Visualization of gene expression patterns by in situ hybridization

Tags: embryos, room temperature, gentle agitation, FIXATION, TISSUE PREPARATION, probe, specimens, specimen, molten wax, 30 minutes, tissue sections, wax block, temperature, hybridization experiments, embedded tissue, probes, gene, cause tissue, autoradiography, hybridization, alkaline phosphatase, SSC-CHAPS, SSC, sections, wax blocks, EtOH, IN SITU HYBRIDIZATION, Oxford University Press, RNA, dripping wax
Content: Visualization of gene expression Patterns by in situ Hybridization Urs Albrecht1, Gregor Eichele1, Jill A. Helms2, Hui-Chen Lu1 1 Department of Biochemistry Baylor College of Medicine One Baylor Plaza Houston, TX 77030 2 Department of Orthopaedic Surgery University of California at San Francisco, San Francisco, CA 94143 Proofs to G. Eichele FAX (713) 798 3203, E-mail: [email protected] 1
1. INTRODUCTION 2. TISSUE PREPARATION 2.1. FIXATION 2.2. DEHYDRATION 3. IN SITU HYBRIDIZATION ON SECTIONS USING RADIOACTIVE PROBES 3.1. EMBEDDING 3.2. SECTIONING 3.3. PREPARATION AND SYNTHESIS OF THE RIBOPROBE 3.4 HYBRIDIZATION STEPS 3.4.1. PREPARATION OF SECTIONS 3.4.2. PREHYBRIDIZATION 3.4.3. HYBRIDIZATION 3.4.4. POSTHYBRIDIZATION WASHES 3.5. AUTORADIOGRAPHY AND VISUALIZATION 3.5.1. APPLICATION OF EMULSION 3.5.2. DEVELOPING 3.5.3. VIEWING AND PHOTOGRAPHY 3.6. TROUBLESHOOTING 4. IN SITU HYBRIDIZATION ON WHOLE MOUNTS 4.1. PREPARATION AND SYNTHESIS OF RIBOPROBES 4.2. FIXATION OF EMBRYOS 4.3. REHYDRATION, PROTEINASE K TREATMENT, POST FIXATION, AND ACETYLATION 4.4. PREHYBRIDIZATION AND HYBRIDIZATION 4.5 POSTHYBRIDIZATION WASHES, RNASE TREATMENT AND POST-RNASE WASHES 4.6. ANTIBODY INCUBATION AND WASHES 4.7. alkaline phosphatase COLOR REACTION 2
4.8. EMBEDDING AND SECTIONING W.M. EMBRYOS 4.8.1. PARAFFIN SECTIONS 4.8.2. FROZEN SECTIONS 5. REFERENCES 3
1. INTRODUCTION The identification of the expression pattern of genes in embryonic and adult tissues provides critical information about the temporal and spatial action of genes, and thus constitutes a first important step in understanding their function. The most straightforward method to visualize gene expression patterns in space and time is in situ hybridization on tissue sections or on whole mount specimens. The sole prerequisite for carrying out in situ hybridization is having tissue preserved by methods that maintain the integrity of mRNA, and having a fragment of expressed sequence. Such fragments can readily be prepared by subcloning of previously isolated cDNA clones or by RT-PCR methods 1,2. The principle underlying in situ hybridization is to anneal a labeled DNA- or RNA-probe to the cellular RNA made accessible either by sectioning of the tissue of interest or by making whole mounts permeable to the probes3-5. In situ hybridization is very sensitive in particular when antisense RNA probes are used. The procedure consists of four steps: (1) The preparation of tissue (fixation, dehydration, embedding and sectioning), (2) the preparation of the probe, (3) the application and annealing of the probe and subsequent removal of the non-hybridized probe by RNase digestion and washing, and (4) the visualization of the expression pattern (autoradiography, color reaction). Riboprobes used in in situ hybridization experiments are either labeled with 35[S] UTP or with a hapten (most frequently digoxygenin). In the first case, hybridization is detected by autoradiography, while in the second case antibodies are used to visualize the hapten. The decision of whether to determine a pattern of expression on sections or on whole mounts is largely determined by the nature of the specimen. Larger pieces of tissues or older embryos are usually not suitable for the whole-mount procedure. Once an epidermis covers the embryo, it becomes impermeable for most reagents including riboprobes. However, even larger specimens are fine for the whole mount procedure if the gene of interest is expressed on the surface of the specimen. Larger specimens can be dissected into parts, and those may be sufficiently permeable for the whole mount approach. For whole mounts, digoxygenin-labeled probes are used since autoradiography required for the detection of radiolabeled probes is not applicable to three 4
dimensional objects. Also note that whole mounts subjected to hybridization with a digoxygenin probe can be sectioned and examined under Nomarski optics. In situ hybridization on sections using 35[S] UTP labeled or digoxygenin labeled probes is a method ideal for larger specimens such as the brain. While offering less contrast than radiolabeled probes, digoxygenin probes can provide cellular resolution. It goes without saying that whole mounts and section approaches are complementary, and both may be required for a detailed expression analysis of a developmentally regulated gene. Essential to all in situ hybridization analyses is the use of controls that demonstrate probe specificity. A commonly accepted control is the use of sense riboprobes. The hybridization of antisense and sense riboprobes in parallel experiments and under identical conditions is particularly important if a gene appears to be broadly expressed. Once hybridization and washing conditions are established, hybridization with the sense control may be omitted. However, when a new tissue or significantly different embryonic stage is examined, then the use of sense probe controls is strongly recommended. It is difficult to determine which portion of a cDNA is most suitable for hybridization. If the gene of interest is member of a gene family, we advise the use of a riboprobe complementary to the 3' untranslated region which is usually not conserved. However, a riboprobe designed to recognize the protein coding region gives good results in most cases, since the protocols described here use an RNase digestion step followed by a stringency wash. Even a few mismatches are sufficient for RNase to cleave the probe leading to the removal of the cleaved probe fragments during the stringency wash. This chapter is designed to discuss each of the steps in sufficient detail in order to enable the reader to carry out in situ hybridization. However, we assume that the reader is sufficiently familiar with basic molecular biology techniques (subcloning, restriction analysis, electrophoresis etc.), and thus do not describe such fundamentals (for more information see ref. 6). The protocols we chose were successfully used in our laboratory using embryos and tissues from mouse7,8, chicken9, and human7,10. 2. TISSUE PREPARATION 2.1. FIXATION 5
The general purpose of fixation is to preserve good histological detail. Because the strength of the hybridization signal is dependent upon the availability of cellular mRNA that has been preserved in the tissue throughout the procedure, fixing protocols that cross-link excessively are not suitable. Glutaraldehyde and formaldehyde are particularly suitable for in situ hybridization protocols. It is important to keep in mind that not all tissues require the same amount of fixation, and that steps in the protocol are interdependent. For example, shorter fixation times or the use of gentle fixatives usually require a shorter protease digestion. A good and popular fixative for most embryonic and adult tissues is 4% paraformaldehyde in phosphate buffered saline (PBS). Fixations are usually performed at 4°C, with gentle agitation. All dissections are performed in sterile, ice-cold PBS. The tissue is rinsed in fresh PBS after dissection is complete, and placed into the chosen fixative for approximately 12 hours. Convenient containers for fixation include glass scintillation vials (20 ml volume), or if smaller tissues are being fixed, 24-well plates. While fixations of larger tissue pieces can be extended to days, care should be taken not to fix an excessive amount of tissue in a single vial. A useful guideline is that the volume of the fixative should be 30 times of the volume of tissue to be fixed. Alternative fixation agents include Bouin's or Carnoy's reagent that both have been successfully used in our laboratory for in situ hybridization of sections. Bouin's fixative is particularly useful when calcified tissues need to be sectioned (older embryos), since this fixative softens hard tissues. In the case of Carnoy's and Bouin's fixative, contents in the amniotic fluid become very solid and are hard to remove after fixation. Thus embryos should be washed in saline and extra embryonic membranes should be removed before fixation. PFA and MEMFA are equally suitable to fix embryos for whole mount studies. RECIPES MEMFA: 0.1 M MOPS (3-[N-morpholino]propanesulfonic acid), pH 7.4; 2 mM EGTA; 1 mM MgSO4; 3.7% formaldehyde. A stock solution is prepared not containing the formaldehyde. 1 part of a 37% formaldehyde stock solution is added to 9 parts of buffer/salt mix just prior to use. 4% paraformaldehyde/PBS: 100 ml of paraformaldehyde (PFA) fixative is prepared by heating 6
90 ml H20 and 100 µl 2 M NaOH in a microwave oven to 50°C. The solution must be basic to aid in the dissolution of the PFA. Add 4 g of PFA (use a face mask) and stir in a hood until the PFA is dissolved. Add 10 ml 10x PBS, filter through a Whatman paper filter and let cool to room temperature. If necessary, adjust the pH back to 7.4 by the addition of 100 µl 2 M HCl. If the PFA is too acidic, it will destroy DNA; if it is too basic it will destroy RNA. Store the solution at 4°C, sealed with parafilm to prevent fixation of other proteins stored in the refrigerator. If kept refrigerated, the solution is usually good for about one week. Carnoy's fixative: 100% EtOH (60 ml), chloroform (30 ml), acetic acid (10 ml) Bouin's fixative: saturated picric acid aqueous (75 ml), formalin (25 ml), acetic acid (5 ml). Suitable for larger embryos since this fixative decalcifies the ossified tissue. To fix mouse embryos older than 17 dpc, either the belly has to be opened up or the tissue of interest should be isolated by dissection from the rest of the body for better penetration of fixative. The skin of the embryo from this stage onwards is impermeable to the fixative. 2.2. DEHYDRATION During dehydration, water is removed from the specimen so it can be embedded in paraffin. Moreover, in a dehydrated state, specimens can be stored at -20°C for months without noticeable RNA degradation. Embryos are taken through a graded ethanol/salt series, beginning with physiologic salt concentration and ending in 100% ethanol (EtOH). The length of each wash is dependent upon the specimen size. Stage 6-14 chick embryos, or mouse embryos younger than day 9, are washed for 10 min per step. In the case of a stage 20 chick embryo or a day 11 mouse embryo, 15-30 min is sufficient; day 16 mouse embryos are washed for 60- 90 minutes in each step. Very large tissues should be dehydrated for 2-4 hrs per step, with the 70% EtOH step being extended overnight. This may need to be adjusted if the tissue is less permeable (i.e. from an older embryo or an adult). Transferring the specimen through the dehydration steps too rapidly results in specimens that cannot be sectioned. If water is not completely replaced by ethanol, or if the wax is too hot or the wax permeation period too short (see 3.1.), air bubbles develop and appear as holes in the paraffin sections which results in tearing of the section. 7
To dehydrate, subject tissue to the following steps at 4°C on a rocking platform moving at a gentle rate (20 times per min): 1. 0.9% NaCl 2. 30% EtOH in 0.9% NaCl 3. 50% EtOH in 0.9% NaCl 4. 70% EtOH in H2O 5. 90% EtOH in H2O 6. 100% EtOH At this point, embryos can be stored indefinitely at -20°C. Dehydration with Carnoy's and Bouin's fixatives: Carnoy's fixed tissues are transferred to 100% EtOH 15 min, 4°C (twice) and are stored in 100% EtOH at -20°C. Bouin's fixed embryos are transferred to 70% EtOH 15 min, 4°C. Wash in 70 % EtOH, replace until the yellow picric acid is gone, otherwise this agent crystallizes in the embryo. Transfer specimen to 95% EtOH, and then to 100% EtOH, 15 min, 4°C (each twice) and store in 100% EtOH at -20°C. Specimens to be used for whole mounts fixed in PFA are dehydrated as described above (steps 1 to 6) except that methanol is used instead of EtOH. If specimens were fixed in MEMFA, wash with 0.9% NaCl (2-5 min), transfer to 90% methanol and store at -20°C. 3. IN SITU HYBRIDIZATION ON SECTIONS USING RADIOACTIVE PROBES 3.1. EMBEDDING Paraffin wax should be melted ahead of time, either by placing a container in a warming oven overnight, or by microwaving the wax for approximately 10 min and leaving a small amount of wax unmelted. Care should be taken to not overheat the wax, since this destroys its ability to form the proper lattice structure when the wax cools. Discard unused wax after 2-3 days. A small aliquot of the wax should be placed in a separate beaker, and approximately 10 min before use an equal amount of xylene is added. Discard the unused portion of the xylene:wax solution immediately in a receptacle in a hood (xylene is toxic). Prior to embedding, process the tissues through the following solutions on a rocking 8
plate, using the same length of time as was employed when transferring tissue from 0.9% NaCl to ethanol: 1. 100% EtOH , at room temperature 2. EtOH/xylene 1:1, at room temperature 3. xylene, at room temperature 4. xylene/wax, at 58°C (does not need to be rocked) 5. wax, 3 times at 58°C After the final wax wash, prepare a suitable container for the tissue, such as a small weigh boat or a commercially available plastic mold. Gently swirl the vial to suspend the tissue in the melted wax, and quickly pour into the container; avoid creating bubbles in the process. The tissue can be oriented using forceps that have been warmed over an alcohol flame. If the tissue fails to pour from the vial, add more wax and try again. Bubbles in the molten wax can be removed by heating the forceps and gently moving it through the wax. Bubbles that are in close proximity to the tissue should be removed, as they will create voids in the wax that will make sectioning more difficult. The wax blocks should be allowed to cool at room temperature without disruption, and can be stored at room temperature indefinitely. Do not refrigerate or freeze wax blocks. Tissue fixed with Bouin's fixative and stored in 100% EtOH is embedded as follows. Transfer specimen to methylbenzoate with three exchanges at room temperature and after the last change wait until the specimen sinks to the bottom of the vial. Transfer to benzene (twice, room temperature), benzene wax (1:1, 58°C), wax ( twice, 58°C) and pour into mold. Methylbenzoate and benzene are used instead of xylene since the latter hardens the tissue and makes it too brittle for sectioning. HINTS AND PROBLEMS 1. To aid in positioning the tissues in wax, stain tissues with Eosin Y dye in EtOH prior to embedding. 1% stock of alcoholic eosine: 1 g of Eosin Y is dissolved in 20 ml of water and 80 ml of 95% EtOH are added. Prior to use, mix 1 part stock solution with 3 parts of 80% EtOH 2. Minimizing the length of the xylene step is advisable, since extended exposures cause tissue to become brittle. 9
3. Overheated wax shrinks after it has been poured, creating a depression in the wax block. Do not attempt to re-pour wax into this depression as it will not adhere to the original wax, causing separation when the wax block is mounted for sectioning. Instead, the entire wax block is melted by placing it back into the oven for the minimum time necessary to completely melt the wax. Repour and let cool to room temperature. 3.2. SECTIONING Mounting the tissue onto a wooden block (or an equivalent support) is easily performed if there is sufficient wax around all sides of the embedded tissue. Determine the plane of section that is desired, then spread a small amount of molten wax onto the block, and heat up a metal spatula until it is very hot. Place the warmed spatula against the wax on the block and simultaneously lower the tissue-containing wax block towards the top surface of the heated spatula. Quickly remove the spatula and firmly place the now-melted surface of the wax block into the heated wax on the wooden block. Re-heat the spatula and melt the four sides of the wax block slightly, allowing the dripping wax to fill in any space between the wooden block and the tissuecontaining wax block; allow sample to cool at room temperature. In order to produce a ribbon of tissue sections that is straight and wrinkle-free, the wax block must have 90° angles. If the bottom edge is uneven, or either side is longer than the other, the ribbon will twist making it difficult to position the sections onto a slide. Gradually shave down the wax block around the tissue with a clean razor blade, preserving enough wax at the periphery so that the tissue occupies approximately one-half of the total wax surface. Take care to remove only small amounts of wax at a time so that the block does not crack. Sections are best cut in a place with few air currents and constant temperature. Position a water bath (set between 45-50°C, use autoclaved water), and a slide warmer set at 38°C in close proximity. The position and angle of the microtome blade, the speed at which to cut and the optimal thickness of the section are determined empirically. To begin, use a blade angle of 10°, 7 µm thickness and a slow cutting speed. If the sections tend to wrinkle, it may be due to compression as a result of cutting too fast. Thus slow down when passing through the tissue. In 10
addition one can also reduce the angle of the knife. If the sections fail to form a ribbon and come off as individual sections, it is frequently due to the angle of the blade; adjust blade to a larger angle. If you detect a scratch or a split in the wax section, first check that there are no obvious nicks in the blade. If so, reposition the microtome blade so that a new zone is being used; use extreme caution when doing this so as not to cut yourself. Dust particles in the block underneath the specimen also can cause failure to form a ribbon. Shave a small amount of wax off of the bottom edge of the wax block (the edge where the knife first contacts the block). Once you have succeeded in making ribbons of sections, lay them onto a clean piece of paper. It is particularly helpful to use black paper since you can see the tissue in the wax with more contrast. After you have accumulated a number of ribbons, you can begin to mount them. Cut the ribbons into sections that are shorter than the slide you are using; the ribbon will expand approximately 10% as it warms up. Place the ribbon gently into the waterbath, taking care to place the side of the ribbon that faced the blade in contact with the water (shiny side of ribbon). Watch the ribbon closely; as the wax melts and expands, small wrinkles should disappear. Do not allow the water to get too warm as the wax will begin to melt, and in doing so will tear the tissue. To mount the tissue, position a slide underneath the ribbons and, using forceps to guide the sections, slowly lift the slide out of the water. Care should be taken not to touch the tissue since the wax will melt slightly and become adherent to the forceps. Allow the slides to dry on the slide warmer for a minimum of 2 hours, then store them in a cool (4 to 20°C), dry place. Include a desiccant in the box, then seal the box. Avoiding water condensation is critical, since in the presence of water, RNase can start cleaving the cellular RNA. RECIPES There are a number of protocols for treating slides to aid in adherence of the wax sections. We present two methods, one of which cleans and etches the slides slightly using TESPA (3aminopropyltrioxyethyl silane); these slides are stable indefinitely. The other method uses polyL-lysine; although the tissue adheres very well, poly-L-lysine is labile and the slides are more 11
difficult to prepare. TESPA: Place slides in a stainless steel rack and dip in 10% HCl/70% EtOH for 2 min, followed by distilled water and then 95% EtOH, 2 min each. Dry the slides either in a baking oven at 150°C for 15-20 min or in a speed-vac, then cool to room temperature. Dip slides for 10-15 sec. in 2% TESPA in acetone, followed by two quick washes in acetone and one in water. Dry at 42°C and place in a vacuum or desiccated box. Poly-L-lysine: Place slides in a stainless steel rack and wash in a 1% solution of Linbro ("7X") (Flow Labs Inc.) for 15 minutes. Rinse the slides extensively in deionized water (5 times, 2-3 min each), then bake them at 150°C for 1 hour. Cool to room temperature before immersing the slides in a freshly prepared solution of 100 µg/ml poly-L-lysine in autoclaved deionized water for 1 hour at room temperature. Allow the rack to drain and air dry for a few hours. Store in a vacuum or desiccated box for up to 2 weeks. 3.3. PREPARATION AND SYNTHESIS OF THE RIBOPROBE Single-stranded RNA probes are prepared using plasmid vectors containing a polylinker bordered by promoters of T3 and T7 bacteriophage RNA polymerases. Probe synthesis reactions are carried out using a linearized plasmid as a template. The plasmid is linearized by digesting with a restriction enzyme that cuts at a single site downstream of the insert. The site should be oriented so that the synthesized RNA probe will be terminated as a result of polymerase run-off. In order to synthesize a probe that hybridizes to mRNA, the antisense strand of the cloned insert must be used as the template. Following restriction enzyme digestion, the template is treated with proteinase K, then extracted in phenol/chloroform and precipitated in ethanol. This proteinase K treatment removes ribonucleases from the template reaction. The linearized template is then resuspended in diethyl pyrocarbonate (DEPC) treated water or TE buffer (10 mM TrisCl, 1 mM EDTA, pH 7.4). TEMPLATE PREPARATION cDNAs should be digested to yield both sense and antisense orientations of the template. The 12
sense probe serves as an important control for non-specific hybridization. At least 10 µg of DNA should be digested initially. Determine that the restriction digestion has gone to completion by running a small aliquot of the reaction out on an agarose gel; the presence of the uncleaved template will generate RNA transcripts that contain polylinker and vector sequence and may be a source of background signal. Add 50 µg/ml of proteinase K to the restriction buffer and incubate for 30 min at 37°C, followed by two phenol/chloroform (1:1 v/v) extractions and ethanol precipitation. Resuspend the digested, proteinase K-treated cDNA in TE made with DEPCtreated water. TRANSCRIPTION REACTION A typical transcription reaction is described which will generate high-specific activity probes (transcription kits are commercially available). Add the solutions in the following order, keeping the reagents on ice and using caution to avoid contamination with RNases (gloves, autoclaved tubes and tips, etc.). 6 µl of 5x transcription buffer (5x is 200 mM Tris-HCl pH 8.0, 40 mM MgCl2, 250 mM NaCl, 10 mM spermidine) 1 µl of 1 µg/ µl of restricted, proteinase K-treated, linearized DNA template add sufficient DEPC-treated water to make a final volume of 30 µl 1 µl of 10 mM rATP 1 µl of 10 mM rCTP 1 µl of 10 mM rGTP 1 µl of RNAsin (40 U/µl) 1 µl of 0.75 M DTT (use DTT that has been thawed once) 5 - 10 µl of 400-800 Ci/mmol, 10 µCi/µl 35[S] UTP Mix well, spin and then add 10 units of T3 or T7 RNA polymerase. Incubate at 37°C for 2 to 3 h. To remove the template DNA, use DNase I that is RNase-free, and add the following to the 30 µl transcription reaction: 19 µl of DEPC-treated water, 1.7 µl of 0.3 M MgCl2, 2 units of DNase I; incubate at 37°C for 15 min. 13
The probe is precipitated by adding the following to the transcription reaction: 100 µl of DEPC-treated water, 100 µl of 1 mg/ml yeast tRNA (previously purified by phenol extraction), 250 µl of 4 M ammonium acetate and 1 ml of EtOH. Vortex well, precipitate 5 min on ice, spin 10 min and remove supernatant. This precipitation is repeated once. Aspirate off the remaining supernatant using a pulled-out Pasteur pipet. A speed-vac can also be used for this purpose, however, use caution since RNA pellets may be difficult to resuspend if over-dried. Resuspend pellet in hybridization buffer (see Section 3.4.2.). Alternatively, unincorporated nucleotides are removed by passing the transcription reaction over a column containing RNase-free Sephadex G50. If hydrolysis is necessary to reduce the size of the transcripts (longer than 1- 1.5 kb while the optimal probe size is 0.5 - 0.8 kb), dissolve the probe in water and add an equal volume of 80 mM NaHCO3, 120 mM Na2CO3. Incubate at 60°C for t minutes, and t = (Lo - Ld)/0.11 Lo L, where Lo is the original probe length in kilobases and Ld is the desired probe length.
DETERMINATION OF INCORPORATION
Typically, greater than 50% of the input 35[S] UTP is incorporated into the RNA transcription
reaction. The use of labeled UTP as the single source of this nucleotide, limits the mass yield of
RNA probe. The following calculation is used to determine the amount of RNA generated in the
transcription reaction:
total number of µCi added x 13.2 x % incorporation________ =
ng of RNA
synthesized
number of 35[S]dNTP used x specific activity of radioisotope
Percent incorporation is determined by total counts divided by incorporated counts. Count an
aliquot of riboprobe and an aliquot of the reaction mix taken prior to precipitation in 2 ml water
miscible scintillation fluid in the 14C channel. The approximate specific activity of freshly
prepared RNA is expected to be 109 cpm/µg of RNA. If necessary, store the probe at -80°C for
up to a week (fresh probes are better since older probes display strand breaks).
14
3.4. HYBRIDIZATION STEPS 3.4.1. DEWAXING AND FIXATION OF SECTIONS All solutions used for prehybridization should be RNase free. This is achieved by autoclaving for 45 min. It is convenient to prepare 10x stock solutions, and make dilutions as required. Most of the prehybridization solutions can usually be reused if they are kept RNase-free. Paraformaldehyde (PFA) solutions can be re-used twice, but should be discarded thereafter. PFA is stored at 4°C. Proteinase K and acetylation solutions must be made fresh each time. We use a Tissue-Tek II staining station from Miles for the steps described here. It is advisable to cover all solutions containing ethanol with plastic wrap to prevent evaporation. Place slides (in a rack) in the following solutions in the order listed: 1. Histo-Clear (National Diagnostics) or another xylene substitute to remove wax, 10 min, twice. 2. Ethanol series, starting at 100% and progressing to 95%, 70%, 50%, 30%. Agitate the slides in each solution until equilibrated (approximately 1 min each). 3. 0.9% NaCl, 5 min 4. 1x PBS, 5 min 5. 4% PFA, 20 min (this solution can be re-used below) 6. 1x PBS, 5 min 7. Subject slides to proteinase K treatment: 20 µg/ml proteinase K in 50 mM Tris-HCl pH 7.6, 5 mM EDTA, 5 min. After this step, the tissue is particularly susceptible to RNases; use extreme caution. 8. 0.2 M HCl, 5 min (can be left out) 9. 1x PBS, 5 min 10. 4% PFA, 20 min 10. Acetylation: The purpose of this step is to acetylate amino groups in tissues to reduce electrostatic binding of probe12. Acetylation also blocks binding of probe to poly-L-lysine treated slides. These steps must be performed in a hood or another well-ventilated place. Suspend the slide rack above a rapidly rotating stir-bar in a solution of 0.1 M triethanolamine-HCl (TEA; pH 8.0). Add 600 µl of acetic anhydride; wait 5 min, then add 600 µl more of acetic anhydride. 15
Incubate for a total of 10 min. Care should be taken not to introduce water into the acetic anhydride stock; acetic anhydride would rapidly hydrolyze to acetic acid in the presence of water. Thus if the agent smells like acetic acid it should be discarded. 12. 1x PBS, 5 min 13. 0.9% NaCl, 5 min 14. 30% and 50% EtOH, 1 min each 15. 70% EtOH, 5 min (this step is longer to make sure to remove all salt deposits) 16. 80%, 95%, 100%, 100% EtOH, 1 min each Air-dry slides in a dust-free, RNase-free place, then store in a sealed container at room temperature with desiccant for up to a few days. Theoretically, de-waxed slides can be stored indefinitely; however, they are much more susceptible to RNase once the tissue has been treated with proteinase K and also prone to oxidation. 3.4.2. PREHYBRIDIZATION Sections may be incubated with hybridization buffer not containing the probe in order to reduce a high background; this may help especially if the mRNA of interest is rare. For 25 x 75 mm slides with 24 x 50 mm coverslips, 100 µl of hybridization mix per slide is sufficient. Be sure to prepare extra mix, because the viscosity of the hybridization mix leads to large pipetting errors. To the mix, add 1/100 th volume of 25 mM alpha-S-thio ATP (this is crucial for the reduction of background). The mix is then heated in 90 to 100 °C water for 2 min and cooled to room temperature. Carefully place the hybridization mix onto the dry slide, then lower a cover slip onto the slide with forceps. This is easiest if you drop one end of the coverslip onto the slide, then slowly lower the rest. Try to avoid bubbles, but if they do form, make sure that hybridization mix completely covers every section. Incubate slides in horizontal position for 1 hour at 50°C in a chamber whose bottom is filled with several layers of Whatman paper soaked in 50% formamide/2x SSC. We have used chambers consisting of transparent plastic obtained in hardware or department stores. We place the slides on a grid rack built from plastic pipets. Prehybridization has disadvantages. First, it necessitates removing the coverslip covering 16
the section which may damage the tissue. Second, the exact probe concentration can not be determined for the subsequent hybridization since hybridization buffer remains on the slide and will dilute the probe. The alternative, a hybridization performed at a higher temperature, can also decrease background and has the advantage of being quicker and not requiring the removal of the coverslip. RECIPE Hybridization buffer: 50% deionized formamide, 0.3 M NaCl, 20 mM Tris-HCl (pH 8), 5 mM EDTA, 10% dextran sulfate (heat gently and rock to dissolve), 0.02% Ficoll, 0.02% bovine serum albumin (RNAse free), 0.02% polyvinylpyrrolidone, 0.5 mg/ml yeast RNA (RNase free). Make 1 ml aliquots and store at -80°C until use. 3.4.3. HYBRIDIZATION An optimal signal-to-noise ratio is achieved when probe concentrations are just sufficient to saturate the complementary cellular mRNAs. However, this concentration is usually unknown, and the optimal probe concentration is therefore determined empirically. Initially, the amount of probe used in a hybridization reaction should be in 10-fold or greater excess of the transcripts of interest. A good starting place is to use a high specific activity probe (109 cpm/µg riboprobe) at a concentration of 2-6 x 106 cpm/slide (2-6 ng). We have found, however, that using half or even less (1/10th) of that amount can yield even nicer in situs when the expression of the gene of interest is confined to a small area. The correlation is not intuitive; it seems that the lower the expression level of the mRNA species, the lower the optimum concentration of the probe. It's probably a good idea to start off with the 1 ng per slide and then to subsequently titrate to find the optimal concentration. The rate of hybridization depends on the concentration of probe. We found that over night hybridizations are sufficient, although times as short as five hours have been successfully used. If the sections were prehybridized, remove the coverslip by tilting the slide and simply letting it slide off. A 26 x 75 mm slide can be covered adequately with 100 µl of hybridization 17
solution (50 µl are sufficient for prehybridized slides); calculate the appropriate amount of probe to add to the hybridization buffer, and place into an Eppendorf tube. Place the tube into a 70°C waterbath for three minutes. It is not necessary to denature RNA probes prior to hybridization. However, heating the hybridization buffer prior to pipetting is helpful due to the viscous nature of the solution. Add 1/10 volume of 1 M DTT, vortex well, spin and aliquot onto each slide. DTT is heat labile, and therefore should not be added to the solution prior to heating. Apply the mix with a Pipetman and gently lower a coverslip onto the slide using forceps by decreasing the angle between the coverslip and slide so that bubbles are forced out. The coverslips must be very clean to minimize bubbles. Do not press down on the coverslip to remove bubbles as this will result in a thinner layer of the hybridization solution. Place the slides into the humidified, sealed chamber (see 3.4.2.). Incubation temperatures are determined empirically; a good starting point is 45-50°C, which is approximately 25°C below the Tm of hybrids formed in situ with a 150 nucleotide long probe and a GC content of 50%. 3.4.4. POSTHYBRIDIZATION WASHES Washes are used to gently remove the coverslip, then to hydrolyze the non-specifically bound probe which has adhered to the slide and the tissue. The optimal temperatures of the washes must be determined empirically; however, a temperature of 50°C will provide the same stringency as the hybridization conditions. Higher temperatures, up to 65°C, can be used to improve background without significant loss of signal. Proceed as follows: 1. Coverslip removal: gently place the slides into a slide holder, using extreme caution to not dislodge the coverslips. Do not attempt to manually remove coverslips since the shear force is likely to damage the tissue. Place the slide rack into a solution of 5x SSC at 50°C, add 40 mM beta-mercaptoethanol (in a hood!). Gently agitate the slides every few minutes; after fifteen minutes the coverslips should have loosened sufficiently so that upon raising a slide out of the holder, the coverslip is left behind in the solution. Discard this solution in the radioactive waste. 2. Treat slides in 50 % formamide, 2x SSC, 40 mM beta-mercaptoethanol at 55-65°C for 30 min. 18
3. Wash slides once or twice in 0.5 M NaCl, 10 mM Tris-HCl (pH 8.0), 1 mM EDTA, 40 mM beta-mercaptoethanol for 30 min at 37°C 4. The non-specifically bound probe is hydrolyzed using 10 - 20 µg/ml RNase A in 0.5 M NaCl, 10 mM Tris-HCl (pH 8.0), 1 mM EDTA, for 30 min at 37°C. RNase A is stored as a stock solution of 10 mg/ml which can be refrozen. If background is a problem, the addition of RNase T1 (at 1 unit/ml) is recommended. 5. Slides are re-equilibrated in 0.5 M NaCl, 10 mM Tris-HCl (pH 8.0), 1 mM EDTA without RNase A for 15 min at room temperature, then subjected again to a high stringency wash (step 2 above) of 50% formamide/2x SSC, 40 mM beta-mercaptoethanol. 6. Wash once in 2x SSC for 15 min, then 0.1x SSC for 15 min, each at room temperature. 7. Dehydrate through an ethanol series: 30%, 50%, 70% EtOH/0.3 M NH4acetate, then in 95% and twice in 100% ethanol. Allow slides to dry. Step 2 (first stringency wash) and step 3 can be omitted if the probe behaves well (not "sticky"). 3.5. AUTORADIOGRAPHY AND VISUALIZATION To determine whether the stringency washes were of a sufficiently high temperature, and to determine the length of time that slides should be exposed to emulsion, the slides are first exposed to a high performance autoradiographic film for 1-3 days. Cronex film (DuPont) is suitable for this purpose. Tape the top edges of the slides to an immobile surface (a used piece of film is ideal) and use care not to allow the slides to come in contact with dust or lint present in the film cassette. Ideally, antisense probe hybridization should reveal distinct hybridization in the sections, possibly only in specific regions such as the nervous system, the heart, or some other subset of tissues. The sense control should be essentially free of signal except for a weak uniform haze on top of the section. If this is not the case, the temperature of the stringency wash needs to be increased. Some genes are broadly expressed as would be revealed with the antisense probe. If this is suspected, it is extremely critical that the sense control be essentially blank. Sometimes there is intense signal around the margins of sections or even on the slide. This can be due to the slides drying out during hybridization. Other causes are discussed in Section 3.6. 19
If it is determined that a higher wash temperature is needed to remove background, the slides can be rehydrated by placing them in 2x SSC for 15 min and then into a solution of 50% formamide/2x SSC/40 mM beta-mercaptoethanol. Rewashing can often remove non-specifically bound probes, however it rarely works as well as having washed initially at a higher temperature. After rewashing, dehydrate the slides as described under 3.4.4, step 5. 3.5.1. APPLICATION OF EMULSION Slides of suitable quality are then dipped in photographic emulsion. Emulsion is prepared by adding 118 ml Kodak NTB-2 emulsion to 200 ml deionized water. Read directions carefully when using this emulsion; most darkroom safelights are not suitable and will expose the emulsion; dark red (number 2) safelight is suitable. The emulsion and water must be warmed at 40-42°C for 30-35 min before mixing, using extreme caution to avoid exposure to any light source. It is convenient to aliquot the emulsion into glass scintillation vials, approximately 10 ml each. Wrap the scintillation vials in two layers of aluminum foil, then place inside a light-tight box. This box should be stored at 4°C and kept away from all forms of beta radiation. Melt an aliquot of the emulsion at 42°C for 5-10 min (keep this time to a minimum since the emulsion tends to break down with heat and time). Do not agitate. Once melted, gently pour into a suitable container such as a slide mailer (e.g. Young Laboratories) using care not to create bubbles. The slides are then grasped at the frosted end, and gently lowered into the emulsion for 3-4 sec; wipe the back of the slide to remove the emulsion and immediately place slide horizontally. The slides must be kept horizontal until the emulsion is completely hardened. This is dependent upon the air-flow in the chamber in which the slides are drying. If a simple box is used, the edges usually require taping to exclude all light. A more convenient option is to either obtain a desiccant box which is air-tight and place silica gel desiccant in it, or to have a box manufactured. If a box is manufactured, it is convenient to have a fan installed. However, note that emulsion can not be dried under vacuum, nor can it be heated as it tends to crack. Drying time is usually 5-7 hours at room temperature. Once dry, the slides can be stored in a sealed black box containing silica gel desiccant. 20
The box should be wrapped in aluminum foil, shielded from beta radiation and stored at 4°C. Use the exposure time of the Cronex film as a guide to determining the length of the emulsion exposure. Usually 3-4 times the length of the Cronex exposure is sufficient. Batches of emulsion should be tested prior to dipping tissue slides. Typically, the silver grain density should be so low as not to create a haze. Moreover, there should be no local clusters of silver grains. 3.5.2. DEVELOPING Remove the slides from 4°C and let them equilibrate to room temperature. Using only the appropriate dark red (number 2) safelight, place the slides into a rack and develop as follows: 1. Kodak D-19 developer, 2 min (do not warm past room temperature) 2. wash in water for 10 - 15 seconds 3. place in fixer for 5 min 4. final rinse in water for 10-15 min After developing, the slides can be counter-stained with a solution of 2 µg/ml Hoechst 33258 in water (stock solution of 10 mg/ml in DMSO, make 40 µl aliquots and store at -80°C). Dip the slides into the solution for two minutes, followed by two minutes in water. Air-dry the slides in a dark place, since the Hoechst stain is light sensitive. Once dry, overlay the slides with 50 µl of a solution of 5 g Canada balsam in 10 ml methyl salicylate, then place a coverslip and wick off the excess mounting medium from the edges. Keep slides flat for 1 week so that the mounting medium dries out. Other mounting media may be used which dry faster. However, some of those exhibit autofluorescence and cannot be used in combination with Hoechst staining. An alternative staining method not involving fluorescence detection involves staining with toluidine blue3. 3.5.3. VIEWING AND PHOTOGRAPHY Hoechst stain is a nuclear dye, which has the advantage of allowing a determination of cell density as well as visualizing tissue morphology. Using the DAPI channel on a epifluorescence microscope, at the same time that one views the sliver grains by transmitted light, allows one to 21
determine the tissue which is expressing the gene of interest. Hoechst dye eventually bleaches under fluorescent light, therefore care should be taken to minimize exposure. To clean the slides for photography, remove residual emulsion with glacial acetic acid on a Q-tip. Remove acid with ethanol. One may also use a new razor blade and scratch off the emulsion on the back of the slide; do not scratch the glass itself since this will cause background in the darkfield. Typically we obtain images of our in situ hybridizations specimens by conventional photography. In this case we take an image of the tissue revealed by blue Hoechst fluorescence and superimpose a darkfield image. To enhance the contrast, we insert a red filter into the light path so that the silver grains become red instead of white. More convenient than classical photographs are images captured with a video camera linked to a computer. These can be "black-and-white" images which are then pseudocolored using programs such as Adobe Photoshop. Such images can readily be stored on CDs and assembled into figures and be printed with a dye-sublimation printer. Electronically stored images can also be transmitted to other sites using computer networks. 3.6. TROUBLESHOOTING Poor tissue quality on sections: Brittle tissue which cracks upon sectioning usually indicates excessive time in xylene or too high temperature of the wax. If air bubbles are associated with the tissue it is an indicator that the tissue is moved too rapidly through the ethanol/xylene/wax solutions. Using old wax can also result in poor tissue embedding. Wrinkled sections, sections coming off of the slides : Wrinkled sections frequently result when the wax block is cut too quickly, or static electricity prevents the ribbon from moving away from the blade. Either slowing down the speed with which the blade passes through the knife, or wetting a Kimwipe and touching it to the wax block to decrease static electricity usually helps. Sections can come off the slides during the protocol when they were wrinkled during sectioning. In addition, inadequate post-fixation with PFA may be the cause; make sure to use fresh PFA. Low transcription from template : First check that the RNA pellet is completely resuspended by vortexing and heating slightly (50°C). Second, check that the appropriate polymerase was used 22
for a given template. Also, check whether the probe is intact by running a small aliquot out on a gel (using care to prevent RNA degradation by RNases). Lastly, extensive amounts of secondary structure which permits intramolecular annealing in the template DNA may prevent a good yield of high specific activity probe. In this case, a better probe may be obtained by using a shorter template or by using another region of the template. High background: This can result from a variety of problems. Too low of a wash temperature is a frequent problem. Try increasing the temperature 5 degrees in both the hybridization and wash steps. A major source of background signal can result when using antisense RNA probes that are contaminated with a small amount of sense strand RNA. A common source of such sense strand synthesis is the initiation of the RNA polymerase from the ends of the template DNA. Such aberrant initiation is more frequent when the ends of the template have a 3' overhang. If even a small amount of the probe is double stranded, it will be resistant to degradation by RNase A. Sense RNA can be isolated away from antisense RNA by gel electrophoresis as described in ref. 11. When using 35[S] labeled probes, either DTT or beta-mercaptoethanol are required to prevent oxidation of sulfhydryl groups. Both agents are heat labile. Therefore it is advisable to add these compounds just before adding the slides to a heated solution, using them once and discarding the unused portion. High background in emulsion: Usually this results from exposure of the emulsion to light or beta radiation. Check that the appropriate safelights are being used and the box in which the slides are being dried is light-tight. Be aware of watches that glow in the dark or beepers with lights. The distance between the safelight and the emulsion should be a minimum of 4 ft. Emulsion removal: At times it may be necessary to remove the emulsion layer from the slide. For example, if the unprocessed emulsion layer has been unintentionally exposed to light, or the processed emulsion layer has high background and the specimens are of sufficient value to warrant recovery. Although it is difficult to remove the emulsion without some damage to the underlying tissue because of the use of alkaline solutions, the following protocols will allow removal of most of the emulsion: 23
If the emulsion has not been hardened by photographic developing, it is reasonably soft and can be removed by placing the slides in a 60°C waterbath for a few minutes to soften the emulsion. Rinse the emulsion off by gentle agitation in the waterbath or under running tap water. Place the slides in Kodak fixer for 3 min to remove the faint bluish stain, followed by washing for 5 min in Running Water to remove all traces of the fixer. After drying completely, the slide can be re-dipped in emulsion. If the emulsion layer has been hardened by photographic processing, it is much more difficult to remove it. First digest the slide for 2-5 min in a 1% solution of NaOH or KOH at room temperature. Any remaining silver grains in contact with the tissue can be removed by placing the slides in a silver reducer such as 7.5% potassium ferricyanide. This is followed by a water rinse, then 3 min in Kodak fixer and 5 min of water washing. The slides can then be redipped. 4. IN SITU HYBRIDIZATION ON WHOLE MOUNTS (W.M.-ISH)3-5 4.1. PREPARATION AND SYNTHESIS OF RIBOPROBES Digoxigenin-labelled riboprobes are prepared in a similar way as radioactive-labelled riboprobes (Section 3.3.). TEMPLATE AND TRANSCRIPTION REACTION It is better to have long inserts (0.6 kb) to increase the sensitivity of detection of transcripts which are expressed broadly or expressed at low levels. The procedure described here will work well with riboprobes ranging from 0.3 to 1.2 kb. Single-stranded riboprobe is prepared by the transcription of linearized plasmid template using digoxigenin-tagged rUTP in the substrate mix. We include 0.2 µCi of -32[P]rUTP as a tracer to determine the yield of the reaction. 4 µl of 5x transcription buffer (5x is 200 mM Tris-HCl pH 8.0, 40 mM MgCl2, 250 mM NaCl, 10 mM spermidine) 1.5 µl of 1 µg/ µl of restricted, proteinase K-treated, linearized DNA template 24
add sufficient DEPC-treated water to make a final volume of 20 µl 2 µl of 10 mM rATP 2 µl of 10 mM rCTP 2 µl of 10 mM rGTP 1.3 µl of 10 mM rUTP (can be replaced by dig-11-rUTP) 0.7 µl of 10 mM dig-11-rUTP 0.5 µl of RNAsin (40 U/µl) 2 µl of 0.1 M DTT (use DTT that has been thawed once) 1 µl diluted -32[P]rUTP (0.2 µCi per µl) Mix well, spin and then add 40 units of T3 or T7 RNA polymerase. Incubate at 37°C for 2.5 to 3 h. Digest template by adding 2 µl of DNase I and incubate for 15 minutes at 37°C. Take 1 µl aliquot of synthesized probe to measure yield of incorporation (probe can be stored on ice or at -20°C meanwhile). DETERMINATION OF INCORPORATION Spot two 0.5 µl aliquots of reaction on two Whatman GF/C glass-fiber filters and let filters air dry. Transfer one filter into 200-300 ml ice cold 5% TCA containing 20 mM sodium pyrophosphate. Swirl the filters in this solution for 2 minutes. Wash two more times. Transfer the filter to 70% EtOH, wash briefly, then let it air dry. This filter and the non-washed filter are placed into scintillation vials, cocktail is added, and the radioactivity is measured in the 32[P] channel. 100% incorporation of all precursors, with substrate concentrations as specified above, should give 26 µg RNA. Yields generally range from 35-45%, but one can use probes even if incorporation is 10 - 20%. A transcription reaction should synthesize about 10 µg probe. PROBE PURIFICATION Precipitate the riboprobe by adding 10 µl 7.5 M NH4OAc and 60 µl 100% ice-cold EtOH. Precipitate at -20°C, wash, air dry the pellet as usual (see also Section 3.3). Resuspend in 10 µl distilled water, then add hybridization buffer to make a solution of 10 µg/ ml. For this and the 25
subsequent steps, water should be sterilized, but not DEPC treated, since this agent interferes with some of the later steps. Probe can be stored at -20°C for months. 4.2. FIXATION OF EMBRYOS Dissect embryos in ice-cold PBS. Remove the tissues (extraembryonic membranes) that will trap reagents. In addition open up cavities (brain, heart) which would otherwise trap reagents. Embryos are fixed for 90 minutes in MEMFA at room temperature. Embryos are then rinsed once in 0.9 % NaCl for 2 minutes, and transferred to 90% MeOH, a medium in which they can be stored at 20°C. In these and the subsequent manipulations we use 1 ml transfer pipets. Embryos become very sticky and fragile after proteinase K treatment; be careful not to damage the embryo when removing liquids or when embryos need to be transferred. Use a black background to visualize the embryos. 4.3. REHYDRATION, PROTEINASE K TREATMENT, POST FIXATION, AND ACETYLATION REHYDRATION Embryos are rehydrated by passage through the following solutions for 10 minutes each step: 1. 75% Methanol/H2O 2. 50% Methanol/H2O 3. 25% Methanol/75% PBT (PBT= PBS, 0.1% Tween 20, first dilute detergent to 10% with water) 4. PBT twice PROTEINASE K TREATMENT AND POSTFIXATION Embryos are transferred to a small petri dish (3.5 cm diameter) containing PBT at room temperature. Remove most liquid and add freshly prepared protease K solution (10 µg/ml) to the 26
dish. Embryos need to be well immersed and are incubated for 3-20 minutes according to their dimension (e.g. chick embryos Hamburger-Hamilton stage 4: 3 minutes; HH stage 8: 6 minutes; for HH stage 22: 20 min). Protease batches can vary in strength; incubation times should thus be optimized empirically. The protease stock should be aliquoted and stored at -20°C. Following proteinase K treatment, rinse twice with PBT. Transfer into 5 ml 100 mM glycine (in PBT, adjust pH to 7.2) and incubate for 5 min at room temperature. Wash twice in PBT, 5 min each. Embryos are then refixed for 30 minutes at room temperature in 4% paraformaldehyde-PBS containing 0.1% Tween (0.2% glutaraldehyde may be added to this fixation if the tissue is particularly fragile, such as early embryos). This is followed by two 5 min washes in 5 ml of PBT. 4.4. PREHYBRIDIZATION AND HYBRIDIZATION Transfer the embryos into 7 ml glass vials with Teflon lined caps. Remove most of the PBT solution followed by underlying the residual PBT with 2 ml of hybridization buffer. Allow embryos to sink into the hybridization solution (may take 20 to 30 min) and remove as much of the upper layer (PBT) as possible, then mix. After 10 minutes remove the hybridization buffer, then add 2 ml fresh hybridization buffer and incubate for 4 hours at 60°C in a very gently rotating water bath (18 rpm). Place vials at room temperature for 5 minutes, and carefully remove the hybridization solution. Add 1.5 ml hybridization buffer containing 0.2 µg/ml digoxigenin riboprobe (1/50 volume of stock). Thoroughly mix by swirling (or inverting if a teflon-lined cap is used) and incubate the embryos overnight in a 60°C water bath. Smaller volumes of hybridization buffer can be used and the probe mix can be reused once. RECIPE Hybridization buffer: 50% deionized formamide, 5x SSC, 5 mM EDTA, 1 mg/ml yeast tRNA (purified with phenol/chloroform, followed by EtOH precipitation), 100 µg/ml heparin, 1x Denhardt's solution, 0.1% Tween 20, 0.1% CHAPS (3-[3-cholamidopropyl)-dimethylammonio]-1-propanesulfonate, Boehringer). 27
4.5 POSTHYBRIDIZATION WASHES, RNASE TREATMENT AND POST-RNASE WASHES POST HYBRIDIZATION WASHES Unless otherwise specified, the washes are performed in a 60°C water bath rotating at a rate of 18 rpm. The washes need to be thorough but gentle. It is important to allow embryos to settle to the bottom of the tube and leave some liquid above them when removing the wash solutions, otherwise the embryos will be flattened and damaged. 1. Rinse embryos twice in 5 ml FSC (50% formamide; 2x SSC; 0.1% CHAPS; 50 mM glycine, pH 7.2) at room temperature. 2. Wash twice in 5 ml FSC, 30 minutes per wash. 3. Wash twice in 5 ml 2x SSC-CHAPS (2x SSC; 0.3% CHAPS; 50 mM glycine pH 7.2), 5 minutes per wash. 4. Resuspend in 1.8 ml SSC-CHAPS (1x SSC; 0.3% CHAPS; 50 mM glycine pH 7.2) at room temperature. RNASE TREATMENT The reaction is carried out for 35 minutes at 37°C in SSC-CHAPS containing 4 µg/ml RNase A (RNase A is dissolved in water at a concentration of 1 mg/ml and frozen at -20°C in aliquots, do not boil the stock) and 20 units/ml RNase T1. Specifically, mix in an Eppendorf tube 191.6 µl SSC-CHAPS, 0.4 µl RNase T1 (100,000 units/ml), 8 µl RNase A (1 mg/ml) and add this RNase cocktail to the embryos. Mix gently and incubate 35 minutes at 37°C, agitating occasionally. POST-RNASE WASHES 1. Wash once in 5 ml 2x SSC-CHAPS at room temperature for 5 min. 2. Wash briefly in 5 ml FSC at room temperature. 3. Wash twice in 5 ml FSC at 60°C with gentle agitation, 30 minutes per wash. 28
4. Wash once in 5 ml 2x SSC-CHAPS at 60°C for 5 minutes. 5. Wash twice in 0.2x SSC-CHAPS (0.2x SSC; 0.3% CHAPS) at 60°C for 15 minutes. 6. Wash once in PBT-CHAPS (PBS; 0.1% Tween 20; 0.3% CHAPS) at 60°C for 5 minutes, cool to room temperature. Note: 60°C works well for most of the probes. If the signal is rather strong and localized, RNase treatment and post-RNase washes can be substituted by two washes with 0.2x SSC-CHAPS for 30 minutes at 55-65°C and then a single wash with PBT-CHAPS for 5 minutes at 55-65°C. However, if this short-cut is applied, the temperature for hybridization has to be optimized. 4.6. ANTIBODY INCUBATION AND WASHES To visualize the digoxygenin hapten, a high-affinity antibody from Boehringer is used. To prevent non-specific binding of the antibody, embryos are pre-blocked with lamb serum. In addition, the polyclonal antibody may be pre-absorbed to embryo powder or to embryos, but this step is not always required. 1. Wash three times in 5 ml PBTB (PBS; 0.1% Triton X-100; 2 mg/ml BSA). 2. Add 1 ml of PBTB containing 40% lamb serum which has been heat inactivated at 55°C for 30 minutes. Block for 1 hour at 4°C, gently rocking the specimens. 3. Replace liquid with 2 ml of PBTB-40% lamb serum containing a 1/2,000 dilution of sheep anti-digoxigenin Fab antibody conjugated to alkaline phosphatase. This solution can be reused more than once. Include 0.1% sodium azide to prevent bacterial growth during storage at 4°C. 4. Incubate overnight at 4°C with gentle rocking. 5. Rinse briefly with PBTB 3 times at room temperature. 6. Rinse 4 times with PBTB for 1 hour per wash, with gentle agitation on a rocking platform. 4.7. ALKALINE PHOSPHATASE COLOR REACTION 1. Rinse the embryos briefly with 4 ml alkaline phosphatase buffer twice (100 mM Tris, pH 9.5; 100 mM NaCl; 50 mM MgCl2; 0.1% Tween 20). 2. Add 3 ml alkaline phosphatase buffer into the vial and wrap it with aluminum foil to prevent 29
light exposure. Color reagents: mix 1 ml alkaline phosphatase buffer with 18 µl nitro blue tetrazolium (75 mg/ml in 75% dimethylformamide), and 14 µl BCIP (5-bromo-4-chloro-3indolylphosphate, 50 mg/ml in dimethylformamide). Add this color reagent mixture to the embryos and gently agitate in the dark for 3-9 hours at room temperature. 3. Observe the development of the stain briefly every half hour and try to catch the best staining (strong signal and low background). Slightly overstain the embryos since some of the background stain will be washed off in the subsequent treatments. The embryos should be examined only briefly, since prolonged illumination will turn the yellow reaction mixture brown, and this may increase the background. 4. Stop the color reaction by post-fixation of the embryos in MEMFA for 30 minutes at room temperature, then rinse specimens in 0.9% NaCl for 2 minutes, and transfer to 90% MeOH. If the background is high, subject embryos to several washes in 90% MeOH at 4°C with gentle rocking (overnight to 2-3 days). The stained embryos can be stored at -20°C. Embryos can be restained if they have not been post-fixed by MEMFA. The embryos that require restaining are kept in PBT at 4°C. 4.8. EMBEDDING AND SECTIONING W.M. EMBRYOS The stained embryos can be photographed in a dissecting microscope with bright- or darkfield illumination. They can subsequently be embedded and sectioned to examine the details of the expression patterns. Although the reaction product using NBT/BCIP is soluble in many organic solvents, it is stabilized by fixation with formaldehyde and the embryos can then be embedded in paraffin wax or OCT in the case of frozen sections. 4.8.1. PARAFFIN SECTIONS 1. Transfer the embryos to 100% EtOH for 10 minutes at room temperature. 2. Transfer to xylene for 5 minutes, twice. 3. Transfer to xylene/paraffin wax (prewarmed at 58°C) for one hour at 58°C. 4. Transfer to paraffin wax (prewarmed at 58°C) and incubate overnight at 58°C. 30
5. Embed the embryo as described in section 3.1. 6. The embryo is sectioned at 10 µm thickness and the sections are placed on Superfrost slides (Fisher) as described in section 3.2. 7. Dewax the slides with xylene 10 minutes twice and then progressively rehydrate until sections are in water. 8. Mount the slide with Biomeda's Crystal/Mount (Cat. No. M02). Place the slides horizontally in an oven set at 70-80°C for at least 10 min. Remove the slides and allow them to reach room temperature. These slides are now ready for microscopy. 4.8.2. FROZEN SECTIONS 1. Rehydrate the embryos progressively to PBS, and rinse with PBS briefly three times. 2. Let the embryos lay on top of 30% sucrose/phosphate buffer (10 mM Na2HPO4, 1.8 mM KH2PO4, pH 7.4 ). Keep at 4°C until the embryos sink to the bottom. 3. Put the embedding mold on top of smashed dry ice and put some OCT into this cooled mold. Place and orient the embryo in the OCT, and wait for it to solidify. 4. When OCT is hardened, cut away the extra OCT. Put some OCT on top of the sample holder of the cryostat at room temperature and place the trimmed tissue block on top of it. The tissue block should be glued by the OCT. Put the sample holder rapidly into crushed dry ice and let it cool so that the tissue block firmly attaches to the holder. 5. Section at 10 µm thickness and mount these sections on Superfrost slides (Fisher) as described in Section 3.2. 6. Air dry the slides completely. 7. Cover the entire slides with PBS for 1 minute and remove PBS by touching the margins on top of paper towels. 8. Put three drops of the Biomeda Crystal/Mount (Cat. No. M02) and mount the slide as described above. Acknowledgements 31
We would like to thank Drs.Ariel Ruiz i Altaba, Clementine Hofmann, Beat Lutz, Olof Sundin and Mr. Calvin Wong who have contributed to the development of these protocols. This work was supported in part by funds from the National Institutes of Health (GE, JH) and from the Swiss National Science Foundation (UA). 32
5. REFERENCES 1. Koopman, P. Analysis of gene expression by reverse transcriptase-polymerase chain reaction. Essential Developmental Biology. A practical approach. Eds. Stern, C.D., Holland, P.W. Oxford University Press, Oxford, 233-242, 1993. 2. Holland, P.W. Cloning genes using the polymerase chain reaction. Essential Developmental Biology. A practical approach. Eds. Stern, C.D., Holland, P.W. Oxford University Press, Oxford, 243-256, 1993. 3. Wilkinson, D.G. In Situ Hybridization. Essential Developmental Biology. A practical approach. Eds. Stern, C.D., Holland, P.W. Oxford University Press, Oxford, 257-276, 1993. 4. Wilkinson, D.G. In Situ Hybridization. A practical approach. Ed. Wilkinson, D.G., Oxford University Press, Oxford, 1992. 5. Li, H.-S., Yang, J.-M., Jacobson, R.D., Pasko, D., Sundin, O. Pax-6 is first expressed in a region of ectoderm anterior to the early neural plate: Implications for stepwise determination of the lens. Dev. Biol., 162, 181-194, 1994. 6. Sambrook, J., Fritsch, E.F., Maniatis, T. Molecular Cloning. A Laboratory Manual. Cold Spring Harbor Laboratory Press. New York. 1989. 7. Reiner, O., Albrecht, U., Gordon, M., Chianese, K.A., Wong, C., Gal-Gerber, O., Sapir, T., Siracusa, L. D., Buchberg, A. M., Caskey C. T. and Eichele, G. Lissencephaly gene (LIS1) expression in the CNS suggests a role in neuronal migration. J. Neuroscience 15, 3730-3738, 1995. 33
8. Parker, S., Eichele, G., Zhang, P., Rawls, A., Sands, A.T., Bradley, A., Olson, E.N., Harper, J.W.and Elledge, S.J. p53-independent expression of p21Cip1 in muscle cells and other terminally differentiating cells. Science 267, 1024-1027, 1995. 9. Lutz, B. Kuratani, S., Cooney, A.J., Wawersik, S., Tsai, S.Y., Eichele, G., and Tsai, M.-J. Developmental regulation of the orphan receptor COUP-TFII gene in developing spinal motor neurons. Development 120, 25-36, 1994. 10. Lutz, B., Kuratani, S., Rugarli, E. I., Wawersik, S., Wong, C., Bieber, F.R., Ballabio, A., and Eichele, G. Expression of the Kallmann syndrome gene in human fetal brain and in the manipulated chick embryo. Human Molecular Genetics 3, 717-723, 1994. 11. Krieg, P.A. and Melton, D.A. In vitro RNA synthesis with SP6 RNA polymerase. Methods in Enzymology, 155, 397-415, 1987. 12. Hayashi, W., Gillam, I.C., Delaney, A.D.and Tener, G.M.. Acetylation of chromosome squashes of Drosophila melanogaster decreases the background in autoradiographs from hybridization with 125I - labeled RNA. J. Histochem. Cytochem 26, 677-679, 1978. 34

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