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[31] Whole-mount immunohistochemistry
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[31] W h o l e - M o u n t I m m u n o h i s t o c h e m i s t r y By CLAYTUS A. DAVIS Introduction Whole-mount immunohistochemistry is the localization of antigens in unsectioned tissues using specific antibodies. 1,2 It can be successfully applied to whole mouse embryos from zygote to 10.0 days of gestation and to dissected tissues at any stage. The intention of this chapter is to provide enough information for the development of a working protocol; to discuss procedure variations; and to indicate possible solutions for the most common problems. Although the whole-mount technique is similar in outline to immunohistochemistry of sectioned material and shares some of the problems, it has several advantages. Most importantly, it quickly and clearly conveys the spatial relationship between antigen distribution and the rest of the embryo. 3 It also requires less effort and working time to complete and permits many samples to be easily processed in parallel. There are also several disadvantages. The elapsed time, from the addition of primary antibody to getting the results, is longer; cellular level detail is poorer; and it does not work for older embryos. The design of a working procedure and its successful application depend on the particular antigen-antibody combination, so there is no one protocol that will work in all situations. The protocol presented in Table I serves primarily as an example and as a reference point for discussing the individual steps. Whole-Mount Procedures
General Points Whole-mount procedures largely consist of a series of incubations, during which reagents must fully penetrate the sample and react, and l M. Costa, Y. Patel, J. B. Furness, and A. Arimura, Neurosci. Lett. 6, 215 (1977). 2 M. Costa, R. Buffa, and E. L. Solcia, Histochemistry 65, 157 (1980). 3 People who are able to visualize well in three dimensions will be able to reconstruct this relationship from serial section data, but this information is very difficult to convey to anyone else. Several c o m p u t e r programs are available that automate the reconstruction and presentation of the data; however, they currently require an inordinate amount of time and effort.
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TABLE I SAMPLE WHOLE-MOUNT IMMUNOHISTOCHEMISTRY PROTOCOLa Step Dissection
Fixation Pretreatment
Primary antibody incubation Washes Secondary antibody incubation Washes Color development
Clearing
Procedure Dissect out the embryos in cold phosphate-buffered saline (PBS), removing all extraembryonic membranes Fix in methanol-dimethyl sulfoxide (DMSO) (4: 1) overnight at 4 ° Blocking endogenous peroxidase: Transfer to methanol-DMSO-30% H202 (4 : 1 : 1) for 4-5 hr at room temperature Storage: Embryos may then be stored in 100% methanol at -20°; rehydrate embryos through methanol series diluted in water: 50%, 20%, PBS, for 30 min each Blocking background binding: Incubate twice in PBSMT (PBS plus 2% instant skim milk powder and 0.1%, v/v, Triton X-100) for 1 hr each at room temperature Incubate overnight at 4 ° with primary antibody diluted in PBSMT Wash 2 times in PBSMT at 4° and 3 times at room temperature for 1 hr each Incubate overnight at 4° with secondary antibody diluted in PBSMT Repeat the above washes, adding a final 20-min wash in PBT at room temperature Incubate embryos in 0.3 mg/ml DAB (diaminobenzidine, e.g., Sigma, St. Louis, MO, D-5637; possibly carcinogenic) + 0.5% NiC12 in PBT (PBS plus 0.2% bovine serum albumin, e.g., Sigma, A-4378, and 0.1%, v/v, Triton X-100) at room temperature for 20 min; add H202 to 0.03% and incubate at room temperature until color density looks good, usually about 10 min Rinse several times in PBT and dehydrate through methanol series: 30, 50, 80, 100% for 30 min each; embryos may be stored in methanol Incubate embryos in BABB [benzyl alcohol-benzyl benzoate (1 : 2)I-methanol (1 : 1) for 20 min; transfer to BABB
a Throughout the entire protocol, embryos should be gently rocked.
subsequent washes, during which excess reagents must be completely removed. To facilitate both, the embryos should be gently rocked or rolled throughout the procedure. Frequent solution changes increase the chances of damaging or losing embryos. To avoid this, transfer the solution rather than handling the
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embryos. Postimplantation embryos may be processed in 15-ml disposable polypropylene tubes. To transfer, stand the tubes upright for a few minutes to let the embryos sink to the bottom, then gently decant the solution. For small embryos, monitor the process over a light box. Check that no embryos have stuck to the inside of the tube cap. In aqueous solutions without detergent, the embryos tend to stick to the tube walls. The incubation of small postimplantation embryos in primary antibody may be done in 1.5-ml cryovials or Reactivials (Pierce, Rockford, IL) to minimize the amount of reagent used. For preimplantation embryos, perform the incubations and washes in a depression slide or microtiter plate. Remove and add reagents by pipette. Avoid drawing up the embryos, however, since they may stick to the inside of the pipette. Monitor the transfers under a microscope. Before using a mouth pipette, remember that some of the reagents are poisonous. A few embryos will be lost during the course of the procedure. Some of the transfers involve a large change of osmotic potential, for example, between aqueous and organic solutions and between different clearing agents. Even when the embryos are fully permeable, a sudden transfer may degrade the appearance. Take the embryos through a graded series of two or three intermediate mixes (e.g., 80, 50, 20%). Mouse embryos of vastly different sizes may produce acceptable whole mounts. As the size increases, the minimum duration of each step also increases. The times indicated in Table I are usually long enough for 10.0day mouse embryos. Observed background staining is proportional to the size of the specimen. It may determine the upper age limit for which the procedure works. For some reason, background is variable between individual large specimens, and so it may be necessary to process many before a few suitable for good photographs are found. Smaller embryos yield uniform results. Dissection
The goal of dissection is to get the embryos from the uterus to the fixative in good condition and in a reasonably short time. A period of 30 rain should be fine. Dissect in PBS (phosphate-buffered saline, without Ca2+/Mg 2+) and keep the embryos on ice as much as possible. Good dissections, especially for the early stages, require practice, steady hands, a well-adjusted dissecting microscope with a good field depth, and undamaged forceps. The stages between egg and late blastocyst are flushed out of the fallopian tubes and uterus. Between implantation and approximately 6.5 days of gestation, the embryos are extremely difficult to find and dissect
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FIG. 1. Flattening an early neurulation embryo. A dorsal view of an 8.0-day embryo is shown. The covering amnion has been removed and two vertical cuts made in the lateral amnion (indicated by arrows). Once the cuts are made, the embryo will flatten out.
away from the mass of decidual tissue. Stages between 6.5 and 7.5 days are manageable but require practice. Later stages are easier. Hogan et al. 4 give detailed instructions for removing the different stages; however, if you are unfamiliar with the dissections, it is best to get a demonstration from someone knowledgeable. During gastrulation and early neurulation (~6.5-8.0 days), the embryo is U-shaped and is bound in the amnion and yolk sac. Although wholemount staining will work on the entire structure, it may be difficult to photograph the results. The embryo will flatten if, before fixing, the amnion cap is dissected off and two cuts are made in the lateral amnion (Fig. 1). After approximately 9.0 days of gestation, the neural tube closes. The lumen of the tube sometimes nonspecifically binds both primary and secondary antibodies, causing high background. To both reduce this and improve penetrability, the thin roof of the hindbrain may be opened and the embryos left to gently rock in PBS at 4° for 10 rain prior to fixing. Torn edges and crushed tissues often show high background staining. Fixation
A perfect fixative would preserve antigenicity and morphology, while leaving the embryo completely open to the passage of antibody and other reagents. Fixatives fall into two groups: those that work by covalently cross-linking proteins, forming a meshwork (e.g., aldehyde fixatives), and 4 B. Hogan, F. Costantini, and E. Lacey, "Manipulating the Mouse Embryo: A Laboratory Manual." Cold Spring Harbor Laboratory, Cold Spring Harbor, New York, 1986.
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those that precipitate the cellular components (e.g., organic solvents). Sternberger discusses their preparation and relevance to immunohistochemistry in more detail. 5 Recipes for two good starting fixatives, one from each group, are given below. The same fixatives used for sectioned material may be tried with whole-mount procedures, although a fixative that works for one may not work for the other. In particular, glutaraldehyde-containing fixatives may generate an almost impenetrable mesh of tightly cross-linked proteins. 4% Paraformaldehyde in PBS. Add 4 g of paraformaldehyde to 50 ml of water plus 1 drop of 1 N NaOH. Dissolve at 37° with shaking for 1 hr. Adjust the pH to approximately 7 with HC1, add an equal volume of 2 x PBS and then filter through Whatman (Clifton, NJ) No. 1 paper. Store at 4 ° and use within 1 day of preparation. Although less convenient, paraformaldehyde is more likely to preserve antigenicity than any other fixative. Dent's Fixative. Dent's fixative is 80% methanol, 20% dimethyl sulfoxide (DMSO). 6 If antigenicity is preserved, this may give a lower background than aldehyde fixatives, and requires less work. Fix on ice in a volume more than 20-fold greater than the sample volume, for between 1 hr and overnight, depending on the size of the embryos. Transfer the embryos to fresh fixative after the first 15-30 min.
Pretreatment Ideally, the pretreatments leave the embryo in such a condition that antibodies and other reagents can freely diffuse through them and bind to only the correct molecules. Which pretreatments are necessary depends on the fixative and the detection scheme used. Blocking Free Aldehydes. Aldehyde-fixed embryos require an incubation in 0.1 M glycine in PBS for 30-60 min at room temperature to block any free aldehyde groups. Blocking Endogenous Enzyme Activity. The fixation protocol may not abolish all endogenous peroxidase activity. At the risk of reducing antigenicity in some instances, the remaining activity may be eliminated by incubation in either PBS plus 0.3-1.0% H202, or methanol or Dent's fixative plus 0.5-5% H2Oz, for 3-4 hr at room temperature. The methanol treatment is preferable because the aqueous reaction may produce destructive oxygen bubbles within the embryo, especially at higher HzO2 concentrations. Most endogenous phosphatase activities are inhibited by includ5 L. A. Sternberger, " I m m u n o c y t o c h e m i s t r y . " Wiley, New York, 1986. 6 j. A. Dent, A. G. Poison, and M. W. Klymkowsky, Development (Cambridge, UK) 107, 35 (1989).
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ing I-2 mM levamisole prior to and during the incubation with the enzyme substrate. 7 If the embryos are still in methanol after completing these blocking steps, they should be rehydrated into PBS through a series of methanol dilutions in water (e.g., 50%, 20%, PBS, for 30 min each). Blocking Nonspecific Antibody Binding. The final pretreatment, antibody dilutions, and washes are all done in PBS plus 0.1-0.2% detergent plus 1-5% protein. The protein may be bovine serum albumin (BSA) or nonfat milk powder (both used at 1-3%) or nonimmune serum from the animal in which the secondary antibody was raised (5-20%, v/v). Nonimmune serum can be heat-treated for 30 min at 56° to inactivate complement. In our hands, milk powder worked the best. The detergent added is usually nonionic: Nonidet P-40, Triton X-100, or Tween 20. Both the protein and the detergent decrease background staining. The excess protein blankets sites in the embryo that interact nonspecifically with the antibodies. The detergent may weaken nonspecific antibody binding. Which protein and detergent will work best needs to be determined empirically for each antigen-antibody combination.
Antibody Incubations Dilute the primary antibody in the last blocking solution (PBS + protein + detergent) and incubate the embryos for a few hours to overnight, depending on their size. Keep the embryos gently rocking or rolling. Long incubations with a low concentration of antibody at 4° will give lower background staining than short incubations with high concentration of antibody. The success of a whole-mount protocol depends greatly on the characteristics of the primary antibody. Antibodies vary, among other things, in their specificity (how well they recognize the correct antigen to the exclusion of others), their affinity (how stable the antigen-antibody interaction is), and their source (polyclonal antisera or monoclonal). An antibody with poor specificity will generate increased background, which, because of the thickness of the material, will readily obscure specific staining. Because the antigen-antibody complex is noncovalent, and bound antibody is in equilibrium with free antibody, some of the bound antibody will be lost during the washes. In sectioned material, where 30 min of washing is usually sufficient, most of the antibody will remain bound, even if it does not have a high affinity. In the whole-mount procedures, 7 B. A. Ponder and M. M. Wilkins, J. Histochem. Cytochem. 29, 981 (1981).
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especially for large embryos, the wash steps may span most of a day, and there is the possibility that much of the signal may be lost if the antibody has a low affinity. A polyclonal antiserum usually contains antibodies recognizing more than one epitope. Consequently, its apparent affinity (acidity) is often higher than that of a monoclonal antibody, which will recognize only one epitope. There are specific problems associated with using mouse monoclonal antibodies on mouse tissues. Self-tolerance makes the production of monoclonal antibodies against epitopes present in the mouse less likely. For this reason, a mouse monoclonal antibody raised against a conserved non-mouse protein may recognize the protein in a wide variety of species except the mouse, since the mouse immune system may preferentially respond to regions of the injected protein that are different from the endogenous version. Also, embryonic or maternal mouse immunoglobulin G (IgG) present in the embryo will bind to any anti-mouse secondary antibody used and so increase background. To keep the embryos as free of mouse immunoglobulin as possible, dissect away maternal and extraembryonic tissues, layer by layer, in different changes of PBS, and wash them thoroughly in PBS before fixing. Remaining immunoglobulins may be blocked by incubating the embryos with unlabeled secondary antibody and then washing prior to the addition of primary antibody. This problem may be avoided by labeling the primary antibody, although this will reduce the sensitivity. A preferable alternative is to use rat monoclonal antibodies. Anti-rat immunoglobulin secondary antibodies which do not cross-react with mouse immunoglobulin are commercially available. Problems arising from self-tolerance will also be reduced. Polyclonal antisera also have disadvantages. Unlike monoclonal antibodies, they contain a majority of nonspecific antibodies, which may cause background. To remedy this, the antiserum may be either diluted, affinitypurified against the antigen, or preadsorbed. If the specific antibody titer is high, then it may be possible to dilute the antiserum to such an extent that the background due to nonspecific antibodies is negligible. Affinity purification produces the cleanest antibody solution but requires 1 week to perform and a large amount of purified antigen; in addition, antibodies that have a very high affinity for the antigen may not elute well, thus stripping the antiserum of its best antibodies. Preabsorption to an acetone powder of embryonic mouse tissue that does not contain any of the desired antigen may remove nonspecifically binding antibodies. Harlow and Lane describe these techniques. 8 s E. Harlow and D. Lane, "Antibodies: A Laboratory Manual." Cold Spring Harbor Laboratory, Cold Spring Harbor, New York, 1988.
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Washes
Five consecutive washes in the blocking solution following antibody incubations are typical. Keep the embryos gently rocking or rolling at either 4° or room temperature. The duration must be determined empirically, but each wash should not need to be longer than 1 hr. If washes are too short, there will be increased background; if too long, there may be a loss of signal. The length of the washes is dictated by diffusion rates and the intensity of nonspecific antibody binding. In general, the rate of diffusion decreases for larger molecules, for larger embryos, and for the denser tissue meshworks generated by some fixatives (e.g., glutaraldehyde). Improving the blocking steps will most often reduce the background better than greatly increasing the wash times. Detection
The binding of primary antibody may be monitored, and the signal amplified, by a number of different methods. For whole-mount techniques, enzyme-linked and fluorochrome assays are usual. Briefly, the complex of antigen and primary antibody is detected by binding a secondary antibody coupled to either a dye, which is then observed by fluorescence microscopy, or to an enzyme, which generates an insoluble colored reaction product from a soluble substrate. Although the fluorochrome assays take less time and localize antigens more precisely, the enzyme-linked assays are preferable because they are more sensitive, permanent, and easier to photograph. Both horseradish peroxidase 9 (HRP) and alkaline phosphatase l° are used. Of the two, HRP is better, because it is smaller and its diaminobenzidine (DAB) reaction product is extremely stable. (Sodium azide, which is often added to protein solutions to prevent bacterial or fungal growth, must not be used during the procedure, since it is a potent inhibitor of HRP.) A variant technique, the avidin-biotin complex H,12 (ABC) method, also works well. The secondary antibody is tagged with biotin and bound to the primary antibody-antigen complex. This in turn is bound to a complex of avidin or streptavidin 13 and biotinylated peroxidase. Because of the additional amplification step, the technique is reputedly more sensitive (although this has been questioned), 5 permitting background arising 9 p. K. Nakane and G. B. Pierce, Jr., J. Histochem. Cytochem. 14, 929 (1966). 10 A. S. Bulman and E. Heydermann, J. Clin. Pathol. 34, 1349 (1981). ii j. L. Guesdon, T. Ternynck, and S. Avarameas, J. Histochem. Cytochem. 27, 1131 (1979). t2 S.-M. Hsu, L. Raine, and H. Fanger, J. Histochem. Cytochem. 29, 577 (1981). ~3 Streptavidin is preferable because of its greater affinity for biotin.
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from nonspecific antibodies to be reduced by further diluting the primary antiserum. Endogenous biotin may be a source of increased background. It may be blocked in the pretreatment step by first incubating the embryos with unlabeled streptavidin or avidin, then with biotin, followed by washing away the excess before adding the primary antibody. 14 Coupled secondary antibodies and ABC kits are available commercially. Irrespective of the chosen detection scheme, each incubation step should be followed by a thorough set of washes in PBS plus detergent plus protein.
Color Development Enzyme-linked assays require the addition of enzyme substrate. If an HRP detection scheme has been chosen then DAB is the best choice. DAB is a potential carcinogen. Treat all waste DAB with bleach. Following the final wash, incubate the embryos in PBS plus 0.3 mg/ml DAB for 30 min. Add H 2 0 2 to 0.03% and follow the color development under a dissecting microscope. If the development takes longer than 15 min, keep the embryos in the dark to avoid light-catalyzed DAB polymerization. To stop the reaction rinse the embryos three times in PBS plus 0.1% (v/v) Triton X-100. If the signal is strong, the color development may be the shortest step in the entire procedure and the HzO z may not completely permeate the embryo before the formation of background staining prompts termination of the reaction. To avoid this, try altering the antibody or reagent concentrations such that the color reaction takes 30 min or longer in the large embryos. The DAB color reaction can be enhanced by including 0.5% NiCI2 in the PBS plus DAB solution. On adding the nickel, a precipitate occasionally forms, which may be removed by filtration through Whatman No. 1 paper.
Clearing To see the staining pattern clearly, the embryos can be made more transparent or more opaque by transfer into solutions of different refractive indices. They are increasingly transparent in the following: methanol < PBS < glycerol < BABB. BABB (Murray's solution), is a mix of benzyl alcohol and benzyl benzoate (1 : 2). It is an obnoxious solution that dissolves polystyrene, but it works very well. Because BABB is nonmiscible with aqueous solutions, embryos should first be dehydrated 14 G. S. Wood and R. Warnke, J. Histochem. Cytochem. 29, 1196 (1981).
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through an alcohol series (20, 50, 100%) and then soaked in alcohol-BABB (1 : 1) before being transferred into BABB. To illustrate the relationship between the staining and the embryo, greater transparency is not necessarily better. Large embryos will need more clearing, but staining in small embryos or near the surface may be best observed if the tissues are more opaque. Background is also less apparent in more opaque embryos.
Microscopy and Photography The embryos are usually photographed submerged in an open dish of clearing medium such that they can be manipulated with forceps into different orientations. Whole mounts are more difficult to photograph than slides. Normal microscopes used for viewing slides are not suitable, since they usually have a shallow field depth. A good dissecting microscope with a camera or one of the microscopes designed for lower magnifications [e.g., Leitz (Wetzlar, Germany) Apozoom, Photomakroskop M400; M10] is preferable. Use a stable viewing platform to avoid vibrations. If the illumination starts to heat the mounting medium, then the resulting convection currents may degrade the image quality by moving the embryo or causing diffraction differences in the medium. Vary lighting conditions to get the best pictures (Fig. 2). Try reflected or dark-field illumination on more opaque embryos when the relation between the staining and the surface of the embryo is to be emphasized. Sometimes a combination of lighting works well. For more transparent embryos, transillumination is best. To photograph deep staining, large embryos must exhibit low background and be made almost completely transparent. Unfortunately, this results in a low contrast between the embryo and the clearing medium in which it sits (Fig. 2B). Staining is usually more obvious in color prints than in black and white, since the eye uses color cues. As journals typically prefer black-and-white figures, try an enzyme-linked detection assay that gives a dark-colored reaction product (e.g., nickel-enhanced DAB staining) for making black-andwhite prints.
Whole-Mount Immunohistochemistry and Scanning Electron Microscopy Scanning electron microscopy (SEM) is the best technique for observing surface detail and morphology. Surface antigens have been detected during SEM by using secondary antibodies coupled to a morphologically
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FIG. 2. Whole-mount immunohistochemistry on different stage embryos. All embryos were processed according to the example protocol given in Table I, using an affinity-purified polyclonal antiserum against the mouse engrailed proteins as primary antibody and a commercial affinity-purified goat anti-rabbit IgG coupled to HRP (Jackson Immunoresearch, West Grove, PA) as secondary antibody, and photographed using a Leitz macroscope and Ektachrome (Kodak, Rochester, NY) color slide film. Anterior is to the right. (A) Dorsal views of early 8.0-day embryos showing staining in the midbrain/hindbrain neuroepithelium. (Left) The DAB reaction was enhanced with NiC12. The embryo was photographed in methanol using reflected illumination. (Right) The DAB reaction was not enhanced. The embryo was cleared in BABB and transilluminated. (B) Lateral view of a 10.0-day embryo showing expression in the midbrain/hindbrain region, the somites, the ventral limb buds, and in the ventral spinal cord and hindbrain (arrows). The DAB reaction was not enhanced. The embryo was cleared in BABB and transilluminated. Bars: 0.3 mm.
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distinct marker 15 or by a detecting secondary electrons emitted from osmium deposited preferentially at the site of DAB polymerization. 16 We have reported a different method that may have some advantages. 17 Nickel ions, added to enhance the HRP/DAB reaction, are incorporated into the growing DAB polymer network. If the nickel is on or near the surface, it is detected during SEM by monitoring for element-specific Xray emissions. Machines that can do this are typically found in metallurgy or geology departments. Although untried, it should be possible to localize several different antigens simultaneously on the SEM image by doing serial whole-mount reactions, incorporating different metals (e.g., nickel and cobalt) at each step. There are some practical considerations to this technique. First, the nickel deposition must be on or within 1 /zm of the surface, since the electron beam does not penetrate far. Second, the embryo surface must be conductive. To accomplish this, coat the surface with as thin a layer of carbon as possible. A thicker coat, or a metal coat, will greatly reduce the sensitivity. Third, the procedure is slow, requiring a few hours to build up an adequate map. Increasing the beam intensity helps. Fourth, nickel emits characteristic soft K a X-rays at 7.427 keV, which may be partially absorbed by the bulk of the embryo. Ensure that the detector is capable of surveying the same region of the embryo as the electron beam.
Storage Embryos may be stored after fixation, after the first blocking steps (e.g., after aldehyde blocking and/or endogenous peroxidase inactivation), and after staining is complete. Embryos in most organic solvents may be safely left for a few days at 4° and for a few months at - 2 0 ° if first transferred into methanol. Aldehyde-fixed embryos may be stored in methanol as well if antigenicity is retained. If not, it may be possible to freeze and store the fixed embryos in liquid nitrogen without severely damaging the morphology. Once stained, the embryos are best stored at - 2 0 ° in methanol. The DAB reaction product is completely stable in methanol and is stable for at least a few days in BABB, but we have noticed variable loss of staining in embryos stored in BABB for several months at 4°. 15 R. S. Molday, in "Techniques in Immunochemistry" (G. R. Bullock and P. Petrusz, eds.), Vol. 2, p. 117. Academic Press, San Diego, 1983. 16 A. L. Hartman and P. K. Nakane, in "Techniques in Immunochemistry" (G. R. Bullock and P. Petrusz, eds.), Vol. 2, p. 103. Academic Press, San Diego, 1983. 17 C. A. Davis, D. P. Holmyard, K. J. Millen, and A. L. Joyner, Development (Cambridge, UK) 111, 287 (1990).
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TABLE II TROUBLESHOOTINGGUIDE Problem
Possible causes
Relevant section of text
Poor morphology
Damageduring dissection Damage during incubations or transfer Poorly fixed embryos Embryos poorly penetrable Inadequate blocking Nonspecific antibody b i n d i n g Wash times too short Wash volumes too small Concentration of antibody or other detection reagent too high Enzyme exposed to substrate for too long Excess enhancement Antigenicity not retained Rare antigen Poor primary antibody
Dissection GeneralPoints Fixation Fixation; Pretreatment; Dissection Pretreatment Pretreatment; Antibody Incubations Washes Washes AntibodyIncubations; Detection
Background
Poor signal
Variable results
Antibody or other detection reagents too dilute Embryos poorly penetrable One of reagents is bad Deep interior staining faint in large embryos Variable background in large embryos
Color Development Color Development Fixation Detection Antibody Incubations; Developing Working Protocol Antibody Incubations; Detection; Developing Working Protocol Fixation; Pretreatment; Dissection (Include positive controls) Dissection; General Points; Color Development General Points
Sectioning For a more detailed examination at the tissue level, embryos stained with D A B can be sectioned. The DAB reaction product is stable in xylene and wax, so follow normal paraffin wax embedding and sectioning procedures. Both staining and b a c k g r o u n d will appear much fainter in the sections than in the whole mounts. Therefore, if the e m b r y o s are to be sectioned, it m a y be desirable to let the color development reaction proceed until the b a c k g r o u n d starts to look quite bad. Developing Working Protocol The whole-mount procedures have a large number of parameters. F a c e d with an untried a n t i b o d y - a n t i g e n combination, the task is to find the set of conditions that give the best results. There are a few practical points that can make this less arduous. F o r the initial tests of a new antibody, use the smallest e m b r y o s that express the antigen well, since they will exhibit cleaner whole-mount
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results. Try the HRP indirect immunohistochemical detection method, since it is reasonably sensitive and straightforward. Fix some embryos in 4% paraformaldehyde and some in Dent's fixative. For each group test 7-8 different serial dilutions of the primary antibody. Working dilutions are typically between 1/20 and 1/1000 for polyclonal antisera or monoclonal antibodies from ascites, and between undiluted and 1/20 for hybridoma culture supernatants. If possible, include another primary antibody known to produce good whole mounts as a positive control, and use it to find a dilution range for the secondary HRP-coupled antibody (typically between 1/200 and 1/1000). Start with overnight antibody incubations at 4 °, and five 1-hr washes. Do not initially enhance the H R P - D A B reaction, since even a small amount of enhanced background may conceal specific staining. Following the initial trials, use the troubleshooting guide to help choose modifications to the procedure. It would take months of work to explore all the possible variations of the whole-mount procedure. If there is no staining or if there is unacceptable background after the first few trials of a new antigen-antibody combination, then it may be worthwhile to start raising more antibodes. Meanwhile, further alterations of the procedure can be tried.
Troubleshooting Table II lists the most common difficulties that may be encountered during the whole-mount procedure, their possible causes, and portions of the text relevant to their solution.
Controls
Artifactual staining is common in immunohistochemical techniques. The same controls are required in whole-mount procedures as in others (e.g., no antibodies, no primary antibody, primary antibody preadsorbed to the antigen). They are fully discussed elsewhere.18 False-negative results are more problematic. They may be due to an insufficiently sensitive procedure, masking or loss of epitopes, or insufficient penetration of the specimen by the antibodies or detection reagents. The last is especially applicable to whole-mount protocols, where the increased thickness and possibly more impermeable basement membranes of older embryos may make exposure of the entire sample t8 Chr. W. Pool, R. M. Buijs, D. F. Swaab, G. J. Boer, and F. W. Van Leeuwen, in "Immunohistochemistry" (A. C. Cuello, ed.), p. 1. Wiley, Chichester, UK, 1983.
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less likely. Finally, no set of controls is perfect, so consistent results using other techniques (e.g., Western blot or RNA in situ hybridization) are desirable. Acknowledgments I t h a n k Drs. I. Gitelman, A. L. Joyner, and N. Bonini for helpful c o m m e n t s .
[32] T e c h n i q u e s for L o c a l i z a t i o n o f S p e c i f i c M o l e c u l e s in Oocytes and Embryos .... By CALVIN SIMERLY and GERALD SCHATTEN
Introduction The fields of cell, developmental, molecular, and reproductive biology and genetics have tremendously benefited from studies on eggs at fertilization. 1'2 These early studies relied on systems like frogs and sea urchins in which fertilization in vitro and embryo culture were performed using simple solutions like seawater or pond water at room temperature. The design of methods for routinely and reliably obtaining excellent in vitro fertilization of many mammals now permits detailed experimentation on molecular and structural features of development in mammals. These investigations have led to many important and unexpected basic discoveries including genomic imprinting3-5; gametic recognition involving unique receptors and galactosyltransferases 6'7 atypical maternal inheritance patterns of the centrosome in mice8; both paternal and maternal inheritance
J T. Boveri, "Zellen-Studien: U e b e r die N a t u r der C e n t r o s o m e n , " Vol. 4. Fisher, Jena, G e r m a n y , 1901. 2 D. Mazia, Exp. Cell Res. 153, 1 (1984). 3 j. M c G r a t h a n d D. Solter, Cell (Cambridge, Mass.) 37~ 179 (1984). 4 M. A. H. Surani, S. C. Barton, and M. L. Norris, Cell (Cambridge, Mass.) 45, 127 (1986). 5 C. Sapienza, A. C. Peterson, J. R o s s a n t , and R. Bailing, Nature (London) 328, 251
(1987). 6 p. M. W a s s a r m a n , Annu. Rev. Cell Biol. 3, 109 (1987). 7 B. D. Shur, in " T h e Molecular Biology of Fertilization" (H. Schatten and G. Schatten, eds.), Cell Biology Ser., p. 38. A c a d e m i c Press, San Diego, 1989. 8 G. Schatten, C. Simerly, and H. Schatten, Proc. Natl. Acad. Sci. U.S.A. 88, 6785 (1991).
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[31] W h o l e - M o u n t I m m u n o h i s t o c h e m i s t r y By CLAYTUS A. DAVIS Introduction Whole-mount immunohistochemistry is the localization of antigens in unsectioned tissues using specific antibodies. 1,2 It can be successfully applied to whole mouse embryos from zygote to 10.0 days of gestation and to dissected tissues at any stage. The intention of this chapter is to provide enough information for the development of a working protocol; to discuss procedure variations; and to indicate possible solutions for the most common problems. Although the whole-mount technique is similar in outline to immunohistochemistry of sectioned material and shares some of the problems, it has several advantages. Most importantly, it quickly and clearly conveys the spatial relationship between antigen distribution and the rest of the embryo. 3 It also requires less effort and working time to complete and permits many samples to be easily processed in parallel. There are also several disadvantages. The elapsed time, from the addition of primary antibody to getting the results, is longer; cellular level detail is poorer; and it does not work for older embryos. The design of a working procedure and its successful application depend on the particular antigen-antibody combination, so there is no one protocol that will work in all situations. The protocol presented in Table I serves primarily as an example and as a reference point for discussing the individual steps. Whole-Mount Procedures
General Points Whole-mount procedures largely consist of a series of incubations, during which reagents must fully penetrate the sample and react, and l M. Costa, Y. Patel, J. B. Furness, and A. Arimura, Neurosci. Lett. 6, 215 (1977). 2 M. Costa, R. Buffa, and E. L. Solcia, Histochemistry 65, 157 (1980). 3 People who are able to visualize well in three dimensions will be able to reconstruct this relationship from serial section data, but this information is very difficult to convey to anyone else. Several c o m p u t e r programs are available that automate the reconstruction and presentation of the data; however, they currently require an inordinate amount of time and effort.
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TABLE I SAMPLE WHOLE-MOUNT IMMUNOHISTOCHEMISTRY PROTOCOLa Step Dissection
Fixation Pretreatment
Primary antibody incubation Washes Secondary antibody incubation Washes Color development
Clearing
Procedure Dissect out the embryos in cold phosphate-buffered saline (PBS), removing all extraembryonic membranes Fix in methanol-dimethyl sulfoxide (DMSO) (4: 1) overnight at 4 ° Blocking endogenous peroxidase: Transfer to methanol-DMSO-30% H202 (4 : 1 : 1) for 4-5 hr at room temperature Storage: Embryos may then be stored in 100% methanol at -20°; rehydrate embryos through methanol series diluted in water: 50%, 20%, PBS, for 30 min each Blocking background binding: Incubate twice in PBSMT (PBS plus 2% instant skim milk powder and 0.1%, v/v, Triton X-100) for 1 hr each at room temperature Incubate overnight at 4 ° with primary antibody diluted in PBSMT Wash 2 times in PBSMT at 4° and 3 times at room temperature for 1 hr each Incubate overnight at 4° with secondary antibody diluted in PBSMT Repeat the above washes, adding a final 20-min wash in PBT at room temperature Incubate embryos in 0.3 mg/ml DAB (diaminobenzidine, e.g., Sigma, St. Louis, MO, D-5637; possibly carcinogenic) + 0.5% NiC12 in PBT (PBS plus 0.2% bovine serum albumin, e.g., Sigma, A-4378, and 0.1%, v/v, Triton X-100) at room temperature for 20 min; add H202 to 0.03% and incubate at room temperature until color density looks good, usually about 10 min Rinse several times in PBT and dehydrate through methanol series: 30, 50, 80, 100% for 30 min each; embryos may be stored in methanol Incubate embryos in BABB [benzyl alcohol-benzyl benzoate (1 : 2)I-methanol (1 : 1) for 20 min; transfer to BABB
a Throughout the entire protocol, embryos should be gently rocked.
subsequent washes, during which excess reagents must be completely removed. To facilitate both, the embryos should be gently rocked or rolled throughout the procedure. Frequent solution changes increase the chances of damaging or losing embryos. To avoid this, transfer the solution rather than handling the
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embryos. Postimplantation embryos may be processed in 15-ml disposable polypropylene tubes. To transfer, stand the tubes upright for a few minutes to let the embryos sink to the bottom, then gently decant the solution. For small embryos, monitor the process over a light box. Check that no embryos have stuck to the inside of the tube cap. In aqueous solutions without detergent, the embryos tend to stick to the tube walls. The incubation of small postimplantation embryos in primary antibody may be done in 1.5-ml cryovials or Reactivials (Pierce, Rockford, IL) to minimize the amount of reagent used. For preimplantation embryos, perform the incubations and washes in a depression slide or microtiter plate. Remove and add reagents by pipette. Avoid drawing up the embryos, however, since they may stick to the inside of the pipette. Monitor the transfers under a microscope. Before using a mouth pipette, remember that some of the reagents are poisonous. A few embryos will be lost during the course of the procedure. Some of the transfers involve a large change of osmotic potential, for example, between aqueous and organic solutions and between different clearing agents. Even when the embryos are fully permeable, a sudden transfer may degrade the appearance. Take the embryos through a graded series of two or three intermediate mixes (e.g., 80, 50, 20%). Mouse embryos of vastly different sizes may produce acceptable whole mounts. As the size increases, the minimum duration of each step also increases. The times indicated in Table I are usually long enough for 10.0day mouse embryos. Observed background staining is proportional to the size of the specimen. It may determine the upper age limit for which the procedure works. For some reason, background is variable between individual large specimens, and so it may be necessary to process many before a few suitable for good photographs are found. Smaller embryos yield uniform results. Dissection
The goal of dissection is to get the embryos from the uterus to the fixative in good condition and in a reasonably short time. A period of 30 rain should be fine. Dissect in PBS (phosphate-buffered saline, without Ca2+/Mg 2+) and keep the embryos on ice as much as possible. Good dissections, especially for the early stages, require practice, steady hands, a well-adjusted dissecting microscope with a good field depth, and undamaged forceps. The stages between egg and late blastocyst are flushed out of the fallopian tubes and uterus. Between implantation and approximately 6.5 days of gestation, the embryos are extremely difficult to find and dissect
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FIG. 1. Flattening an early neurulation embryo. A dorsal view of an 8.0-day embryo is shown. The covering amnion has been removed and two vertical cuts made in the lateral amnion (indicated by arrows). Once the cuts are made, the embryo will flatten out.
away from the mass of decidual tissue. Stages between 6.5 and 7.5 days are manageable but require practice. Later stages are easier. Hogan et al. 4 give detailed instructions for removing the different stages; however, if you are unfamiliar with the dissections, it is best to get a demonstration from someone knowledgeable. During gastrulation and early neurulation (~6.5-8.0 days), the embryo is U-shaped and is bound in the amnion and yolk sac. Although wholemount staining will work on the entire structure, it may be difficult to photograph the results. The embryo will flatten if, before fixing, the amnion cap is dissected off and two cuts are made in the lateral amnion (Fig. 1). After approximately 9.0 days of gestation, the neural tube closes. The lumen of the tube sometimes nonspecifically binds both primary and secondary antibodies, causing high background. To both reduce this and improve penetrability, the thin roof of the hindbrain may be opened and the embryos left to gently rock in PBS at 4° for 10 rain prior to fixing. Torn edges and crushed tissues often show high background staining. Fixation
A perfect fixative would preserve antigenicity and morphology, while leaving the embryo completely open to the passage of antibody and other reagents. Fixatives fall into two groups: those that work by covalently cross-linking proteins, forming a meshwork (e.g., aldehyde fixatives), and 4 B. Hogan, F. Costantini, and E. Lacey, "Manipulating the Mouse Embryo: A Laboratory Manual." Cold Spring Harbor Laboratory, Cold Spring Harbor, New York, 1986.
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those that precipitate the cellular components (e.g., organic solvents). Sternberger discusses their preparation and relevance to immunohistochemistry in more detail. 5 Recipes for two good starting fixatives, one from each group, are given below. The same fixatives used for sectioned material may be tried with whole-mount procedures, although a fixative that works for one may not work for the other. In particular, glutaraldehyde-containing fixatives may generate an almost impenetrable mesh of tightly cross-linked proteins. 4% Paraformaldehyde in PBS. Add 4 g of paraformaldehyde to 50 ml of water plus 1 drop of 1 N NaOH. Dissolve at 37° with shaking for 1 hr. Adjust the pH to approximately 7 with HC1, add an equal volume of 2 x PBS and then filter through Whatman (Clifton, NJ) No. 1 paper. Store at 4 ° and use within 1 day of preparation. Although less convenient, paraformaldehyde is more likely to preserve antigenicity than any other fixative. Dent's Fixative. Dent's fixative is 80% methanol, 20% dimethyl sulfoxide (DMSO). 6 If antigenicity is preserved, this may give a lower background than aldehyde fixatives, and requires less work. Fix on ice in a volume more than 20-fold greater than the sample volume, for between 1 hr and overnight, depending on the size of the embryos. Transfer the embryos to fresh fixative after the first 15-30 min.
Pretreatment Ideally, the pretreatments leave the embryo in such a condition that antibodies and other reagents can freely diffuse through them and bind to only the correct molecules. Which pretreatments are necessary depends on the fixative and the detection scheme used. Blocking Free Aldehydes. Aldehyde-fixed embryos require an incubation in 0.1 M glycine in PBS for 30-60 min at room temperature to block any free aldehyde groups. Blocking Endogenous Enzyme Activity. The fixation protocol may not abolish all endogenous peroxidase activity. At the risk of reducing antigenicity in some instances, the remaining activity may be eliminated by incubation in either PBS plus 0.3-1.0% H202, or methanol or Dent's fixative plus 0.5-5% H2Oz, for 3-4 hr at room temperature. The methanol treatment is preferable because the aqueous reaction may produce destructive oxygen bubbles within the embryo, especially at higher HzO2 concentrations. Most endogenous phosphatase activities are inhibited by includ5 L. A. Sternberger, " I m m u n o c y t o c h e m i s t r y . " Wiley, New York, 1986. 6 j. A. Dent, A. G. Poison, and M. W. Klymkowsky, Development (Cambridge, UK) 107, 35 (1989).
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ing I-2 mM levamisole prior to and during the incubation with the enzyme substrate. 7 If the embryos are still in methanol after completing these blocking steps, they should be rehydrated into PBS through a series of methanol dilutions in water (e.g., 50%, 20%, PBS, for 30 min each). Blocking Nonspecific Antibody Binding. The final pretreatment, antibody dilutions, and washes are all done in PBS plus 0.1-0.2% detergent plus 1-5% protein. The protein may be bovine serum albumin (BSA) or nonfat milk powder (both used at 1-3%) or nonimmune serum from the animal in which the secondary antibody was raised (5-20%, v/v). Nonimmune serum can be heat-treated for 30 min at 56° to inactivate complement. In our hands, milk powder worked the best. The detergent added is usually nonionic: Nonidet P-40, Triton X-100, or Tween 20. Both the protein and the detergent decrease background staining. The excess protein blankets sites in the embryo that interact nonspecifically with the antibodies. The detergent may weaken nonspecific antibody binding. Which protein and detergent will work best needs to be determined empirically for each antigen-antibody combination.
Antibody Incubations Dilute the primary antibody in the last blocking solution (PBS + protein + detergent) and incubate the embryos for a few hours to overnight, depending on their size. Keep the embryos gently rocking or rolling. Long incubations with a low concentration of antibody at 4° will give lower background staining than short incubations with high concentration of antibody. The success of a whole-mount protocol depends greatly on the characteristics of the primary antibody. Antibodies vary, among other things, in their specificity (how well they recognize the correct antigen to the exclusion of others), their affinity (how stable the antigen-antibody interaction is), and their source (polyclonal antisera or monoclonal). An antibody with poor specificity will generate increased background, which, because of the thickness of the material, will readily obscure specific staining. Because the antigen-antibody complex is noncovalent, and bound antibody is in equilibrium with free antibody, some of the bound antibody will be lost during the washes. In sectioned material, where 30 min of washing is usually sufficient, most of the antibody will remain bound, even if it does not have a high affinity. In the whole-mount procedures, 7 B. A. Ponder and M. M. Wilkins, J. Histochem. Cytochem. 29, 981 (1981).
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especially for large embryos, the wash steps may span most of a day, and there is the possibility that much of the signal may be lost if the antibody has a low affinity. A polyclonal antiserum usually contains antibodies recognizing more than one epitope. Consequently, its apparent affinity (acidity) is often higher than that of a monoclonal antibody, which will recognize only one epitope. There are specific problems associated with using mouse monoclonal antibodies on mouse tissues. Self-tolerance makes the production of monoclonal antibodies against epitopes present in the mouse less likely. For this reason, a mouse monoclonal antibody raised against a conserved non-mouse protein may recognize the protein in a wide variety of species except the mouse, since the mouse immune system may preferentially respond to regions of the injected protein that are different from the endogenous version. Also, embryonic or maternal mouse immunoglobulin G (IgG) present in the embryo will bind to any anti-mouse secondary antibody used and so increase background. To keep the embryos as free of mouse immunoglobulin as possible, dissect away maternal and extraembryonic tissues, layer by layer, in different changes of PBS, and wash them thoroughly in PBS before fixing. Remaining immunoglobulins may be blocked by incubating the embryos with unlabeled secondary antibody and then washing prior to the addition of primary antibody. This problem may be avoided by labeling the primary antibody, although this will reduce the sensitivity. A preferable alternative is to use rat monoclonal antibodies. Anti-rat immunoglobulin secondary antibodies which do not cross-react with mouse immunoglobulin are commercially available. Problems arising from self-tolerance will also be reduced. Polyclonal antisera also have disadvantages. Unlike monoclonal antibodies, they contain a majority of nonspecific antibodies, which may cause background. To remedy this, the antiserum may be either diluted, affinitypurified against the antigen, or preadsorbed. If the specific antibody titer is high, then it may be possible to dilute the antiserum to such an extent that the background due to nonspecific antibodies is negligible. Affinity purification produces the cleanest antibody solution but requires 1 week to perform and a large amount of purified antigen; in addition, antibodies that have a very high affinity for the antigen may not elute well, thus stripping the antiserum of its best antibodies. Preabsorption to an acetone powder of embryonic mouse tissue that does not contain any of the desired antigen may remove nonspecifically binding antibodies. Harlow and Lane describe these techniques. 8 s E. Harlow and D. Lane, "Antibodies: A Laboratory Manual." Cold Spring Harbor Laboratory, Cold Spring Harbor, New York, 1988.
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Washes
Five consecutive washes in the blocking solution following antibody incubations are typical. Keep the embryos gently rocking or rolling at either 4° or room temperature. The duration must be determined empirically, but each wash should not need to be longer than 1 hr. If washes are too short, there will be increased background; if too long, there may be a loss of signal. The length of the washes is dictated by diffusion rates and the intensity of nonspecific antibody binding. In general, the rate of diffusion decreases for larger molecules, for larger embryos, and for the denser tissue meshworks generated by some fixatives (e.g., glutaraldehyde). Improving the blocking steps will most often reduce the background better than greatly increasing the wash times. Detection
The binding of primary antibody may be monitored, and the signal amplified, by a number of different methods. For whole-mount techniques, enzyme-linked and fluorochrome assays are usual. Briefly, the complex of antigen and primary antibody is detected by binding a secondary antibody coupled to either a dye, which is then observed by fluorescence microscopy, or to an enzyme, which generates an insoluble colored reaction product from a soluble substrate. Although the fluorochrome assays take less time and localize antigens more precisely, the enzyme-linked assays are preferable because they are more sensitive, permanent, and easier to photograph. Both horseradish peroxidase 9 (HRP) and alkaline phosphatase l° are used. Of the two, HRP is better, because it is smaller and its diaminobenzidine (DAB) reaction product is extremely stable. (Sodium azide, which is often added to protein solutions to prevent bacterial or fungal growth, must not be used during the procedure, since it is a potent inhibitor of HRP.) A variant technique, the avidin-biotin complex H,12 (ABC) method, also works well. The secondary antibody is tagged with biotin and bound to the primary antibody-antigen complex. This in turn is bound to a complex of avidin or streptavidin 13 and biotinylated peroxidase. Because of the additional amplification step, the technique is reputedly more sensitive (although this has been questioned), 5 permitting background arising 9 p. K. Nakane and G. B. Pierce, Jr., J. Histochem. Cytochem. 14, 929 (1966). 10 A. S. Bulman and E. Heydermann, J. Clin. Pathol. 34, 1349 (1981). ii j. L. Guesdon, T. Ternynck, and S. Avarameas, J. Histochem. Cytochem. 27, 1131 (1979). t2 S.-M. Hsu, L. Raine, and H. Fanger, J. Histochem. Cytochem. 29, 577 (1981). ~3 Streptavidin is preferable because of its greater affinity for biotin.
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from nonspecific antibodies to be reduced by further diluting the primary antiserum. Endogenous biotin may be a source of increased background. It may be blocked in the pretreatment step by first incubating the embryos with unlabeled streptavidin or avidin, then with biotin, followed by washing away the excess before adding the primary antibody. 14 Coupled secondary antibodies and ABC kits are available commercially. Irrespective of the chosen detection scheme, each incubation step should be followed by a thorough set of washes in PBS plus detergent plus protein.
Color Development Enzyme-linked assays require the addition of enzyme substrate. If an HRP detection scheme has been chosen then DAB is the best choice. DAB is a potential carcinogen. Treat all waste DAB with bleach. Following the final wash, incubate the embryos in PBS plus 0.3 mg/ml DAB for 30 min. Add H 2 0 2 to 0.03% and follow the color development under a dissecting microscope. If the development takes longer than 15 min, keep the embryos in the dark to avoid light-catalyzed DAB polymerization. To stop the reaction rinse the embryos three times in PBS plus 0.1% (v/v) Triton X-100. If the signal is strong, the color development may be the shortest step in the entire procedure and the HzO z may not completely permeate the embryo before the formation of background staining prompts termination of the reaction. To avoid this, try altering the antibody or reagent concentrations such that the color reaction takes 30 min or longer in the large embryos. The DAB color reaction can be enhanced by including 0.5% NiCI2 in the PBS plus DAB solution. On adding the nickel, a precipitate occasionally forms, which may be removed by filtration through Whatman No. 1 paper.
Clearing To see the staining pattern clearly, the embryos can be made more transparent or more opaque by transfer into solutions of different refractive indices. They are increasingly transparent in the following: methanol < PBS < glycerol < BABB. BABB (Murray's solution), is a mix of benzyl alcohol and benzyl benzoate (1 : 2). It is an obnoxious solution that dissolves polystyrene, but it works very well. Because BABB is nonmiscible with aqueous solutions, embryos should first be dehydrated 14 G. S. Wood and R. Warnke, J. Histochem. Cytochem. 29, 1196 (1981).
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through an alcohol series (20, 50, 100%) and then soaked in alcohol-BABB (1 : 1) before being transferred into BABB. To illustrate the relationship between the staining and the embryo, greater transparency is not necessarily better. Large embryos will need more clearing, but staining in small embryos or near the surface may be best observed if the tissues are more opaque. Background is also less apparent in more opaque embryos.
Microscopy and Photography The embryos are usually photographed submerged in an open dish of clearing medium such that they can be manipulated with forceps into different orientations. Whole mounts are more difficult to photograph than slides. Normal microscopes used for viewing slides are not suitable, since they usually have a shallow field depth. A good dissecting microscope with a camera or one of the microscopes designed for lower magnifications [e.g., Leitz (Wetzlar, Germany) Apozoom, Photomakroskop M400; M10] is preferable. Use a stable viewing platform to avoid vibrations. If the illumination starts to heat the mounting medium, then the resulting convection currents may degrade the image quality by moving the embryo or causing diffraction differences in the medium. Vary lighting conditions to get the best pictures (Fig. 2). Try reflected or dark-field illumination on more opaque embryos when the relation between the staining and the surface of the embryo is to be emphasized. Sometimes a combination of lighting works well. For more transparent embryos, transillumination is best. To photograph deep staining, large embryos must exhibit low background and be made almost completely transparent. Unfortunately, this results in a low contrast between the embryo and the clearing medium in which it sits (Fig. 2B). Staining is usually more obvious in color prints than in black and white, since the eye uses color cues. As journals typically prefer black-and-white figures, try an enzyme-linked detection assay that gives a dark-colored reaction product (e.g., nickel-enhanced DAB staining) for making black-andwhite prints.
Whole-Mount Immunohistochemistry and Scanning Electron Microscopy Scanning electron microscopy (SEM) is the best technique for observing surface detail and morphology. Surface antigens have been detected during SEM by using secondary antibodies coupled to a morphologically
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FIG. 2. Whole-mount immunohistochemistry on different stage embryos. All embryos were processed according to the example protocol given in Table I, using an affinity-purified polyclonal antiserum against the mouse engrailed proteins as primary antibody and a commercial affinity-purified goat anti-rabbit IgG coupled to HRP (Jackson Immunoresearch, West Grove, PA) as secondary antibody, and photographed using a Leitz macroscope and Ektachrome (Kodak, Rochester, NY) color slide film. Anterior is to the right. (A) Dorsal views of early 8.0-day embryos showing staining in the midbrain/hindbrain neuroepithelium. (Left) The DAB reaction was enhanced with NiC12. The embryo was photographed in methanol using reflected illumination. (Right) The DAB reaction was not enhanced. The embryo was cleared in BABB and transilluminated. (B) Lateral view of a 10.0-day embryo showing expression in the midbrain/hindbrain region, the somites, the ventral limb buds, and in the ventral spinal cord and hindbrain (arrows). The DAB reaction was not enhanced. The embryo was cleared in BABB and transilluminated. Bars: 0.3 mm.
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distinct marker 15 or by a detecting secondary electrons emitted from osmium deposited preferentially at the site of DAB polymerization. 16 We have reported a different method that may have some advantages. 17 Nickel ions, added to enhance the HRP/DAB reaction, are incorporated into the growing DAB polymer network. If the nickel is on or near the surface, it is detected during SEM by monitoring for element-specific Xray emissions. Machines that can do this are typically found in metallurgy or geology departments. Although untried, it should be possible to localize several different antigens simultaneously on the SEM image by doing serial whole-mount reactions, incorporating different metals (e.g., nickel and cobalt) at each step. There are some practical considerations to this technique. First, the nickel deposition must be on or within 1 /zm of the surface, since the electron beam does not penetrate far. Second, the embryo surface must be conductive. To accomplish this, coat the surface with as thin a layer of carbon as possible. A thicker coat, or a metal coat, will greatly reduce the sensitivity. Third, the procedure is slow, requiring a few hours to build up an adequate map. Increasing the beam intensity helps. Fourth, nickel emits characteristic soft K a X-rays at 7.427 keV, which may be partially absorbed by the bulk of the embryo. Ensure that the detector is capable of surveying the same region of the embryo as the electron beam.
Storage Embryos may be stored after fixation, after the first blocking steps (e.g., after aldehyde blocking and/or endogenous peroxidase inactivation), and after staining is complete. Embryos in most organic solvents may be safely left for a few days at 4° and for a few months at - 2 0 ° if first transferred into methanol. Aldehyde-fixed embryos may be stored in methanol as well if antigenicity is retained. If not, it may be possible to freeze and store the fixed embryos in liquid nitrogen without severely damaging the morphology. Once stained, the embryos are best stored at - 2 0 ° in methanol. The DAB reaction product is completely stable in methanol and is stable for at least a few days in BABB, but we have noticed variable loss of staining in embryos stored in BABB for several months at 4°. 15 R. S. Molday, in "Techniques in Immunochemistry" (G. R. Bullock and P. Petrusz, eds.), Vol. 2, p. 117. Academic Press, San Diego, 1983. 16 A. L. Hartman and P. K. Nakane, in "Techniques in Immunochemistry" (G. R. Bullock and P. Petrusz, eds.), Vol. 2, p. 103. Academic Press, San Diego, 1983. 17 C. A. Davis, D. P. Holmyard, K. J. Millen, and A. L. Joyner, Development (Cambridge, UK) 111, 287 (1990).
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TABLE II TROUBLESHOOTINGGUIDE Problem
Possible causes
Relevant section of text
Poor morphology
Damageduring dissection Damage during incubations or transfer Poorly fixed embryos Embryos poorly penetrable Inadequate blocking Nonspecific antibody b i n d i n g Wash times too short Wash volumes too small Concentration of antibody or other detection reagent too high Enzyme exposed to substrate for too long Excess enhancement Antigenicity not retained Rare antigen Poor primary antibody
Dissection GeneralPoints Fixation Fixation; Pretreatment; Dissection Pretreatment Pretreatment; Antibody Incubations Washes Washes AntibodyIncubations; Detection
Background
Poor signal
Variable results
Antibody or other detection reagents too dilute Embryos poorly penetrable One of reagents is bad Deep interior staining faint in large embryos Variable background in large embryos
Color Development Color Development Fixation Detection Antibody Incubations; Developing Working Protocol Antibody Incubations; Detection; Developing Working Protocol Fixation; Pretreatment; Dissection (Include positive controls) Dissection; General Points; Color Development General Points
Sectioning For a more detailed examination at the tissue level, embryos stained with D A B can be sectioned. The DAB reaction product is stable in xylene and wax, so follow normal paraffin wax embedding and sectioning procedures. Both staining and b a c k g r o u n d will appear much fainter in the sections than in the whole mounts. Therefore, if the e m b r y o s are to be sectioned, it m a y be desirable to let the color development reaction proceed until the b a c k g r o u n d starts to look quite bad. Developing Working Protocol The whole-mount procedures have a large number of parameters. F a c e d with an untried a n t i b o d y - a n t i g e n combination, the task is to find the set of conditions that give the best results. There are a few practical points that can make this less arduous. F o r the initial tests of a new antibody, use the smallest e m b r y o s that express the antigen well, since they will exhibit cleaner whole-mount
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results. Try the HRP indirect immunohistochemical detection method, since it is reasonably sensitive and straightforward. Fix some embryos in 4% paraformaldehyde and some in Dent's fixative. For each group test 7-8 different serial dilutions of the primary antibody. Working dilutions are typically between 1/20 and 1/1000 for polyclonal antisera or monoclonal antibodies from ascites, and between undiluted and 1/20 for hybridoma culture supernatants. If possible, include another primary antibody known to produce good whole mounts as a positive control, and use it to find a dilution range for the secondary HRP-coupled antibody (typically between 1/200 and 1/1000). Start with overnight antibody incubations at 4 °, and five 1-hr washes. Do not initially enhance the H R P - D A B reaction, since even a small amount of enhanced background may conceal specific staining. Following the initial trials, use the troubleshooting guide to help choose modifications to the procedure. It would take months of work to explore all the possible variations of the whole-mount procedure. If there is no staining or if there is unacceptable background after the first few trials of a new antigen-antibody combination, then it may be worthwhile to start raising more antibodes. Meanwhile, further alterations of the procedure can be tried.
Troubleshooting Table II lists the most common difficulties that may be encountered during the whole-mount procedure, their possible causes, and portions of the text relevant to their solution.
Controls
Artifactual staining is common in immunohistochemical techniques. The same controls are required in whole-mount procedures as in others (e.g., no antibodies, no primary antibody, primary antibody preadsorbed to the antigen). They are fully discussed elsewhere.18 False-negative results are more problematic. They may be due to an insufficiently sensitive procedure, masking or loss of epitopes, or insufficient penetration of the specimen by the antibodies or detection reagents. The last is especially applicable to whole-mount protocols, where the increased thickness and possibly more impermeable basement membranes of older embryos may make exposure of the entire sample t8 Chr. W. Pool, R. M. Buijs, D. F. Swaab, G. J. Boer, and F. W. Van Leeuwen, in "Immunohistochemistry" (A. C. Cuello, ed.), p. 1. Wiley, Chichester, UK, 1983.
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less likely. Finally, no set of controls is perfect, so consistent results using other techniques (e.g., Western blot or RNA in situ hybridization) are desirable. Acknowledgments I t h a n k Drs. I. Gitelman, A. L. Joyner, and N. Bonini for helpful c o m m e n t s .
[32] T e c h n i q u e s for L o c a l i z a t i o n o f S p e c i f i c M o l e c u l e s in Oocytes and Embryos .... By CALVIN SIMERLY and GERALD SCHATTEN
Introduction The fields of cell, developmental, molecular, and reproductive biology and genetics have tremendously benefited from studies on eggs at fertilization. 1'2 These early studies relied on systems like frogs and sea urchins in which fertilization in vitro and embryo culture were performed using simple solutions like seawater or pond water at room temperature. The design of methods for routinely and reliably obtaining excellent in vitro fertilization of many mammals now permits detailed experimentation on molecular and structural features of development in mammals. These investigations have led to many important and unexpected basic discoveries including genomic imprinting3-5; gametic recognition involving unique receptors and galactosyltransferases 6'7 atypical maternal inheritance patterns of the centrosome in mice8; both paternal and maternal inheritance
J T. Boveri, "Zellen-Studien: U e b e r die N a t u r der C e n t r o s o m e n , " Vol. 4. Fisher, Jena, G e r m a n y , 1901. 2 D. Mazia, Exp. Cell Res. 153, 1 (1984). 3 j. M c G r a t h a n d D. Solter, Cell (Cambridge, Mass.) 37~ 179 (1984). 4 M. A. H. Surani, S. C. Barton, and M. L. Norris, Cell (Cambridge, Mass.) 45, 127 (1986). 5 C. Sapienza, A. C. Peterson, J. R o s s a n t , and R. Bailing, Nature (London) 328, 251
(1987). 6 p. M. W a s s a r m a n , Annu. Rev. Cell Biol. 3, 109 (1987). 7 B. D. Shur, in " T h e Molecular Biology of Fertilization" (H. Schatten and G. Schatten, eds.), Cell Biology Ser., p. 38. A c a d e m i c Press, San Diego, 1989. 8 G. Schatten, C. Simerly, and H. Schatten, Proc. Natl. Acad. Sci. U.S.A. 88, 6785 (1991).
METHODS IN ENZYMOLOGY,VOL. 225
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