten have already been discussed above (see "Synthesis of labelled riboprobes"), including the use of spin-columns or re-precipitation in LiCl/Ethanol to clean the riboprobe, and re-use of the probe solution several times. Unincorporated fluorescein is particularly troublesome, but other haptens can also cause background.

"Trapping" within cavities

An intensely coloured precipitate can often develop within organs that contain internal cavities (e.g. the brain vesicles in 2-4 day embryos, heart, eye, gut, etc.). This seems to be a particularly serious problem with the NBT/BCIP substrate. I am unaware of the causes of this, but we have found that it can be alleviated substantially by perforating these cavities many times with a fine pin or a miniature knife during fixation of the embryo. This appears to improve the exchange of solutions during the washing steps and virtually eliminates this problem. If puncturing is not enough, the dorsal midline of the neural tube may be opened with a microscalpel in two or three places.

Cross-reactivity with other mRNAs

This is also a common problem, particularly when detecting a member of a large family of genes, and also when using short (# 300 nt) riboprobes where the stringency of hybridization has to be reduced (see above). It is difficult to control for this problem in every case, but it is recommended to determine the distribution of specific gene products by comparing results obtained with at least two different riboprobes derived from non-overlapping regions of the same gene (see "Designing appropriate controls" above). Use of the 3' untranslated end of the cDNA is often a good way to produce very specific probes (even ones that do not cross-react between closely related avian species, which is useful for analysing chick/quail chimaeras; see Izpisúa-Belmonte et al., 1993), but one has to be aware that there may be several alternatively spliced forms of the mRNA affecting this region, with different expression patterns.

Non-specific binding of antibodies

The commercially available anti-fluorescein and anti-digoxigenin antibodies purchased from Boehringer-Mannheim give very low levels of non-specific binding to avian tissues provided that they have been preabsorbed against chick embryo powder as described in Table 4, and that the embryos have been blocked efficiently prior to addition of antibody. If non-specific binding of the antibody remains a problem despite having preabsorbed it, the cause should be determined empirically by omitting one step of the protocol at a time.

Endogenous peroxidase or phosphatase activity

The protocol given in this chapter has been successful in our hands in eliminating all endogenous phosphatase and peroxidase activity that may interfere with the detection procedures. The bleaching step in 6% H2O2 following fixation of older embryos is designed to eliminate residual peroxidase, particularly in erythrocytes, as well as reducing eye pigmentation which may obscure signals in this region. The proteinase-K treatment and the high temperature of hybridization both contribute to reduce or eliminate phosphatases. If endogenous phosphatase remains a problem, the situation may improve by incubating the embryos in a 1 mM solution of levamisole for 1-2 hours prior to incubation in each alkaline phosphatase coupled antibody.

No signal or low sensitivity

Lack of signal, despite knowledge by the investigator that a particular gene is expressed at a given stage of development, probably indicates contamination with RNases (provided that other steps in the protocol have been carried out correctly and that the probe is good). We have found that the initial fixation step is important, but do not know the reasons for this. If no signal is detected or if only weak reactions are obtained, it is useful to start trouble-shooting by reducing the duration of the initial fixation of the embryos to just 30 min. In our experience, contamination with RNases during subsequent handling of the embryos is not a problem. If the steps follow one another swiftly, RNase-free solutions are only needed in the hybridization solution itself, and gloves need only be worn between the proteinase-K digestion and the end of the post-hybridization washes (up to the first TBST wash). Furthermore, the vials containing the embryos need not be RNase free because the proteinase-K digestion should remove these from the inner surfaces of the vial.

When detecting mRNAs expressed at low levels, the most sensitive detection techniques should be used (e.g. NBT/BCIP on digoxigenin-labelled probes), and detection of this mRNA should be carried out first. It is important to remember that signals will be stronger if the riboprobe is longer (and therefore contains more molecules of hapten per riboprobe molecule), so it is useful to design longer riboprobes for the more weakly-expressed molecule.

In the case of embryonic antigens, the use of a polyclonal antiserum for weakly-expressed antigens is preferable to a monoclonal antibody. Polyclonal secondary antibodies recognising several regions of the primary immunoglobulin can also be advantageous.

Persistent alkaline phosphatase activity from the first antibody

This can be an important problem when using several alkaline phosphatase labelled antibodies directed against different haptens, and detected sequentially. No method of inactivation is fool-proof. Those described to date include: (a) heating at 70EC overnight (this intensifies the blue colour of NBT/BCIP and the magenta of Magenta Phos, but can remove Fast Red and ELF reaction products); (b) fixation in 4% paraformaldehyde (not very effective if used alone); (c) incubation in 1mM levamisole (again not very effective if used alone); (d) two 15 min washes in absolute methanol (this will intensify NBT/BCIP and Magenta Phos as well as Fast Red to some extent, but will remove ELF); (e) incubation twice for 10 min in 100 mM glycine, pH 2.25 at room temperature.

In our hands, the acid glycine incubation, followed by incubation overnight at 70EC and then by fixation in 4% paraformaldehyde (as well as a short wash in methanol if alkaline phosphatase has been revealed by NBT/BCIP) works best. However, if the order of these steps is altered, the procedure becomes much less effective. Occasionally, we have found that the acid glycine breaks down double-stranded RNA with resulting loss of the labelling hapten. We do not yet know the reasons for the variability between experiments.

Acknowledgements

I am most grateful to Drs. David Ish-Horowicz, Juan Carlos Izpisúa-Belmonte, Randy Johnson, Chris Kintner, Hermann Rohrer, Jonathan Slack and Cliff Tabin for numerous tips and valuable advice over many years. Our research involving these methods is currently funded by the National Institutes of Health, the Muscular Dystrophy Association and the Human Frontier Science Program.

References

Avivi, C., Rosen, O. and Goldstein, R.S. (1994). New chromogens for alkaline phosphatase histochemistry: salmon and magenta phosphate are useful for single- and double-label immunohistochemistry. J. Histochem. Cytochem. 42, 551-554.

Bueno, D., Skinner, J., Abud, H. and Heath, J.K. (1996). Double in situ hybridization on mouse embryos for detection of overlapping regions of gene expression. Trends Genet. 12, 385-387.

Izpisúa-Belmonte, J.C., De Robertis, E.M., Storey, K.G. and Stern, C.D. (1993) The homeobox gene goosecoid and the origin of the organizer cells in the early chick blastoderm. Cell 74, 645-659.

Jowett, T. and Yan, Y.-L. (1996). Double fluorescent in situ hybridization to zebrafish embryos. Trends Genet. 12, 387-389.

Kamachi, Y., Sockanathan, S., Liu, Q., Breitman, M., Lovell-Badge, R. and Kondoh, H. (1995). Involvement of SOX proteins in lens-specific activation of crystallin genes. EMBO J. 14, 3510-3519.

Levin, M., Johnson, R.L., Stern, C.D., Kuehn, M. and Tabin, C.J. (1995) A molecular pathway determining left-right asymmetry in chick embryogenesis. Cell 82, 803-814.

New (1955) A new technique for the cultivation of the chick embryo in vitro. J. Embryol. exp. Morph. 3, 326-331.

Riddle, R.D., Johnson, R.L., Laufer, E. and Tabin, C. (1993) Sonic hedgehog mediates the polarizing activity of the ZPA. Cell 75, 1401-1416.

Sambrook, J., Fritsch, E.F. and Maniatis, T. (1989). Molecular cloning - A laboratory manual. 2nd Edition. New York: Cold Spring Harbor Laboratory Press.

Stern, C.D. (1993a). Avian embryos. In: Essential Developmental Biology: A Practical Approach. (C.D. Stern and P.W.H. Holland, eds.) IRL Press at Oxford University Press, Oxford. pp. 45-54.

Stern, C.D. (1993b). Immunocytochemistry of embryonic material. In: Essential Developmental Biology: A Practical Approach. (C.D. Stern and P.W.H. Holland, eds.) IRL Press at Oxford University Press, Oxford. pp. 193-212.

Stern, C.D. (1993c). Simple tips for photomicrography of embryos. In: Essential Developmental Biology: A Practical Approach. (C.D. Stern and P.W.H. Holland, eds.) IRL Press at Oxford University Press, Oxford. pp. 67-78.

Streit, A., Sockanathan, S., Perez, L., Rex, M., Scotting, P.J., Sharpe, P.T., Lovell-Badge, R. and Stern, C.D. (1997) Preventing the loss of competence for neural induction: roles of HGF/SF, L5 and Sox-2. Development (in press)

Uwanogho, D., Rex, M., Cartwright, E.J., Pearl, G., Healy, C., Scotting, P.J. and Sharpe, P.T. (1995). Embryonic expression of the chicken Sox2, Sox3 and Sox11 genes suggests an interactive role in neuronal development. Mech. Dev. 49, 23-36.

Yamada, T., Placzek, M., Tanaka, H., Dodd, J. and Jessell, T.M. (1991). Control of cell pattern in the developing nervous-system: polarizing activity of the floor plate and notochord. Cell 64, 635-647.

Figure legends

Fig. 1: Linearisation of the vector DNA and transcription with fluorescein-labelled UTP. A. Map of the commonly used vector pBlueScript II KS (Stratagene), and sequence of the polylinker region. The position of the T3 and T7 polymerase sites and direction of transcription from each are shown. B. Agarose gel, stained with ethidium bromide, showing size markers ("ladder"), linearised plasmid DNA ("DNA") and transcription reaction prior to DNAse treatment and precipitation with LiCl ("trans."). Note the large amount of unincorporated fluorescein-labelled UTP remaining, and the relationship between the amount of transcribed riboprobe and the amount of DNA template.

Fig. 2: Examples of results obtained with different combinations of probes and chromogens. A. Stage 10 chick embryo hybridised with a probe for the transcription factor Sox2 (expressed throughout the neural tube - Kamachi et al., 1995; Uwanogho et al., 1995; Streit et al., 1997) that had been labelled with digoxigenin and detected with MagentaPhos and Tetrazolium Red and with a probe for the secreted TGF-β-superfamily member cNR1 (=nodal; expressed in the left lateral plate - Levin et al., 1995) that had been labelled with fluorescein and detected with NBT/BCIP. The two colours can be distinguished easily, particularly because the domains of expression do not overlap. Bright field optics. B. Embryo at stage 8, hybridised with the same probes as in A., but this time Sox2 was visualised with Fast Red and cNR1 with NBT/BCIP. Note the disturbing yellowish background from the Fast Red, particularly in extraembryonic regions. Dark field optics (which makes the problem more acute). C. Here, an embryo at stage 5 was injected with a retroviral vector encoding the full-length goosecoid gene (Izpisúa-Belmonte et al., 1993). After 3 days' incubation, the embryo was fixed (about stage 23) and hybridized with a fluorescein-labelled probe for goosecoid (detected with NBT/BCIP) and with a digoxigenin-labelled probe for Sonic hedgehog (Riddle et al., 1994), detected with Magenta Phos and Tetrazolium Red. Dark field optics. D. Embryo overexpressing goosecoid as for C., hybridized with a goosecoid probe labelled with digoxigenin (detected with NBT/BCIP) and then processed by immunoperoxidase (with DAB/H2O2) using an antibody against a neurofilament-associated protein (NFp) (antibody 3A10; Yamada et al., 1991).

Table 1. Transcription mix:

Table 2. Hybridization solution:

Table 3. 10x TBST

Table 4. Preabsorbing antibody

To preabsorb antibody, proceed as follows:

(a) weigh X mg of embryo powder*, where X=2x number of mls of final vol. of Ab. solution needed into an Eppendorf.

(b) add 500 μl TBST and vortex for 20 seconds

(c) heat to 70EC for 30 minutes, vortex again 20 sec. and spin down at low speed for 1 min ‑ just enough to get the powder to form a loose pellet

(d) discard the supernatant

(e) wash the pellet 3-5 times with 500 μl TBST, spinning low speed each time (this is to remove any fat that may be floating on the supernatant. Repeat until no more fat is seen at the top of the supernatant

(f) resuspend the pellet in 100 μl blocking buffer (not TBST) for every bottle of embryos, mixing gently (not shaking)

(g) add antibody so that the final concentration will be 1:5,000. (For example: if final incubation in antibody overnight will have 1 ml per tube, the embryos will now be sitting in 1ml blocking buffer. If 5 bottles will be stained, add 1 μl antibody to the 500 μl blocking buffer with the embryo powder). Leave on a rocking table to absorb for 1-2 hours at room temperature.

(h) after absorbing antibody with powder, spin down at high speed (to make a hard pellet) in the microfuge for 3 min. Keep the supernatant and discard the pellet.

*Embryo powder is made as follows: Homogenise embryos (ideally of the same stage as those being stained) in a minimum volume of ice-cold PBS, through a syringe. Add 4 volumes of ice-cold acetone, mix and incubate on ice for 30 min. Centrifuge at 10,000 g for 10 min, discard the supernatant, and then wash the pellet with ice-cold acetone and spin again. Spread the pellet out and grind it into a fine powder on a sheet of filter paper. Air-dry the powder and store it at 4EC.

Table 5. NTMT

[1]Note: unreacted DAB is a potent carcinogen, but can be inhibited by treatment with bleach (sodium hypochlorite), which causes it to precipitate. Wear gloves while handling DAB and place any solutions or objects that have come into contact with it in a 1:50 dilution of household bleach in tap water to inactivate.