Chapter 18 Transgenic Mice: Production and Analysis of Expression

CHAPTER 18

Transgenic Mice: Production and Analysis of Expression Alexander Faerman and Moshe Shani Institute of Animal Science Agricultural Research Organization The Volcani Center Bet Dagan 50250, Israel

I. Introduction 11. Production of Stable Transgenic Mice A. Choice of Expression Vectors and DNA Preparation for Microinjection B. Superovulation C. Media Preparations D. Embryo Recovery E. Preparation of Holding and Microinjection Pipets F. Microinjection: Equipment and Setup G. Vasectomy and Embryo Transfer H. Identification of Transgenic Mice 111. Production of Transient Transgenics A. Advantages IV. Analysis of Expression at the R N A Level A. RNA Isolation B. Northern Blots C. RNase Protection Assay D. RT-PCR E. In Situ Hybridization to Tissue Sections F. Whole-Mount in Situ Hybridization of Mouse Embryos V. Analysis of Expression at the Protein Level A. CAT Assay B. LacZ Staining VI. Future Directions References

METHODS IN CELL BIOLOGY. VOL. 52 Copyright 0 1998 by Academic Prrrr. All ngho i w ~ i - h 7 9 ~ / 9snx o n

of

reproduction in any form reserved

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I. Introduction Production of transgenic mice become one of the most powerful tools to dissect complex developmentalprocesses such as genetic determination, pattern formation, cell lineages, cis-acting elements controlling gene expression, etc. In the study of myogenesis, transgenesis was exploited to address several important topics:

1. The cis-acting DNA sequences involved in the developmental regulation and tissue specificity of structural and regulatory genes, Elements controlling the expression of the skeletal muscle actin (Shani, 1986; Petropoulos et al., 1989; Asante et al., 1994), troponin (Hallauer et al., 1988; Banerjee-Basu and Buonanno, 1993), myosin light chain 2 (Shani, 1985), M creatine phosphokinase (Johnson et al., 1989), aldolase (Salminen et al., 1994), acetylcholine receptor (Gundersen et al., 1993), myogenin (Cheng et al., 1992, 1993; Yee and Rigby, 1993; Fujisawa-Sehara et al., 1993), and MyoD (Goldhamer et al., 1992, 1995) were characterized. 2. The mechanisms confemng fiber-type specificity. Transgenic mice carrying the rat troponin I slow gene (Banejee-Basu and Bounanno, 1993),the quail fast skeletal muscle troponin I (Hallauer et al., 1993), the rat fast skeletal muscle myosin light chain 1 (Donoghue et al., 1991), and the human aldolase A genes (Salminen et al., 1994) revealed a complex transcriptional requirements for the different muscle fibers. 3. The consequences of overexpressing muscle specific genes in skeletal muscle. To this end transgenic mice overexpressing the B subunit of creatine b a s e (Brosnan et al., 1993), the c-ski (Sutrave et al., 1990), or the rat myosin light chain 2 (Shani et al., 1988) genes were studied. 4. The consequences of ectopic expression of the myogenic regulatory genes MyoDl (Faerman et al., 1993) or myf5 genes (Miner et al., 1992; Santerre et al., 1993). 5. Delineating the cis-acting elements responsive to regulation by the nerve (Banerjee-Basu and Buonanno, 1993; Buonanno et al., 1993). 6. The possible existence of regional specification along the rostrocaudal axis (Donoghue et al., 1992, Grieshammer et al., 1992). 7. The efficiency of correcting muscular dystrophy in mdu mice (Matsumura et af., 1993). The scope of the present chapter is to describe in detail the methodology of transgenic mouse production, mainly focusing on the most commonly used technique of microinjecting into fertilized eggs, based on the experience of our laboratory. This chapter also describes methods for the analysis of transgenic mice with an emphasis on the in situ technologies.

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11. Production of Stable Transgenic Mice A. Choice of Expression Vectors and DNA Preparation for Microinjection

An expression vector should contain the appropriate promoter/enhancer sequences, the entire protein-codingregion, and a polyadenylationsignal. Although cDNAs have been used successfully in a number of studies, they are in general poorly expressed. Therefore, including genomic sequences containing introns within the expression vector is highly recommended. If the entire gene is not yet available, minigenes carrying combinations of genomic and cDNA can also be considered. There appears to be no upper limit to the size of microinjected DNA. DNA cloned into cosmids or yeast artificial chromosomes is efficiently incorporated into the mouse genome (Schedl et al., 1993). The DNA to be microinjected should be free of vector DNA sequences. The presence of prokaryotic vector sequencesmay affect expression of the introduced gene. The purity of the DNA is of utmost importance. Care must be taken to remove organic solvents and salt, which are highly toxic to fertilized eggs. We routinely isolate genomic fragments by electroelution followed by pwification of the eluted fragment by elutip-d columns (Schleicher & Schuell, Inc., Keene, New Hampshire), described in detail as follows:

1. Digest plasmid DNA (20-30 pg) with the appropriate restriction enzyme(s). 2. Electrophorese the digested DNA on 0.8-1% agarose gels at 25 volts. 3. Cut the gel fragment under UV illumination and place it in a dialysis tube. 4. Electroelute the DNA at 80 V for 1-2 hr. 5. Reverse the current for 30-60 sec. 6. Adjust to the low salt solution (0.2 M NaC1, 20 mM Tris-HC1 pH 7.5, 1 mM EDTA) and slowly pass it through an elutip-d column, preequilibrated with the same buffer. 7. Wash the column with 2-3 ml of the low-salt buffer, and dry it completely. 8. Elute the DNA with 0.4ml of the high-salt solution into a sterile Eppendorf tube and add 2 volumes of ethanol. Store overnight at -20°C. 9. Dissolve the pellet in TE (10 mM Tris-HC1, pH 7.5, 1 mM EDTA). 10. Determine the DNA concentration either by fluorometer or by running an alliquot in an analytical agarose gel with known amounts of DNA molecular markers. 11. Dilute the DNA to the desired concentration (for most purposes 1-3 pg/ ml of a 5-kb DNA fragment) with the injection buffer (10 mM Tris-HC1pH 7.5, 0.1 mM EDTA). It can be stored at -20°C. Most importantly, each DNA preparation should be checked for toxicity to fertilized eggs. We test the ability of each DNA preparation to allow development of microinjected eggs to the two-cell stage. If a large proportion of microinjected

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eggs are blocked at the one-cell stage, or show signs of morphological deterioration, the DNA solution is diluted twofold or more until about 80% of the eggs reach the two-cell stage. B. Superovulation 1. Mouse Strains Most mouse inbred strains reproduce poorly and the efficiency of gene integration is considerably lower than F1 or F2 hybrid strains. F2 hybrid zygotes generated by breeding C57BI/6J X SJL F1 mice have been most extensively studied. Other working combinations include C57BI/6J X C3H, C57BU6J X CBA, or BalblC X C57BU6J. We use the inbred mouse strain FVB/N, established at the NIH, as a donor strain for fertilized eggs (Takeo et aL, 1991). This mouse strain is characterized by vigorous reproductive performance and consistently large litters. In addition, fertilized FVBNeggs contain large and prominent pronuclei, which facilitate microinjection.

2. Mouse Colony

An access to an adequate animal care facilities is essential for the production and maintenance of transgenic mice. Mice should be housed under controlled conditions of temperature, humidity, and light cycle. The health of the colony must be monitored routinely, and serum samples should be tested for the presence of antigens of infectious viruses. Maintaining mice in pathogen-free conditions is advantageous. However, it is also extremely expensive. 3. Hormone Treatment Our mouse colony is under a 12-hr lighddark cycle. The end of the light cycle is at 6 P.M. and the end of the dark cycle is at 6 A.M. Superovulation is initiated by injecting 5 i.u. pregnant mare serum (PMS) (Sigma, St. Louis, Missouri, or Gestyl from Organon, or Folligon from Intervet Laboratories) between 2 and 3 P.M. followed by injecting 5 i.u. human chorionic gonadotropin (hCG) (Sigma) two days later between 12 and 1 P.M. Females injected with hCG are placed individually with stud males. Vaginal plugs are checked on the following morning. C. Media Preparation

We use CZB (Chatot et aL, 1989)and HEPES-CZB media for flushing, microinjection, and culturing of one-cell mouse embryos to the two-cell stage (Tables I and 11). Glutamine is added to the medium from a fresh 100-mM glutamine stock immediately before use. Media are prepared in disposable polystyrene tissue-culture tubes, using endotoxin-free, tissue-culture grade water (Sigma

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Table I Recipe for 100 ml 1OX Stock Solutions Stock A

Stock B Stock C Stock D Stock E

4.110 g NaCl 0.356 g KCI 0.293 g MgS04 . 7H20 0.161 g KH2PO4 6.180 g 60% sodium lactate 0.040 g EDTA 0.060 g penicillin 0.050 g streptomycin 2.100 g NaHCOs 0.010 g phenol red 0.360 sodium pyruvate 2.520 g CaC12 . 3H20 5.958 g Hepes 0.010 g phenol red

Chemical Co. No. W3500).Media are filter-sterilized through 0.22-pm Millipore filters in disposable filter holders, gassed with 5% COz, and stored at 4°C. We prepare fresh media every 2 weeks. The osmolarity of all media are tested by freezing-point depression and ranged from 274 to 295 mosmol. D. Embryo Recovery

Fertilized mouse embryos are recovered from the oviducts of plugged females as follows: 1. Sacrifice mice by cervical dislocation. Dissect the oviduct free of the ovary and uterus and place in a small Petri dish containing warm CZB medium. Bring an oviduct into a depression glass slide containing HEPES-CZB and 0.3 mg/ml hyaluronic acid under the dissecting microscope. Table I1 Preparation of 10 ml CZB and HEPES-CZB Stock A

B C D E Glutamine (200 mM) HZO BSA (Sigma A4378)

CZB

Hepes-CZB

1 ml 1 ml 0.1 ml 0.1 ml 0.05 ml 1.15 ml

lml 0.1 ml 0.1 ml 0.1 ml 0.9 ml 0.05 ml 1.15 ml 5.0 g

5.0 g

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2. The fertilized eggs are located at the ampule, which because of superovulation is expanded and transparent. Through its thin wall, masses of fertilized eggs surrounded by cumulus cells can easily be observed. These masses are released from the ampule to the medium by tearing the thin wall with watchmaker’s forceps. 3. The cumuluscells are dispersed in 3-5 min because of the enzymatic activity, and the eggs are taken in a small volume and washed twice in 2 ml CZB medium. Fertilized eggs can be distinguished from unfertilized eggs by their prominent second polar body. E. Preparation of Holding and Microinjection Pipets

1 . Holding Pipet Holding pipettes are made of 5 - ~ Drummond 1 microdispenser tubes, Cat No. 105G, outside diameter 1.0 mm, as follows: a. Tubes are drawn by hand over a small Bunsen flame to an outside diameter of about 100 pm. b. The drawn capillary is clamped to the micoforge (e.g., Narashige, MF-79) and a clean break is made by positioning the filament next to the pipet, heating the filament until the glass starts fusing to the filament, cooling the filament for a few seconds, and pulling the tube away from the filament. c. Orient the capillary so that the tip is facing down and is brought close to the filament. The two should be in the same plane and in a sharp focus. The filament is turned on and the glass begins to melt. Stop heating the filament when the internal diameter of the tube is reduced to about 20 pm.

2. Microinjection Pipets We use the thin-wall glass capillaries with internal filament (TWlOOF-4 of World Precision Instruments, Inc.) with an outside diameter of 1.0 mm. There

is no need to clean or pretreat these capillary tubes in any way. In fact, such pretreatments should be avoided.

a. Mount a pipet in the Sutter PC-84 puller. This puller is designed for maximum control and flexibility. The temperature, the force, the distance, and the time for a pull are all programmable with a defined sequence of up to 16 steps. b. Choose an appropriate program following the detailed instruction of the manual. For DNA microinjection, a tip size of 0.3-0.5 pm gives an ideal flow. c. Determine the tip size of the pulled capillaries using the Sutter LW-87 tip calibration device. d. Store pulled pipets for several hours in a dust-free box. e. Load DNA into the microinjection pipet by placing the base of the pipet into the DNA solution. The solution is drawn by capillary action to the tip.

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F. Microinjection: Equipment and Setup

1 . Equipment

We use the fixed-stageNikon Diaphot inverted microscope with long-workingdistance Nomarski objectives and Narashige NT micromanipulators. The micromanipulators are arranged so that the syringe micrometer on the right side, controlling the holding pipet, is connected via polyethylene tubing to the instrument collar on the left micromanipulator. The polyethylene tubing connected to the holding pipet is filled up with Fluorinet (Sigma) and all air bubbles expelled from the system. The micromanipulator on the right side, controlling the microinjection needle, is connected via polyethylene tubing to the picoinjector PLI-100 (Medical Systems Corp.). The pico-injector is a single-channel, pneumatic, digital injector that delivers small liquid volumes precisely by applying a regulated pressure for a set time.

2. Microinjection Chamber Homemade 75 X 38 m m glass chamber slide is used. Several stripes of HepesCZB medium are placed in the middle of the slide and covered with silicone oil. Fertilized eggs (20-30) are positioned at one end of the microdrop under the dissecting microscope. 3. Microinjection a. Place the chamber on the microscope stage. The elongated microdrops should be parallel to the Y axis of the stage and the embryos are at the front of the stage. Bring the embryos into focus with the lower-power objective (X10). b. Insert the holding pipet into the instrument collar, making sure that all air bubbles are expelled. With the micrometer, push the Fluorinet into the pipet until a drop is hanging at the tip. Move the chamber to the rear of the stage and carefully lower the pipet into the microdrop. Make sure that the angle is such that only the tip of the pipette will touch the glass slide. Move the chamber into the front of the stage and suck up an embryo with the micrometer. c. Insert the microinjection pipet into the instrument collar. Set up the picoinjector to deliver a constant positive pressure that prevents back flow and clogging of the pipet. Align the pipet parallel to the holding pipet and slowly lower it into the microdrop. Do not allow the pipet to reach the bottom of the drop, as it might break. Also, do not allow the two pipets to collide. Switch from the X10 to the X 2 5 objective. d. With an embryo firmly held by the holding pipet, bring the male (usually bigger) pronucleus into focus. Bring the tip of the microinjection pipette into a focus, with the fine vertical drive of the manipulator, next to the zona pellucida. Move the pipette into the pronucleus, avoiding the nucleoli, and microinject by

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pressing the foot switch of the pico-injector. If the pronucleus swells, injection was successful (Fig. 1).If no swelling is visible, then either the tip is blocked or the pipette was not inserted into the nucleus. In the former case the pressure can be increased, whereas in the latter, the pipet should be withdrawn and

Fig. 1 Injection into the male pronucleus of mouse fertilizedegg. (A) Orienting the microinjection

pipet to the optical plane of the male (larger) pronucleus. (B) Insertion of the pipet and injection of the DNA solution. (C) Withdrawal of the pipet.

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realigned at the optical plane of the pronucleus prior to the second attempt. Mouse embryos are usually resistant to multiple penetrations if the plasma membrane is not ruptured. e. After a successful injection, withdraw the pipet and move the chamber to the back of the stage. Expel the embryo from the holding pipet by applying pressure to the micrometer and move the chamber to the front of the stage again to suck up the next embryo. f. When all the embryos in that drop have been injected, take the injection chamber to the dissecting microscope, transfer the healthy-looking eggs into a drop of CZB medium under silicone oil, and place them in the incubator. g. Take 20-30 fresh eggs into the next microdrop and repeat the microinjection steps just described.

4. Troubleshooting Microinjection into the pronuclei of fertilized eggs is still endowed with numerous technical problems. The rule of the thumb is that trying to overcome flow problems of a micropipet is better than replacing it. Basically, problems arise in the action of either the injection needle or the holding pipet. a. The needle is blocked. This is the most common problem and is likely due to proteins sticking to the tip during penetration and withdrawal. Apply momentary high pressure with the pico-injector to clear the clogged pipets. Alternatively, the tip of the pipet can be glanced against the holding pipet to increase the size of the opening. If these remedies do not clear the pipet, replace it. b. Aged pipets become very sticky and upon withdrawal, nuclear material is removed as well. Even if such an embryo does not lyse immediately, it will not survive. The pipet should be discarded. c. The pressure of injection can push the nucleoli from the nucleus to the cytoplasm. Such embryos do not survive. d. Avoid the nucleoli! These are very sticky organelles and when the pipette is withdrawn nucleolar material will be dragged along. This will kill the egg and clog the pipet. Pipets clogged with nucleolar material must be replaced. e. The holding pipet, Oil droplets may clog the pipet. In addition, if the tip size is too big, embryos may be sucked up and clog the pipet. G. Vasectomy and Embryo Transfer

The most common practice is to transfer embryos several hours after microinjection into the oviducts of 1-day pseudopregnant mice. However, transferring can be performed after overnight culturing of injected embryos at the two-cell stage to 1-day pseudopregnant females.

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1. Vasectomy Anesthetize an adult male mouse by intraperitonealinjection of 0.4 ml Avertin. Place the mouse on its back and after gently pushing the testes into the abdominal cavity, make a small (about 1-cm) transverse incision just above the preputial gland. With blunt forceps, pull out the testicular fat pad until the testes are exposed. Cut as much of the vas deferens as possible and repeat the procedure with the other testes. Replace the testes into the abdominal cavity and apply two wound clips to the skin. We do not suture the body wall. Vasectomized mice are allowed to recover for about 2 weeks before they are caged individually and mated with CD1 females that serve as foster mothers for microinjected embryos. Avertin is prepared by dissolving 2.5 g 2,2,2-tribromoethanol (Aldrich) in 5 ml ten-amyl alcohol in warm tap water. When it is completely dissolved, add doubledistilled water to a final volume of 200 d.Store at 4°C in the dark.

2. Embryo Transfer Pipet Introduce air bubbles into a finely drawn Pasteur pipet and up to 15 embryos in a minimum volume of CZB medium.

3. Embryo Transfer Anesthetize a pseudopregnant CD1 mouse by intraperitoneal injection of Avertin. Place the mouse on the stage of a dissecting binocular (e.g., Zeiss SV8) and make a longitudinal incision at back midline at the level of the uterus. With blunt forceps pull out the ovarian fat pad along with the ovary, oviduct, and some of the uterus. Clip a serafine to the fat pad and spread it on the skin so that the ovary and oviduct remain outside the body wall. The opening of the infundibulum always faces the tail. Apply a few drops of 0.1% adrenaline and tear the ovarian bursa with two watchmaker forceps. Adrenaline treatment prevents bleeding from the injured vascularized bursa for several minutes. Push the bursa under the ovary and identify the infundibulum in the crevice between the ovary and oviduct. Insert the embryo transfer pipette into the infundibulum, being careful not to tear the delicate tissue. Expel the embryos and air bubbles and slowly remove the pipette. Return the ovary to the abdominal cavity. Repeat the procedure on the other side. Close the skin with a wound clip and keep the mice warm until they recover from the anesthesia. Embryo transfer into the oviduct is considered to be the most difficult step in the production of transgenic mice. Therefore, one should allow some time to practice with nonmanipulated eggs. H. Identification of Transgenic Mice

1. Tyrosinase Gene Marker Screening for positive transgenic mice is labor and time consuming, requiring DNA analyses of tail biopsies (see later discussion) by Southern blot hybridiza-

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tion or PCR. Therefore, the idea that transgenic mice can be identified at birth is very appealing. It was demonstrated that introducing a functional tyrosinase gene into albino mice led to pigmentation of the eye and skin with high penetrance (Beermann et al., 1991). It is also established that coinjection of two independent genes into fertilized mouse eggs results in most cases in cointegration of both genes at the same chromosomal site. Therefore, coinjecting the gene of interest with the tyrosinase gene would lead to cointegration and facilitate identification of transgenic animals over the albino genotype. However, caution should be excercised in adopting this approach, since we and others could not reproduce these results.

2. Tail DNA Analyses This is the traditional method for screening transgenic animals. a. Anesthetize 3-week-old mice with 0.3-0.4 ml of Avertin and cut about 0.5 cm of the tail into a microcentrifuge tubes. b. Digest the tail biopsy for several hours in 0.5 ml solution containing 100 mM EDTA, 10 mM NaCl, 10 mM Tris-HC1, pH 8.0,1% SDS, and 200 pg/ ml proteinase K at 55°C in a shaking water bath. c. Extract the digest with phenolkhloroforndisoamyl alcohol followed by extraction with chloroform/isoamyl alcohol. d. Adjust to 2.5M ammonium acetate and precipitate the DNA with 1volume of cold isopropanol by spooling on a glass rod. e. Wash the DNA with 70% ethanol and dissolve in 50 pl of 10 mM TrisHCl, pH 8.0,l m M EDTA (or 0.1 mM EDTA if PCR is used for the screening). This method yields DNA that is usually clean enough for dot blots, restriction digests, and Southern blots, as well as for PCR. Although isolating DNA from a large number of mice is laborious and expensive, it is highly reliable. There are several protocols for quicker DNA isolation that avoid extractions in organic solutions. However, such protocols are less reliable and might even be misleading. If one must employ these procedures, appropriate positive internal-control primers should be included.

111. Production of Transient Transgenics A. Advantages

Production of stable transgenic strains is time and labor consuming. Usually the analysis of these mice awaits the F1 or F2 generation. In addition,maintaining a large number of strains requires investing substantial resources in adequate animal facilities. However, in many cases, such as for the identification of cisacting control elements conferring a particular pattern of gene expression or for

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the analysis of the consequences of dominant gene expression on embryonic development, a quicker means would be to produce transgenic embryos and analyze them directly instead of establishing stable strains for each integration event. We have used this approach to dissect the regulatory elements of the human and quail myoD genes (Goldhamer et al., 1992,1995; Pinney et al., 1995), as well as to study the consequences of ectopic myoD expression on mouse embryos (Faerman et al., 1993).

IV. Analysis of Expression at the R N A Level A. RNA Isolation

Total tissue RNA is isolated by the urea-LiC1method of Auffray and Rougeon

(1980). All solutions should be sterilized and prepared in DEPC-treated water.

Care must be taken at all steps to prevent RNase activity.

Place a tissue in 5 ml of 6 M ured3 M LiCl in a sterile plastic tube on ice. Add few drops of 1-octanol to prevent foaming. Homogenize for 30 to 60 sec in a Polytron homogenizer at full speed. Leave at 4°C overnight and spin for 30 min at 10,000 rpm in the cold. 4. Dissolve the pellet in 2 ml solution containing 7 M urea, 2% SDS, 0.35 M NaC1,l mM EDTA, 10 mM Tris-HC1, pH 8.0, and 50 pglml heparin. 5. Extract the RNA with phenolkhlorofodisoamyl alcohol and then with chloroform/isoamyl alcohol. 6. Adjust to 2.5 M ammonium acetate and add 2.5 volumes cold ethanol. 7. Pellet the RNA at 10,OOO rpm for 15 min in the cold and dissolve in 100200 pl sterile water. RNA concentration is determined by reading the optical density at 260 nm. Store at -70°C until used. 1. 2. 3. 3.

B. Northern Blots 1. Prepare a 1.5% agarose gel of 10 X 15 X 6 cm dimension by dissolving 1.5 g agarose in 74 ml water. Cool to 60-65°C and add 10 ml of 1OX MOPS buffer (0.2 M MOPS, 50 m M sodium acetate, 10 mM EDTA, pH 7.0) and 16 ml of 37% formaldehyde solution in fume hood. 2. Dissolve a pellet of 20 pg total RNA in 5 pl DEPC-treated water and 19 p1 dye buffer (100 p1 1OX MOPS, 160 pl formaldehyde, 500 p1 formamide, 100 pl glycerol, and 0.1% bromphenol blue). 3. Heat the sample at 65°C for 5 min and immediately quench in ice-water

bath. Add 0.5 pglml ethidum bromide. 4. Run the gel at 10 Vlcmfor about 2-3 hr, until the bromphenol blue migrates about 10 cm.

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5. Soak the gel in 1OX SSC twice for 20 min each with gentle shaking. This removes most of the formaldehyde. 6. Photograph the gel on a short-wave transilluminator. 7. Transfer the RNA in 1OX SSC by capillary action onto Genescreen membrane. 8. Fix the RNA to the membrane by placing the nylon membrane with the RNA facing down on a short-wave transilluminator for 5 min. 9. Prepare 32P-labeledprobes (RNA or DNA) using standard procedures. 10. Prehybridize in 50% formamide, 5X SCC, 5X Denhardt, 50 mM sodium phosphate buffer, pH 6.8, 250 pglml sheared denatured salmon sperm DNA, 100 pg/ml yeast tRNA, and 1%SDS. Prehybridization temperature is determined by the type of probe being used. For RNA probe use 60-65"C, for DNA probe use 50-55°C. 11. Hybridize in 50% formamide, 5X SSC, I X Denhardt, 20 mM sodium phosphate buffer, pH 6.8, 100 pg/ml sheared denatured salmon sperm DNA, 100 pg/ml tRNA, 1%SDS, and 10% dextran sulfate. C. RNase Protection Assay

The RNase protection assay is a highly sensitive assay to detect and quantitate mRNA. 1. Linearize the plasmid DNA on the 3' of the insert with the appropriate restriction enzyme. 2. Synthesize RNA probe using SP6, T3, or T7 RNA polymerase in a reaction mixture containing 32P-UTP(specific activity 800 CUmmol), 0.5 pg linearized plasmid, 500 p M each ATP, GTP, and CTP, 12.2 p M UTP, 30 pCi 32P-UTP, 16 units RNasin, 10 mM NaCl, 40 mM Tris-HC1, pH 7.5, 10 mM DTT, and 10 units of the RNA polymerase. Incubate for 1 hr at 37°C. 3. Remove the template DNA with 20 pg/ml RNase-free DNase in the presence of 10 pg carrier tRNA and 20-40 units RNasin. Incubate for 15 min at 37°C. Stop the reaction with 2% SDS. 4. Add ammonium acetate to 2.5 M and ethanol-precipitate. Dissolve the pellet in 200 pl hybridization buffer (80% deionized formamide, 0.4 M NaCl, Pipes, pH 6.4, and 1 mM EDTA). 5. Dissolve a pellet containing 10 pg total RNA and 10 pg tRNA in hybridization buffer containing about 200,000 cpm of the RNA probe. 6. Heat the sample at 85°C for 5 min and incubate overnight at the appropriate temperature under water. 7. Stop the hybridization by adding 180 pl of a solution containing 0.3 M NaCl, 10 mM Tris-HC1, pH 7.5,5 mM EDTA, 8 pg RNase A, and 0.4 pg RNase T1. Incubate for 1hr at 37°C.

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8. Add 10~ 1 2 0 % SDS and 50 pg proteinase K and incubate at 37°C for 15 min. 9. Add 10 pg carrier tRNA, ammonium acetate to 2.5 M,and 2.5 volumes of ethanol to precipitate the hybrids. 10. Dissolve the pellet in 5 p1 running buffer and electrophorese on 5% sequencing gel. D. RT-PCR

The greatest advantage of RT-PCR for detection and quantitation of specific RNAs is its speed. However, because of its sensitivity, false-positive results caused by contaminationwith minute amounts of DNA are the major restriction. 1. cDNA synthesis with random primers. In a total volume of 40 pl, add 1OX Taq polymerase buffer (500 mM KC1,100 mM Tris-Ha, pH 8.3,15 mM MgClz, 0.1 mg/ml gelatin), 2 pg total RNA, 100 pmol hexamers, 1.25 mM each dNTP, 40 units RNasin, 200 units Moloney murine leukemia virus reverse transcriptase. Incubate for 1 hr at 42°C. 2. PCR in a total volume of 50 pl containing 1OX taq polymerase buffer, 10 pl cDNA, 100 pmol oligonucleotide primers, 5 pl DMSO. Heat at 95°C for 2 min. Add 2 units Taq DNA polymerase. Annealing and elongation temperature and time should be tested for each set of primers. It is recommended to include internal positive control primers for each reaction. Suitable primers could be as follows: L-pyruvate b a s e 3’ S16 ribosomal protein

5’ GGGTCAGTTGAGCCACACTCG3’

5’ AAGCAACGTAGCAGCATGGAA 5‘ AGGAGCGATITGCTGGTGTGGA 3’ 5’ GCTACCAGGCCTITGAGATGGA

The number of amplification cycles depends on the method of detection. If ethidium bromide staining is the ultimate goal, then 30-40 cycles should be performed. If RNA quantitation is required, the reaction should be stopped while still linear and the amount synthesized detected by Southernblot hybridization using an internal oligonucleotide as a probe.

E. I n Sit# Hybridization to Tissue Sections In situ hybridization (ISH) refers to the set of methods combining cytological and histological techniques with methods of nucleic acid hybridization. This combination allows detection of specific nucleotide sequences within cells and tissues, thus providing visualization of genes and/or gene products. The si@cance of ISH cannot be overestimated because of its high sensitivity and resolution, permitting detection of single-copy genes on chromosome preparations or several hundred mRNA molecules in single cells on tissue sections. ISH became

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the method of choice to visualize the spatio-temporal pattern of gene expression during embryogenesis. There are several categories of ISH according to the probe used (DNA, RNA, or oligonucleotide),the method of labeling its detection (radioactivevs. nonradioactive), and the target (DNA vs RNA). Important theoretical as well as practical aspects of ISH technique are comprehensively discussed in several reviews (Wilkinson and Green, 1990; Polak and McGee, 1990; Wilkinson, 1992; Wilkinson and Nieto, 1993; Sassoon and Rosenthal, 1993). Protocols described in this chapter are used routinely in our laboratory to study the expression of muscle regulatory and structural genes in normal and transgenic embryos and can be adopted for other tissue systems. We describe procedures for detecting mRNA on tissue paraffin sections, using 35S- or digoxigenin-labeled riboprobes, and the visualization of gene expression using the whole-mount ISH on postimplantation (8.5-12.5 dpc) mouse embryos.

1. Radioactive in Sit# Hybridization to Tissue Sections a. Tissue Preparation

All steps (tissuepreparation, sectioning, and prehybridization treatments) must be carried out in RNAse-free conditions with sterile solutions and equipment. All reagents used for ISH are kept separately. Fixution. Freshly prepared fixative is made by mixing 4 g paraformaldehyde with 70-80 ml water. Ten microliters of 10 N NaOH is added and the mixture is heated in a fume hood to 70"C, until the powder is dissolved. After cooling, 10 ml of 1OX PBS is added and the volume is adjusted to 100 ml with water. The pH should be 7.4. Fix excised tissues in ice-cold fixative for 16-24 hr. The thickness of tissue blocks must not be more than 3-5 mm. The volume of fixative must be 20-30 times the volume of the tissue sample. Purufin Embeding. Wash fixed tissues in PBS at +4"C (2 X 15 min). Dehydrate through a series of ethanols: 50%, 70%, 95% and three changes of absolute ethanol. Then incubate in chloroform (twice). Impregnate in three changes of molten paraffin wax (e.g., Paraplast) at 60°C. Duration of incubations depends on the size of tissue blocks. For mouse embryos at developmental stages of 9.5-11.5 days, 30 min incubation at each step is sufficient. For later embryonic stages or large pieces of adult tissue, incubations must be prolonged. Processing can be interrupted and the material be stored in 70-100% EtOH at 4°C. Following impregnation, the sample is oriented in the molten paraffin with heated needles and left at room temperature until it solidifies. Paraffin blocks can be stored indefinitely at +4"C. Sectioning. High-quality sections are prerequisite for unequivocal ISH results. Therefore, it is a good practice to select only the best and most informative sections in terms of morphology. This can be done by collecting some sections for histological evaluation and choosing neighboring sections according to the presence and quality of the desirable structures for ISH. Sections are collected

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on slides subbed with 3-aminopropyltriethoxysilane(TESPA) to prevent detachment of sections during the harsh treatments of the ISH procedure. The slides are prepared by washing in hot 10% laboratory detergent in water, rinsing in running hot water and twice in distilled water, baking in an oven at 200-250°C for 2-3 hr, dipping in 2% TESPA in acetone for 10 sec, rinsing twice in acetone and once in DEPC-treated water, and drying at room temperature or at 37°C. Subbed slides can be stored indefinitely in dust-free conditions. Float separate sections or ribbons of serial sections (4-6 pm) on a drop of 2% EtOH in DEPCtreated water on a subbed slide. Place the slide on a slide-warming table heated at 40-45°C. Let them dry overnight at 40-50°C and store in a refrigerator. Under these conditions slides can be maintained over a year. b. Riboprobe Preparation Riboprobes are synthesized by in vitro transcription reaction with SP6, T3, or 'I7 RNA polymerases as described in Section 3 IV.D, using [CZ-~~SIUTP and 1 p10.75 M dithiothreitol (DTT). Importantly, there is no need to hydrolyze the probe. The amount of probe applied is critical. More than 105 cprdpl results in a significant increase in the background, whereas the intensity of the hybridization signal remains essentially the same. For most purposes we use probes at 2 X lo4cprdpl, which results in exposure time that varies from 3 days (for highly abundant RNAs) to 3 weeks (for low-abundance RNAs).

c. Prehybridiration The aim of the prehybridization step is to make the target RNA accessible to the probe. This is done by treating the biological samples with proteinase K. The treatment described next appears to be suitable for different tissues and for various probes. The most important parameter is the temperature of hybridization, which must be determined empirically for each probe. All washing steps can be performed in Coplin jars (glass or plastic) for a small number of slides or in a stainless steel slide rack, for 30-50 slides. Solutions must be preheated prior to treatment. If not stated otherwise, incubations are at room temperature.

1. Bring the slides to room temperature and deparaffinize by 2 X 10-min changes in xylene. 2. Rehydrate sections by passing sequentially through absolute ethanol (2 X 5 min), 95%, 70%, and 50% ethanols (2 min each). 3. Wash in DEPC-treated water (5 min). 4. Incubate in 2X SSC for 30 min at 70°C. 5. Wash in DEPC-treated water (5 min). 6. Treat with proteinase K solution (2 p g / d in 0.2 M Tris-HC1, pH 7.6, 0.1 M EDTA) for 20 min at 37°C. 7. Refm in 4% paraformaldehyde in PBS for 20 min. 8. Treat slides in freshly prepared 0.2% glycine in PBS for 5 min.

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9. Wash in DEPC-treated water for 5 min. 10. Transfer slides into freshly prepared triethanolamine solution (5.53 ml triethanolamine per 0.5 liter DEPC-treated water) and add acetic anhydride (1.25 ml per 0.5 liter). Incubate for 5 min with continuous mixing over a magnetic stirrer. Add the same amount of acetic anhydride and incubate for another 5 min. Wash the slides in DEPC-treated water for 5 min. The purpose of the acetylation step is to decrease the nonspecific binding of the probe to the section. However, in our hands this step did not lead to noticeable improvement of the results. 11. Wash in DEPC-treated water for 5 min. 12. Dehydrate in graded ethanols (2 min each) 50%, 70%, 95%, 100%. 13. Air-dry the slides and proceed to the hybridization step. d. Hybridization 1. Apply the hybridization solution (Table 111) containing the probe onto pretreated sections (approximately 20 p1 per cm2). Do not allow the drop to dry.

2. Cover the drops with precut pieces of Parafilm and displace air bubbles. 3. Arrange slides horizontally (with sections facing up) in a slide box containing filter paper soaked 50% formamide 5x SSC, and seal the box with an electric tape. For a large number of slides we use a stainless steel slide rack placed vertically in a heat-sealable bag. Incubate overnight at the hybridization temperature (50-65°C). e. Posthybridization Washings

To prevent the oxidation of the 35S-labeledprobe, leading to high background, all solutions used prior to RNAse digestion must contain 10 mM D l T or 1%pmercaptoethanol. 1. Place slides in an appropriate container with 5x SSC, 1% P-mercaptoethanol at 65°C. Incubate for 30-60 min until the Parafilm pieces disengage. 2. Wash slides in 2x SSC, 50% formamide, 1%mercaptoethanol at 65°C for 30 min followed by washing in 2x SSC at 37°C 3 X 5 min. Table III

Composition of In Situ Hybridization Buffers 50% formamide 4 X SSC (pH 8.0) 1 X Denhardt’s 0.5 mg/ml herring sperm DNA 0.25 m g / d yeast RNA 10 mM DTT 10%dextran sulfate

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3. Treat the slides with RNAse A (10 pg/ml) in 0.4 M NaCl, 0.01 M Tris-HC1 (pH 7.5),5mM EDTA for 30 min at 37°C. 4. Repeat the high-stringency washing (step 2) followed by 2x SSC washing. 5. Wash in 3 liters of 0 . 1 ~SSC at 37°C for 15 min. 6. Dehydrate rapidly through 50,70,90% ethanols containing 0.3 M N€€,AC and finally by two changes of absolute ethanol. Air dry.

f. Autoradiography

For a rapid assessment of the hybridization,expose slides to P-max film (Amersham) for 12-24 hr. The hybridization signal on the film can give some indication about the abundance of the transcripts and the localization of the expressing cells. To localize the signal more precisely, high-resolution autoradiography using nuclear track emulsion is required. Important theoretical and practical aspects of histoautoradiography are discussed in Rogers (1979). 1. In the darkroom under safelight conditions (e.g., 25-watt lamp with Ilford 920 filter), melt Kodak NTB-2 emulsion gel in a 50-ml graduated plastic tube containing double-distilled water (1 :1) for 1-2 hr at 43°C. 2. Dip a clean blank slide into the emulsion and remove slowly. Repeat this step with blank slides until no air bubbles appear at the surface of the slide. 3. Dip the hybridized slides into the emulsion. The volume of emulsion needed to cover a standard histological slide is 0.3-0.5ml. 4. Withdraw the slides from the emulsion vertically and blot excess emulsion. 5. Dry slides vertically in the dark for 2-3 hr at room temperature. Using a slide dryer significantly accelerates drying. 6. Store dipped slides overnight at room temperature in a light-tight box. 7. Add into each box a bag with silica gel, seal the box with electric tape, and store at +4"C.

To determine the optimal exposure time (several days to several weeks), test developments should be performed. g. Development and Staining

Warm the box to room temperature for 1-2 hr to avoid condensation. Solutions used for developmentmust be freshly prepared and brought to room temperature. The volumes of developer and fixer should be 5-10 ml per slide. 1. Develop slides in Kodak D-19 developer for 3 min in the dark. 2. Rinse briefly in distilled water. 3. Fix in 30% sodium thiosulfate for 5 min. 4. Wash in at least three changes of distilled water (20 min each) and dry.

For cytologicalobservation the sections must be stained. Usually, light nuclear

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staining allows visualization of both cells and silver grains. We routinely use hematoxylin (Mayer’s or Erlich’s) for counterstaining: 1. The slides are incubated in hematoxylin for 10-30 sec, then washed in 2-3 changes of distilled water followed by washing under tap water. 2. When the color of the sections turn from purple to blue, wash again in distilled water and air-dry. 3. To prepare permanent preparations, put dried slides into xylene and then mount with covedlips using DPX (BDH) or other mountant.

The hybridization signal can be visualized and photographed using either bright- or dark-field illumination. With bright-field illumination, silver grains appear as black dots above the section. If the signal is strong, bright-field photography is adequate. However, when the signal is moderate or weak, dark-field illumination should be used. Under these conditions the signal appears as bright shining dots over a black background and no tissue details can be discerned. Therefore, the standard presentation of radioactive in situ hybridization includes a combined black-and-white photograph of the same section taken with brightand dark-field optics (Fig. 2A,B). Some hematoxylin stained tissues shine brightly under dark-field illumination. To overcome this problem, photographs of unstained sections can be taken with phase-contrast optics or the dark-field photographs can be taken after destaining in ethanol. Another way of presenting results is to make a double exposure of bright- and dark-field images of the same frame on a color film.This results in a combined color picture displaying both silver grains and tissue section details.

2. Nonradioactive in Situ Hybridization with Digoxigenin-Labeled Probes Nonradioactive ISH (nrISH) is based on probes labeled with haptens such as biotin, digoxigenin (dig), or fluorescein. Antisense riboprobes are prepared by in vitro transcription using the appropriate modified nucleotides. The protocols for nonradioactive and radioactive ISH are essentially the same. The most significant difference is the detection of the bound probe. With digoxigenin-labeled probes, anti-digoxigenin antibodies conjugated with marker enzyme (horseradish peroxidase or intestinal alkaline phosphatase) are employed for the immunohistochemical detection. There are several advantages of nrISH, the most important of which is the high resolution not achievable with 35S-labeledprobes and autoradiography. In addition, it avoids the problem of handling, storage, and wasting of radioactive materials, and the histochemicaldetection is quicker, simpler, and more controllable than autoradiographic detection of rISH signal. a. Synthesis of Dig-Lubefed Probes Dig-labeled antisense riboprobes are synthesized using a Dig RNA labeling kit (Boehinger-Mannheim) according to the manufacturer’s protocol. Alternatively,

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Fig. 2 In situ hybridizations.Bright-field (A) and dark-field (B) photomicrographsof a transverse section through a somite of 9.0-dpc mouse embryo hybridized to "S-labeled myogenin riboprobe. (C) Photomicrographof aparasagittal section through the cervical somites of 11.5-dpcmouse embryo hybridized to dig-labeled myoD riboprobe. (D) Whole-mount in situ hybridization showing myogenin expression in 9.5-dpc mouse embryo.

one can use the components of the in v i m transcription reaction purchased from other brands in combination with Boehringer's nucleotide mixture containing dig-labeled UTP. The probe can be stored at -70°C for months. The amount of probe synthesized can be quantitated using dig-labeled control RNA (Boehringer-Mannheim) as a standard. Serial dilutions of the probe and

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control RNA are spotted onto a nylon membrane and revealed with anti-dig antibody conjugate according to the manufacturer’s protocol. For most purposes a concentration of 0.2-1.0 pg/ml of probe is needed. However, it is recommended to determine the optimal concentration by running trial hybridizations with serial dilutions of the probe. The composition of the hybridization buffer is the same as for rISH but without DTT. b. Prehybridization and Hybridization Tissue section treatments and hybridization conditions for nrISH do not differ from these described for rISH. Notably, in contrast to rISH, acetylation decreases the background of hybridizations with dig-labeled probes. Perform stages 1-10 of the prehybridization treatment protocol for rISH and hybridize at 50-65°C as described earlier. c. Posthybridization Washings and Immunodetection The slides are washed according to steps 1-6 of the rISH posthybridization protocol. There is no need to add D l T or @-mercaptoethanol to the washing solutions. The imrnunohistochemicaldetection step is performed at room temperature. Avoid drying of the sections at all steps. 1. Wash slides 3 X 5 min each in TBS (0.15 M NaC1,50 mM Tris-HC1, pH 7.4). 2. Apply a blocking solution (50-100 pl TBS with 3% normal goat or sheep serum). Incubate in a humid chamber for 30-60 min. 3. Tip off excess blocking solution and do not wash. 4. Apply alkaline phosphatase-conjugated anti-digoxigenin antibodies (antidigoxigenin-AP, Fabfragments, Boehringer-Mannheim) diluted 1:500 in TBS with 1%normal goat or sheep serum and incubate for 2 hr. If background staining (due to nonspecific ineraction of the conjugate with the section) appears, the conjugate can be absorbed with heat-inactivated acetone powder prepared from liver of the species used as the source of material for hybridization (Table IV). 5. Wash 3 X 10 min in TBS. 6. Wash 2 X 5 min each in alkaline phosphatase buffer (APB - 100 mM TrisHC1, pH 9.5, 150 mM NaCl, 25 mM mgC12). 7. Reveal alkaline phosphatase activity by incubating sections in the dark with a freshly prepared staining solution. To prepare staining solution, dissolve 5 mg levamisole in 10 ml APB and add 50 pl stock solution of 5-bromo-4-chloro-3indolyl phosphate, 4-toluidine salt (BICP), and 37.5 pl stock solution of nitro blue tetrasolium chloride (NBT).

BCIP stock solution: 50 m g h l in dimethylformamide. NBT stock solution: 100 mg/ml in 70% dimethylformamide. Stock solutions are stable for several months at -20°C in the dark. Ready-for-use stock solutions can be purchased from Boehringer-Mannheim.

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Table IV Preparation of Liver Acetone Powder ~~~

~~

1. Homogenize liver in a minimal volume of PBS on ice. 2. Add 4 volumes of cold acetone and mix well. Keep on ice for 30 min with occasional mixing. Centrifuge at l0,oaOg for 10 min and discard the supernatant. 3. Resuspend the pellet with cold acetone and mix vigorously. Keep on ice for 10 min. Repeat centrifugation. 4. Air-dry the pellet at room temperature on a clean piece of filter paper and store at 4°C in an airtight container. To prepare 1 ml of absorbed antibodies (1:500): 1. To an Eppendorf tube containing 2 mg acetone liver powder add 0.5 ml TBS and heat at 70°C for 30 min. Cool on ice. 2. Add 2 pl anti-digoxigenin antibodies alkaline phosphatase conjugate and incubate with shaking on ice for 1hr. 3. Centrifuge at 10,OOOg for 15 min at +4"C. To the supernatant add 10 p1 normal goat or sheep serum and 0.5 ml TBS.

The appearance of staining varies significantly depending on the abundance of the mRNA. With abundant mRNAs, staining appears after 15-30 min (and up to several hours) of incubation at room temperature. Longer incubations (from 12-16 hr to several days) must be performed at +4"C because of higher stability of the substrate solution at low temperature. Staining is monitored by periodic microscopic observations. The reaction is stopped by washing in TE (50 mM Tris-HC1,l mM EDTA, pH 8.0). The slides are rinsed in distilled water and counterstained with methyl green. Cover the sections with water soluble mountant (e.g., glycerol-gelatin or GVA mountant, Zymed). An example of an nrISH result is presented in Fig. 2C. As an alternative to alkaline phosphatase, horseradish peroxidase conjugates can be used. However, this alternative is recommended only for abundant mRNAs, because of the instability of the peroxidase substrate solutions and the inhibition of enzyme activity by the color product. Nevertheless, for double ISH, employing both 35S- and dig-labeled probes, peroxidase detection is preferable because the product of the alkaline phosphatase reaction interfereswith autoradiography. Another advantage of the peroxidase reaction product is its insolubility in xylene, allowing the use of xylene-based mountants (e.g., DPX, Permount) with superior optical properties. For the peroxidase detection protocol a slight modification must be introduced. After deparaffinization in xylene, sections are incubated for 30 min in absolute methanol with 0.3% H202to inhibit endogenous peroxidase activity. The sections are then transferred into absolute ethanol and processed according to the nrISH protocol as described earlier. The immunodetection is performed as follows: 1. Wash slides in 0 . 1 ~ SSC followed by 3 X 5 min rinses in PBS. 2. Incubate for 45 min in PBS with 3% of normal goat or sheep serum.

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3. Tip off excess fluid and apply anti-digoxigenin antibodies conjugated to peroxidase (anti-digoxigenin-POD, Fabfragments, Boehringer-Mannheim) diluted 1:100 and preabsorbed (if needed) with the appropriate acetone liver powder, and incubate in a humid chamber for 1 hr. 4. Wash in PBS (3 X 10 min). 5. Incubate with freshly prepared substrate solution: 0.15 M NaC1,O.l M TrisHCl, 0.01 M imidazole, pH 7.6,0.5 mg/ml3.3-diaminobenzidinetetrahydrochloride, and 0.01% Hz02. Staining develops within several minutes and incubation longer than 30 min does not result in the intensification of staining.

F. Whole-Mount in Situ Hybridization of Mouse Embryos The major advantage of whole-mount ISH ( W I S H ) is the ability to study the spatial pattern of gene expression directly in the embryo, instead of the timeconsuming sectioning, hybridization, and three-dimensional reconstructions. We tested several protocols of W I S H and found the one developed by Barth and Ivarie (1994), for quail embryos, to be relatively simple and highly reproducible for 8.5- to 12.0-dpcstage mouse embryos. W I S H also employs dig-labeled riboprobes. 1. Fixation of Embryos

Dissect embryos free of extraembryonic membranes in PBS and fix overnight in 4% paraformaldehyde in PBS. Wash fixed embryos in 2-3 changes of 70% EtOH and proceed for hybridization or store in 70% EtOH at -20°C (for several months). Using watchmaker’s forceps and tungsten microneedles, open body cavities to allow free flow of solutions; otherwise, entrapped reagents may cause unacceptable high background. This can be performed while embryos are still in paraformaldehyde or in alcohol. With unturned embryos (8.0-8.5 dpc stage), just remove the extraembryonic membranes and tear the remains of the amnion, to stretch the embryo. With 9.0-10.0 dpc embryos, we make a sagittal incision in the frontal surface of the forebrain to open the brain cavity. From this embryonic stage, the heart must also be teased. With embryos of 10.5-12.0 dpc stages, additional incisions are made in the forebrain vesicles and the roof of the fourth ventricle must be ruptured. During pre- and posthybridization washings, embryos are kept in a roundbottom polypropylene 10-15 ml tube (e.g., Corning 17 X 100 mrn tube, Cat. No. N-25225). Incubations with probe and antibody solutions are done in 2-ml polypropylene test tubes with screw caps (e.g., Sarstedt Cat. No. 72.693). Use polyethylene Pasteur pipets (e.g., Sterilin, Cat. No. PP89) to remove solutions from tubes and to transfer embryos from tube to tube (for bigger embryos, cut the tip of the pipet). When changing solutions, always leave some liquid in the tube to avoid flattening of embryos.

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For washings, the tubes are placed horizontally on a rocking platform. Washings are performed at room temperature, unless stated otherwise. Hightemperature incubations are performed in a hybridization oven.

2. Hybridization Protocol a. Take embryos from 70% EtOH, transfer to PBT.3 (PBS with 0.3% Triton X-100), and wash, 3 X 5 min. b. Treat with Proteinase K (30 pg/ml in PBT.3) at 37%C. With 8.5-9.0 dpc embryos incubate for 10 min, with 9.5 dpc embryos incubate for 15 min, and with 10.5-12.0 dpc embryos incubate for 30 min. c. Rinse in freshly prepared 0.2% glycine in PBT.3 for 5 min. d. Rinse in PBT.3 3 X 5 min. e. Transfer embryos into 2-ml polyethylene tubes with screw caps and fill each tube with a mixture (1 :1)of PBT with hybridization buffer. Incubate for 10 min. f. Replace with hybridization buffer and prehybridize by rotating the tubes in a hybridization oven for 2-3 hr at 65°C. g. Replace with preheated (70"C, 15 min) hybridization buffer containing the probe (250 ng/ml) and hybridize in a hybridization oven at 65°C for 24 hr. h. Remove the hybridization solution (do not discard the hybridization solution; it can be stored at -20°C and reused at least five additional times) and rinse with hybridization buffer without dextran sulfate: once briefly, once for 10 min, once for 20 min at 65°C in a hybridization oven, and once overnight at 65°C in a hybridization oven. i. Transfer embryos into 15-ml tubes, rinse with shaking in PBT.3 3 X 30 min at room temperature, and proceed to the immunodetection step.

3. Immunodetection All steps of the immunodetection are performed at room temperature on a rocking platform. a. Rinse with PBT.3 containing 2% blocking reagent (Boehringer-Mannheim) for 1hr. The 10% stock solution of the blocking reagent is prepared according to manufacturer's protocol, treated with DEPC, autoclaved,and stored at -20°C. b. Absorb the anti-dig antibodies with heat-inactivated mouse-liver acetone powder. To prepare 4 ml of absorbed conjugate (1:2OOO in PBT.3 with 2% blocking reagent), weigh into an Eppendorf tube 4 mg acetone liver powder, add 0.5 ml PBT.3, and heat at 70°C for 30 min. Cool on ice, add 2 p1 of antidigoxigenin antibodies alkaline phosphatase conjugate (Boehringer-Mannheim), and incubate with shaking on ice for 1 hr. Centrifuge at 10,OOOg for 15 min at +4"C. To the supernatant add 800 p 10% blocking reagent and bring to 4 ml with PBT.3.

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c. Incubate embryos with preabsorbed antibodies for 2.5 hr in 2-ml tubes. d. Rinse briefly with PBT.3 and transfer embryos into 15-ml tubes. Rinse 2 X 5 min with PBT.3. e. Rinse with PBT.5 (PBS with 0.5% Triton X-100) for 3 X 20 min. f. Rinse APB.3 (APB with 0.3% Triton X-100) for 15 min. g. Incubate overnight with APB.3. h. Place embryos in 75 X 10 mm glass test tubes, fill the tubes with APB containing 10% poly(viny1alcohol) (APBPVA), and incubate for 30 min. [APBI PVA is prepared by dissolving poly(viny1 alcohol) (MW 30,000-7O,OOO, Sigma P-8136) in APB in a boiling water bath. i. Replace the APBPVA solution with alkaline phosphatase substrate solution, prepared by adding 4.5 pl NBT and 3.5 p BCIP per ml of APBPVA. Incubate in the dark with periodic inspection under the stereomicroscope until the desired intensity of staining is achieved. The rate of color development varies from 1-2 to 24 hr. To avoid overstaining, overnight incubations can be performed at +4"C. To stop the reaction, wash twice in TE. Hybridized embryos can be stored in TE at +4"C in the dark. Photographs of early embryos (up to 9.5 dpc) are taken with a photomicroscope equipped with low-power (2X-4X) objectives (Fig. 2D). If an embryo is too transparent, the staining on the other side of the body is revealed as well and reduces the quality of the pictures. To make embryos less transparent, dehydrate partially in 70% EtOH. Alternatively, embryos can be fixed in 10% formalin.

V. Analysis of Expression at the Protein Level A. CAT Assay

One of the most popular reporter genes for the characterization of control elements in transgenic mice is the bacterial chloramphenicol acetyl transferase (CAT) gene. The qualitative CAT assay is performed as follows: 1. Homogenize tissue samples in 1-5 m10.25 M Tris-HC1, pH 7.8, using Polytron homogenizer. 2. Heat the homogenate for 10 min at 65°C. Centrifuge, collect the supernatant, and store at -70°C. 3. Determine protein concentration and adjust to 20-100 ug, in a final volume of 60 pl. 4. Prepare a mixture containing 20 p1 20mM acetyl coenzyme A, 0.2 pCi 14C-chloramphenicol,and 5 p1 20 mM Tris-HC1, pH 7.8. Adjust the volume to 40 p1 with H 2 0 and add it to the protein sample.

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5. Incubate at 37°C for 1 hr. Stop the reaction with 1 ml ethyl acetate and vortex vigorously. Centrifuge, collect the supernatant, and dry in a Speedvac. 6. Dissolve in 20 pl ethyl acetate, load on a thin-layer chromatography (TLC) plate, and chromatograph with chlorofomdmethanol (955). Dry the plate and autoradiograph.

The quantitative CAT assay is done according to Sleigh (1986): 1. To the protein extract (54 pl) add 20 pl 8 mM chloramphenicol, 5.7 pl 2.0 M Tris, pH 7.8, and 20 p10.1 pCi 14C-acetylcoenzyme A in 0.5 mM nonradioactive acetyl coenzyme A. 2. Incubate for 1 hr at 37°C. 3. Extract with 200 pl ethyl acetate and remove 160 p1 of the supernatant to a minivial containing 2 ml xylene/xylofluor (9 :1). 4. Reextract the sample with 200 pl ethyl acetate and add 200 p1 of the supernatant to the minivial. 5. Read radioactivity in a scintillation counter, 6. A standard curve is made with a commercial CAT enzyme (P. L. Biochem) at a range of 0.002-0.08 unitslreaction.

B. LacZ Stab

‘g

The bacterial LacZ gene, encoding bacterial /3-galactosidase, provides an ideal in situ visual marker for the analysis of regulatory elements in the developing mouse embryoand to follow the spatiotemporalpattern of lacZ in whole embryos. It is also extensively used as a vector for promoter or enhancer traps. The main advantage of lacZ reporter genes is the high sensitivity of detection at the singlecell level (Fig. 3). 1. Dissect embryos in PBS and transfer them into ice-cold freshly prepared 4% paraformaldehyde in PBS. The volume of fixative must exceed 20-30 times the total volume of fixed material. For better penetration of fixative and staining solutions into internal parts of the body, embryos older than 12-13 dpc must be “opened”: make a longitudinal cut in the midline of the abdomen or cut the embryo into two halves along the body axis. In embryos older than 16 dpc the skin should be removed. 2. Incubate for 2-24 hrs at 4°C. 3. Wash embryos in PBS at room temperature (at least X 3) for 30-60 min. The volume of PBS must exceed 20-30 times the total volume of fixed material. 4. Incubate while rocking in staining solution at 37°C for the first 2-3 hr and then at 30°C overnight. Staining solution contains 0.1% X-gal (a 4% stock solution is prepared in dimethylformamide),2 mM MgC12,0.01% (w/v) sodium desoxycholate, 0.02% (vlv) NP-40, 5 mM K3Fe(CN)6, 5 m M We(CN),, and 10% X 10 PBS. The solution is passed through a 0.22-pm filter prior to staining.

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Fig. 3 Expression of the bacterial lacZ reporter gene. Histochemical detection of lacZ activity in

transgenic mouse embryo (10.5 dpc) carrying the reporter gene driven by the regulatory elements of the quail myoD gene. (A) Whole-mount stained embryo. The staining is concentrated mainly in the somites. (B) A sagittal section through the cervical somites of the same embryo showing staining in myotomal cells.

5. Wash embryos in several changes of tap water at room temperature, until the yellow staining is removed (10 min to several hours, depending on the size of the embryos). 6. Place the embryos in 50% EtOH (15 min to 1hr, depending on the size of the embryo) and finally in 70%EtOH. Stained embryos can be store in 70%EtOH. To make embryos more transparent, further dehydrate by passing them through the following reagents: 70%, 80%,95%, 3 X loo%,1:1mixture of 100% EtOH with isobutyl alcohol, pure isobutyl alcohol, 1:1mixture of isobutyl alcohol with mineral oil, and pure mineral oil. The duration of each step vanes according to the size of the embryo: from 15 min (embryos prior to 9.5 dpc) to 1 hr (at later stages). The procedure can be stopped at any EtOH stage. However, too long contact of stained embryos with isobutyl alcohol should be avoided.

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7. For embedding into paraffin, transfer embryos into melted paraplast at 56-60°C and impregnate in three changes of melted paraplast during 1.5-3 hr. Orient the embryos in the embedding tray, remove the tray from the oven, and let it cool down. 8. Paraffin sections can be counterstained with nuclear fast red and mounted with any permanent mountant. However, to avoid stain dissolution care must be taken to minimize the time of exposure to xylene.

VI. Future Directions It appears that transgenic technology by microinjection into mouse fertilized eggs contributed to a number of questions related to myogenesis. Some of these topics are indicated in the Introduction. However, one direction for improvement is to enlarge the size of introduced DNA into the mouse genome. For example, the control elements of the myogenic regulatory gene myf5, the first of the myoD gene family to be expressed during somite formation and differentiation, are not yet known. This gene is closely linked to the MRF4 on human chromosome 12. Attempts to express genomic myf5 gene in transgenic mice have failed to recapitulate the embryonic pattern of expression (Patapoutian et af., 1993). Perhaps control elements of this gene are located further upstream or downstream. The ability to introduce very large DNA fragments, either cosmids or yeast artificial chromosomes, into the mouse genome would permit addressing this question. Another question of interest is whether each of the myogenic regulatory genes are expressed in distinct cell populations during myotome formation. This can be addressed by studying the consequences of expression of toxigens driven by specificelements of these genes or by specificactivation of toxigenes in specific myogenic cells types using the cre-lox recombination approach (Sauer, 1993). It is assumed that members of the MyoD family are functionally identical. The multiplicity of genes serves to fine-tune their expression during embryonic and adult stages. To address this question, MyoD can be activated by myogenin regulatory elements and vice versa in wild-type or in null mutants. Finally, gene trap is one of the most exciting approaches to identify novel genes involved in the establishmentof pattern formation and specificcell lineages. Although it is a blind test, many interesting patterns leading to the identification of novel genes have been reported (for a review, see Skarnes, 1993). We have recently identified a transgenic mouse strain in which the lacZ reporter gene is driven by putative regulatory DNA sequences of an endogenous gene expressed specifically in myogenic cells that migrate from the ventrolateral edge of the somite to populate the limb buds. The spatio-temporal pattern of expression of this gene is distinct from that of another gene (Pax3) expressed in these cells (Williams and Ordahl, 1994). Other$genesimportant in the specification of the myogenic cell lineages and in patterning skeletal muscles would likely be identified using this approach.

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