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Electron microscopy, immunostaining, cytoskeleton visualization, in situ hybridization, and three-dimensional reconstruction of Xenopus oocytes
Methods 51 (2010) 11–19
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Methods journal homepage: www.elsevier.com/locate/ymeth
Review Article
Electron microscopy, immunostaining, cytoskeleton visualization, in situ hybridization, and three-dimensional reconstruction of Xenopus oocytes Szczepan M. Bilinski a, Mariusz K. Jaglarz a, Matthew T. Dougherty b, Malgorzata Kloc c,* a
Department of Systematic Zoology, Institute of Zoology, Jagiellonian University, Krakow, Poland Biomedical Computation and Visualization Laboratory, Baylor College of Medicine, National Center for Macromolecular Imaging, Houston, TX 77030, USA c Department of Surgery, The Methodist Hospital and The Methodist Hospital Research Institute, Houston, TX 77030, USA b
a r t i c l e
i n f o
Article history: Accepted 9 December 2009 Available online 16 December 2009 Keywords: Oogenesis Xenopus Ultrastructure Immunostaining 3D reconstruction
a b s t r a c t Although the overwhelming development of molecular techniques in recent decades has made ultrastructural studies less popular, to the point that ultrastructural interpretation is becoming a dying art, it still remains an indispensable tool for cell and developmental biologists. The introduction of EM-immunocytochemistry and three-dimensional visualization methods allows us to complement the knowledge gained from ultrastructural and molecular approaches. Because the first clues about the functions of newly discovered genes often come from the subcellular localization patterns of their proteins or RNAs, in this chapter we describe the methods that allow for precise ultrastructural localization and visualization of protein and RNA molecules within the compartments, organelles, and cytoskeleton of Xenopus oocytes. Ó 2009 Elsevier Inc. All rights reserved.
1. Introduction The electron microscope (EM) was invented in 1931 by German engineers Ernst Ruska (awarded Nobel Prize for this invention in 1986) and Max Knoll. Flourishing during 1950s–1970s, the application of EM in cell and developmental biology resulted in a plethora of discoveries, including descriptions of the ultrastructure of the nucleus (reviewed in [1]) and various cytoplasmic organelles in Xenopus oocytes and embryos ([2,3], reviewed in [4]). Although ultrastructural studies became less popular (and certainly less publishable) with the extraordinary progress and development of molecular techniques, they still remain an indispensable tool for cell and developmental biologists. Therefore, it is worth emphasizing, as has been done most recently by Stenmark [5], how much of our current understanding of cellular and molecular processes we owe to ultrastructural studies. While the introduction of EM-immunocytochemistry allowed us to complement the knowledge gained from ultrastructural and molecular approaches, the use of EM, the number of accomplished electron microscopists who are able to interpret ultrastructural data, the number of people who are adept at electron microscopy, and the number of electron microscope facilities are declining at an alarming rate both in USA and in * Corresponding author. Address: Immuno-Biology Laboratory, The Methodist Hospital Research Institute, 6565 Fannin St., SM 8-034 Houston, TX 77030, USA. Fax: +1 713 793 7151. E-mail address: [email protected] (M. Kloc). 1046-2023/$ - see front matter Ó 2009 Elsevier Inc. All rights reserved. doi:10.1016/j.ymeth.2009.12.003
Europe [6–8]. This is especially unfortunate considering the fact that very often the first clue as to the function of a newly discovered gene comes from the localization pattern of its protein or RNA within cellular organelles or cell/embryo subcellular compartments. In this chapter we describe the methods that enable precise ultrastructural localization and visualization of protein and RNA molecules within the compartments, organelles, and cytoskeleton of Xenopus oocytes.
2. Post-embedding immunogold EM In the immunogold EM technique, the localization of an antigen is revealed indirectly by the application of primary antibody followed by secondary antibody conjugated to colloidal gold (5– 30 nm) or nanogold (0.8–1.4 nm) particles (Figs, 1A–D, 2A). The advantage of using the gold tags lies in their small size, electrondense nature, and strong scattering of electrons, which allow for enhanced contrast between the label and the tissue. In Xenopus oocytes, this technique has been used to study, for example, the formation of germinal granules [9]. The post-embedding immunogold EM procedure can be divided into five steps: (a) (b) (c) (d) (e)
oocyte/ovary fixation, oocyte/ovary embedding, ultrathin sectioning, immunogold labeling of sections, staining/contrasting of immunolabeled sections.
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2.1. Oocyte/ovary fixation Xenopus oocytes or small pieces of the ovaries are fixed in 2–4% formaldehyde, freshly made from paraformaldehyde (Sigma–Aldrich) in 0.1 M Sörensen’s phosphate buffer (pH 7.2–7.4), or, alterna-
tively, 0.5–2% glutaraldehyde (EM grade, Ted Pella, Inc.) in the same buffer. Glutaraldehyde fixation generates better preservation of ultrastructure but usually has a deleterious effect on tissue immunoreactivity. Glutaraldehyde is a strong denaturating agent whose protein cross-linking properties may result in modifications
Fig. 1. (A–D) Post-embedding immunogold EM. A. XVLG (vasa homologue, primary antibody: mouse anti- XVLG, gift from K. Ikenishi, Osaka City University, Osaka, Japan) in nuage material (arrow) attached to the nuclear envelope of the germinal vesicle (gv); c – cytoplasm. B. Accumulations of the nuage material (arrows) labeled with anti-Sm antibody (primary antibody: Y-12, gift from J. Gall, Carnegie Institution for Science, Baltimore, USA). gv – germinal vesicle, c – cytoplasm. (C) Mitochondrial cement between mitochondria (m) labeled with anti-Sm antibody. (D) Nucleolus (nu) in the germinal vesicle labeled with K-121 antibody (Oncogene Research Products, Calbiochem), which recognizes the 5’ terminal 2,2,7-TMG cap of certain small nuclear and nucleolar RNAs. A–D. Secondary antibody conjugated with18 nm gold particles. (A–D) Stage I Xenopus oocytes, see [9] for details. Bars: (A–C) 200 nm, (D) 500 nm.
Fig. 2. (A) Post-embedding immunogold EM. Bundles of intermediate filaments (arrows) in the cytoplasm of the prefollicular cells surrounding young oocytes/cystocytes in the Xenopus ovary labeled with anti-Par3 antibody (Zymed). (B) Extraction method. Fragment of the Triton-extracted oocyte, arrows point to microtubules. (C) Taxol enhanced microtubular cytoskeleton. Electron micrograph of one pole of the mitotic spindle in the dividing cystocyte within the Xenopus ovary. Arrowhead – centriole, arrows – microtubules of the mitotic spindle, asterisks – chromosomes. Bars: (A) 200 nm, (B) 400 nm, (C) 500 nm.
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of the epitope(s) or affect the binding and/or accessibility of the antibody. The choice of fixative and its concentration should be determined for a particular antigen and the corresponding antibody by trial and error. In our experience, fixation of Xenopus oocytes/ovaries in 1% glutaraldehyde or a mixture of 2% formaldehyde and 1% glutaraldehyde works best (depending on the nature of the antigen and antibody). The advantage of using the mixture of fixatives is that formaldehyde penetrates the tissue more rapidly and stabilizes cell structures, which are subsequently fixed more firmly by glutaraldehyde. As a result, a good balance is struck between maintaining cell ultrastructure and, at the same time, retaining high immunoreactivity. In contrast to the standard EM fixation protocol, in immuno-EM there is usually no post-fixation with osmium tetroxide, which may have an adverse effect on antigenicity and may prevent proper UV-induced resin polymerization (e.g., Lowicryl). The fixation step is crucial to the success of the immuno-EM technique. Therefore one should spare no effort in ensuring the structure of the oocytes/ovaries is preserved as closely to its natural state as possible, while keeping antigen properties uncompromised. The viability of an antigen should be tested in a range of conditions (different fixatives, resins, buffers, etc.). It is important to remember that at each step many factors may influence the final result, so keeping an accurate and detailed record of the procedures used is strongly recommended. This should include the fixative type, concentration, and time-course. Because so many variables may affect the final outcome, it is often necessary, especially in the case of previously untested antibodies, to perform preliminary immunocytochemical experiments using light and/or fluorescence microscopy. This enables a large number of variables to be tested in a more efficient, time- and cost-saving manner. If conducted properly, such preliminary experiments can also provide crude information about the distribution of the antigen(s) in the analyzed cell/tissue, which subsequently can be further refined in the samples processed for electron microscopy. 2.2. Oocyte/ovary embedding Xenopus oocytes can be embedded in practically any resin used in EM studies, but Epon 812 or its equivalents [e.g., Poly/Bed 812 (Polysciences, Inc.), EMbed 812 (Electron Microscopy Sciences)], and Lowicryl K4 M (Electron Microscopy Sciences) are used most frequently. A word of caution should be added regarding Epon embedding. This epoxy resin polymerizes best at 60 °C for 64 h, and there are some indications that this may reduce or even completely abolish immunoreactivity. Therefore, for immuno-EM studies it is advisable to conduct the polymerization at a lower temperature (50 °C) and reduce its time to 48 h. In general, acrylic resins such as Lowicryl K4 M and HM20, L.R. White, and L.R. Gold are better suited for immuno-EM as they do not polymerize into three-dimensional crosslinked networks, in contrast to epoxy resins. However, embedding in these resins is more cumbersome because they require anaerobic conditions for polymerization and usually prolong exposure to indirect UV light (360 nm) at low temperature (e.g. Lowicryl K4 M).
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dle than Lowicryl sections. However, prior to antibody labeling, they require gentle etching with sodium hydroxide and/or hydrogen peroxide to remove a thin layer of the resin and to reveal antigenic determinants/epitopes (Section 2.6). 2.4. Immunogold staining of sections The optimal concentrations of primary and secondary antibodies should be determined by trial and error (the useful trial concentration is usually 1:50–1:200 for primary antibodies and 1:200– 1:500 for the secondary ones, see also below). It is good practice, or even necessity, to perform simultaneous control labeling experiments with each localization attempt. Such experiments should be carefully designed to avoid potential artifacts that may lead to false conclusions. The results of labeling experiments should be regarded with a certain degree of skepticism until the results are verified by control experiments. However, it is worth noting that some control experiments are not always necessary or feasible (e.g., adsorption of antibodies with an excess of a studied antigen or with homogenized tissue (ovary/oocytes) containing the antigen), especially if commercial or previously tested antibodies are used. Both negative and positive controls should be utilized whenever possible. The negative control is done by simply omitting the primary antibody while keeping the rest of the labeling procedure unchanged. This permits the estimation of the ‘‘background” level of labeling due to non-specific interactions between the secondary antibody and the cell constituents, the resin, or both. The results of the negative control also facilitate adjustments to the secondary antibody concentration. The positive control, on the other hand, requires the use of a primary antibody that has previously been demonstrated to cross-react with antigen(s) of the given species (in this case X. laevis). By observing tissue labeling in the positive control, one can be more confident that the absence of labeling with a newly tested antibody did not result from technical problems. Another possible control experiment is to use pre-immune serum (or normal serum from a non-immunized animal of the same species in which the antibody was generated) instead of the primary antibody. Secondary antibodies conjugated to gold particles (5–30 nm in diameter) are available from several vendors, including Sigma–Aldrich, Jackson ImmunoResearch Lab., Inc., and Nanoprobes, Inc. It should be mentioned that it is also possible to visualize the distribution of two (or more) antigens in the same section. This procedure is termed ‘‘multiple labeling” and requires the application of secondary antibodies with gold particles of sufficiently different sizes (e.g., 5 and 20 nm). 2.5. Staining/contrasting of immunolabeled sections Because osmium is not used in this procedure (Section 2.1), tissue contrast is often very low. Therefore, staining procedures with Reynold’s solution (lead citrate) and uranium acetate solutions are usually slightly longer than in the standard EM protocol. However, the length of staining depends on the fixation procedure and the resin type used for embedding. Extended staining, even though it improves contrast, may obscure the gold particles.
2.3. Ultrathin section preparation 2.6. Protocol for post-embedding immunogold EM Gray or silver sections (40–70 nm thick) are cut with an ultramicrotome and transferred onto nickel or gold grids (copper grids are not recommended due to increased risk of artifactual precipitates). To prevent grid bars from potentially obscuring places of interest, it is best to use single-slot grids or, alternatively, grids with very thin cross-bars and a high open area. However, singleslot grids require prior coating with formvar (polyvinyl formal; Polysciences, Inc.) or carbon films to give extra support to ultrathin sections. In our experience, Epon sections are easier to cut and han-
(1) Fix the oocytes/ovaries in 2–4% formaldehyde, freshly made from paraformaldehyde in phosphate buffer (pH 7.2–7.4) for up to 1 h at room temperature or in 0.5–2% glutaraldehyde in the same buffer for 30 min; alternatively, use a mixture of 2% formaldehyde and 1% glutaraldehyde in the same buffer for 30 min. (2) Rinse several times in phosphate buffer (usually 3 10 min) at room temperature.
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(3) Dehydrate, infiltrate, and embed the oocytes in the resin of choice (according to the manufacturer’s protocol). (4) Cut sections with an ultramicrotome and collect them on coated/uncoated nickel grids. (5) Etching: incubate1 Epon sections (the etching step is not necessary for the Lowicryl sections) in a drop of 0.25% sodium hydroxide in 100% ethanol for 1–5 min, followed by 3 washes in 100% ethanol. (Hold grid by the edge with fine forceps and dip several times in each of three small glass beakers containing 100% ethanol, or, alternatively, grids can be gently rinsed using a squirting wash bottle.) After rinsing 3 times in distilled water, immerse the grids separately in a drop of 5–10% hydrogen peroxide (freshly made from a concentrated stock solution) for 5–15 min. Rinse 5 times in distilled water.
(6)
(7)
(8) (9)
(10)
(11)
(12)
The incubation of grids can be carried out in small drops of solution placed on a piece of clean dental wax (each grid in a separate drop) or in small depressions made with the cap of an Eppendorf microtube in a piece of Parafilm (Sigma–Aldrich). The grids can be either fully immersed in the solution or placed on the top of the drop (depending on individual preference). Great care should be taken not to damage the film coating the grids with forceps. Block the non-specific binding sites. Incubate the grids in a drop of freshly made 2–5% bovine serum albumin (BSA) in phosphate-buffered saline (PBS) with the addition of 0.05% sodium azide (NaN3) for 20–30 min at room temperature. A similar concentration of non-fat powdered milk or 1–5% fish gelatin (Sigma–Aldrich) in PBS can be used instead of BSA, depending on the type of antibody and the manufacturer’s recommendations. In some instances, it may be beneficial to briefly expose the grids to glycine (50 mM, 10–30 min) to quench free (unreacted) aldehyde groups. Incubate sections in primary antibody diluted in 1% BSA + PBS + NaN3 overnight at 4 °C or, alternatively, for 2– 4 h at room temperature. The optimal antibody concentration can only be found by trial and error. A good starting point is a 1:50 dilution of monoclonal antibodies and 1:200 for polyclonal antisera, but this varies greatly depending on the antibody and the manufacturer. Rinse 5 times in PBS (several dips each). Label the sections with secondary antibody complexed to colloidal gold particles of required size and diluted in BSA + PBS + NaN3 for 2 h at room temperature. Care should be taken to choose a secondary antibody that is fully compatible with the primary antibody. Again, the optimal antibody dilution should be discovered by trial and error; one can start with a 1:200 dilution. Rinse twice in PBS followed by three washes in distilled water. At this point, leave the grids out to dry or proceed with contrasting. Contrast sections with uranium acetate and lead citrate according to standard EM procedures. The optimum time for each contrasting step has to be established empirically. Finally, after rinsing several times with distilled water and drying, sections are now ready for viewing in TEM.
oocytes. This is due primarily to the presence of numerous cytoplasmic organelles, nutrient/reserve materials accumulated in yolk granules, and lipid droplets that mask the cytoskeletal components/filaments. One way to overcome this difficulty is to remove the ‘‘unwanted” cell constituents with detergent (Triton X-100) extraction. We have adopted a gentle extraction method, originally developed for insect oocytes [10], to Xenopus oocytes. This method is particularly suitable for visualization and analysis of microtubule distribution (Fig. 2B). The most critical step in this method is the Triton extraction. Care should be taken not to prolong this step, as this may result in complete destruction of the cytoskeletal structures. The whole procedure requires 12–13 steps, including three fixation steps: (1) Extraction. (2) Fixation in fixative 1, 15–30 min at room temperature. (3) Rinsing in 0.1 M phosphate buffer pH 7.4, 3 15 min at room temperature. (4) Fixation in fixative 2, 1–1.5 h at 4 °C. (5) Rinsing in 0.1 M phosphate buffer pH 7.4, 3 30 min at room temperature. (6) Fixation in fixative 3, 1–1.5 h at room temperature. (7) Rinsing in distilled water, 3 30 min at room temperature. (8) En bloc staining in 2% aqueous uranyl acetate for 2 h or overnight at room temperature. (9) Rinsing in tap water, 3 5 min. (10) Dehydration in increasing concentrations of ethanol (30% – 2 10 min, 50% – 2 10 min, 70% – 3 10 min, 96% – 3 15 min, 100% – 3 30 min) and in acetone (3 10 min); all steps at room temperature. (11) Embedding in Epon 812 or its equivalents (e.g. Poly/Bed 812 from Polysciences, Inc., EMbed 812 from Electron Microscopy Sciences) according to the manufacturer’s protocol. (12) Cutting block into ultrathin sections. (13) Additional section contrasting (this step is optional). 3.1.1. Extraction buffer Defolliculated Xenopus oocytes or small pieces of ovaries are immersed in the following extraction buffer:
137 mM NaCl 5 mM KCl 1.1 mM Na2HPO4 0.4 mM KH2PO4 4 mM NaHCO3 12 mM MgCl2 2 mM EGTA 50 mM 2-(N-morpholino) ethanesulfonic acid (MES) 5.5 mM glucose 1% Triton X-100
(all chemicals available from Sigma–Aldrich, St. Louis, MO, USA). Extraction time: 5–15 min, depending on the specimen size. It is worthwhile to extract 2–3 bunches of oocytes at 5-min intervals to determine the optimal timing.
3. Methods for cytoskeleton visualization 3.1.2. Fixative 1 3.1. Extraction method 2.5% glutaraldehyde in the extraction buffer. It is challenging to analyze cytoskeleton distribution and spatial organization at the ultrastructural level in cells as large as Xenopus
3.1.3. Fixative 2
1
It is recommended to do this step and the following incubation of sections with the antibodies in a moist chamber; one can be easily prepared using a covered Petri dish lined with moist filter paper.
2% osmium tetroxide, 0.8% potassium ferrocyanide,
S.M. Bilinski et al. / Methods 51 (2010) 11–19
in 0.1 M phosphate buffer pH 7.4. The mixture should be prepared just before use and kept at 4 °C. 3.1.4. Fixative 3 0.15% tannic acid in 0.1 M phosphate buffer pH 7.4. 3.2. Fixation with Taxol Another method of improving visualization of the microtubule cytoskeleton is the addition of taxol to the fixative. Taxol binds tightly to microtubules and stabilizes them in cells and in vitro [11,12]. As no osmium tetroxide is used during the fixation step, the cell membranous structures are only lightly stained, while microtubules and centrioles are highly contrasted/enhanced (Fig. 2C). This method has been used successfully to study the organization of Xenopus germline cysts [13].
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3.3.1. Microtubule stabilizing fixative 2% formaldehyde, 3% glutaraldehyde (EM grade, Ted Pella, Redding, CA), in 0.1 M sodium cacodylate buffer pH 7.3 (Polysciences, Warrington, PA), 10 lM taxol (Cytoskeleton, Denver, CO). 4. Whole-mount (pre-embedding) EM immunostaining of Xenopus oocytes The pre-embedding (whole-mount) immunostaining technique is much faster than performing immunostaining on sections. The only potential limitation is the permeability of the tissue. Thus, this method is great for staining the isolated oocytes and the surface layers of the whole embryo, but it may fail to label deeper embryonic layers (Fig. 3A, C–E). 4.1. Fixation
3.3. Fixation to improve microtubule visualization Fix oocytes in microtubule stabilizing fixative for 30 min at room temperature. Wash in cacodylate buffer 3 10 min. Stain in 0.5% uranyl acetate for 30–60 min. Dehydration and embedding according to the standard EM protocol.
Manually or enzymatically (with collagenase, cat # C6885, Sigma–Aldrich, St. Louis, MO, USA) defolliculated stages-I–VI oocytes are fixed as follows: 4.1.1. For non-cytoskeletal protein immunostaining Oocytes are fixed in 2% formaldehyde plus 0.5% glutaraldehyde (both-EM grade from Ted Pella Inc., Redding, CA, USA:
Fig. 3. Whole mount (pre-embedding) nanogold immunostaining and in situ hybridization. (A) Fatvg protein is present on the surface of vesicles (v) in the stage III Xenopus oocyte (see [17] for details), y – yolk granule. Secondary antibody conjugated with 1.4 nm gold. (B) In situ hybridization, Xcat2 mRNA (arrows) is present within the germinal granule; m – mitochondria. Anti-digoxigenin antibody, conjugated with 0.8 nm gold, and silver enhanced. (C–E) Oocytes labeled with anti-cytokeratin C11 antibody (Sigma– Aldrich) and nanogold-conjugated secondary antibody, and silver enhanced (see [15] for details). Arrows indicate cytokeratin filaments distributed in the vegetal cortex of the stage VI Xenopus oocyte, y – yolk granule. (C) Low magnification view of the fragment of the vegetal cortex of labeled oocyte. Semithin section (0.7 lm) stained with 1% methylene blue in 1% borax. (D and E) Ultrathin sections stained with uranyl acetate and lead citrate. Bars: (A and B) 500 nm, (C) 5 lm, (D and E) 500 nm.
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glutaraldehyde 8% stock, cat# 18421; formaldehyde 16% stock, cat # 18505) in 1 phosphate-buffered saline (PBS) in moleculargrade water for 2 h at room temperature or overnight at 4 °C. Using this concentration of glutaraldehyde is a trade-off between the relatively good preservation of ultrastructure and higher immunoreactivity. Increases in the glutaraldehyde concentration (within a 1–3% range) will improve ultrastructure but will usually decrease immunoreactivity (Fig. 3A) [14]. 4.1.2. For cytokeratin immunostaining Oocytes are fixed in 1–4% formaldehyde (either EM-grade from Ted Pella, Inc. or from 37% formaldehyde stock, cat # F-1635 from Sigma–Aldrich, St. Louis, MO, USA) in 100% methanol (GC resolve grade, cat # A457 from Fisher Scientific, Pittsburgh, PA, USA) overnight at 20 °C. The preservation of the ultrastructure of the components of the oocyte and of the cytokeratin epitope are identical in samples fixed with formaldehyde concentrations ranging from 1% to 4% [15]. 4.2. Post-fixation washes Fixed oocytes are rehydrated and washed with slow shaking on a rotating platform as follows: 100% ethanol 2 10 min, 70% ethanol 15 min, 50% ethanol 15 min, 2 5 min washes in phosphatebuffered saline (PBS) containing 0.05% Tween 20 (cat #166–2404, Bio-Rad Laboratories, Hercules, CA), 10 min in PBS–0.25% Triton X-100 (cat # X100, Sigma–Aldrich, St. Louis, MO, USA), and 2 5 min with PBS–0.05% Tween 20. 4.3. Blocking, antibody incubation, and washing All steps are performed with slow shaking on a rotating platform. Non-specific binding is blocked by incubating 6 h in casein blocking buffer (cat #161–0782, Bio-Rad Laboratories, Hercules, CA) with 0.05% Tween 20 at room temperature. This is followed by incubation with the appropriate primary antibody. For cytokeratin immunostaining, oocytes are incubated with anti-pan cytokeratin clone C-11 mouse monoclonal antibody (cat # C293, Sigma– Aldrich, St. Louis, MO, USA). All primary antibodies should be used at 1:50 dilution in casein blocking buffer containing 0.05% Tween 20, overnight at 4 °C. Oocytes are then washed 4–6 h at room temperature or overnight at 4 °C with PBS–0.05% Tween 20. The overnight washing significantly decreases the background. After washing, oocytes are incubated overnight at 4 °C with secondary antibody (appropriate for the given primary antibody, i.e., anti-mouse, anti-rabbit, etc.) conjugated to 1.4 nm gold (Nanoprobes, Yaphank, NY, USA) and diluted to 1:50 with casein blocking buffer containing 0.05% Tween 20. The next day, oocytes are washed 3 15 min with PBS–0.05% Tween 20, post-fixed for 20 min in 1% glutaraldehyde in PBS– 0.05% Tween 20, washed 3 15 min in molecular-grade water, and silver- or gold-enhanced.
enhancement, oocytes are washed 3 10 min in molecular-grade water and then post-fixed in fixative that contains 2% glutaraldehyde (EM-grade from Ted Pella, Inc., Redding, CA, USA) in 1 PBS. Samples can be kept in this fixative at 4 °C for several days before embedding. 4.5. Embedding and sectioning Whole-mount nanogold-immunostained and silver- or gold-enhanced oocytes are contrasted with 0.5% uranyl acetate only (without osmium tetraoxide, which damages the metal label) and, after dehydration in increasing concentrations of ethanol, infiltrated and embedded in LX-112 medium (Epon substitute; Ladd Research Industries, Burlington, VT, USA). The samples are polymerized in a 70 °C oven for 2 days. Semithin sections (0.7–1 lm) are stained with 1% methylene blue in 1% borax (Fig. 3C). Ultrathin sections (70–100 nm) are cut, stained for 5 min in 1% aqueous uranyl acetate, and 2 min in 1% aqueous lead citrate at room temperature in a Leica EM Stainer (Leica, Deerfield, IL), and examined in a JEM 1010 transmission electron microscope (JEOL, USA, Inc., Peabody, MA) at an accelerating voltage of 80 kV (Fig. 3D–E). Digital images are obtained using AMT Imaging System (Advanced Microscopy Techniques Corp, Danvers, MA). 5. Whole-mount (pre-embedding) EM in situ hybridization of Xenopus oocytes 5.1. Fixation Fixative for electron microscopy in situ hybridization of oocytes (identical to that used for Xenopus embryos and described in Kloc et al. [16]) contains 4% formaldehyde, 0.1% glutaraldehyde (both EM-grade from Ted Pella), 100 mM KCl, 3 mM MgCl2, 10 mM HEPES, 150 mM sucrose, and 0.1% Triton X-100, with a pH of 7.6. The main difference between this fixative and the fixative used for light microscopy in situ hybridization is the presence of glutaraldehyde, MgCl2, and sucrose, which have a stabilizing effect on the membranes and organelles. Defolliculated oocytes are fixed at room temperature for 1–2 h with shaking. 5.2. Post-fixation washes All subsequent treatments are performed with shaking. Fixed oocytes are washed 3 5 min with PBS–0.1% Tween 20 and then treated for 10 min in 10 mg/ml proteinase K in PBS–0.01% Tween 20. This step facilitates the penetration of the probe through the cortical layer of the oocyte. After proteinase K treatment, oocytes are washed for 5 min in 0.1 M triethanolamine (TEA) pH 8.5, then incubated for 10 min in 0.1 M TEA with 0.5% acetic anhydride, washed 3 5 min in PBS-Tween and post-fixed for 20 min in the fixative described above. After post-fixation oocytes are washed 3 5 min in PBS-Tween, and then hybridized overnight at 50 °C.
4.4. Silver or gold enhancement 5.3. Hybridization and post-hybridization washes Silver enhancement is performed with Light Insensitive Silver Enhancer (cat # 2013) and gold enhancement with GoldEnhance (cat# 2113) (both from Nanoprobes, Yaphank, NY, USA), according to the manufacturer’s protocol, for approximately 45 min. Both procedures are done without shaking. Although the general belief is that gold enhancement gives much lower background than silver enhancement, in our hands both types of enhancement work comparably well with very low background. To prevent over-staining, the development of the signal is monitored under the dissecting microscope and stopped (by transferring to molecular-grade water wash) when the oocytes reach a light-brown color. After the
Hybridization and all post-hybridization washes are performed exactly as described previously for light microscopy in situ hybridization [17]. In this citation we also gave a comprehensive list of all chemicals and solutions needed to perform in situ hybridization. 5.4. Antibody incubation After the last post-hybridization wash in PBS-Tween, oocytes are incubated for 15 min in G1 buffer (Boehringer Mannheim/ Roche) and then incubated overnight at 4 °C with a 1:30 dilution
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of anti-DIG 0.8 nm gold (Boehringer Mannheim/Roche) in G2 buffer. As far as we know, Boehringer Mannheim/Roche stopped producing anti-DIG 0.8 nm gold antibody. Although we never tried a different antibody, we believe that this antibody can be successfully replaced with any available anti-digoxigenin antibody conjugated with nanogold (0.8–1 nm) or 5-nm gold (using gold larger than 5 nm will probably create penetration problems). 5.5. Silver or gold enhancement After incubation with the antibody, oocytes are washed 415 min in PBS-Tween, post-fixed in 2% glutaraldehyde in PBSTween, and washed 3 15 min in molecular-grade water. Subsequently, they are silver- or gold-enhanced in Light Insensitive Silver Enhancer or GlodEnhance (Nanoprobes Inc, NY) for approximately 45 min. To prevent over-staining, the development of the signal should be monitored under a dissecting microscope. After the enhancement, oocytes are washed 3 10 min in molecular-grade water and then post-fixed in fixative that contains 4% formaldehyde, 2% glutaraldehyde (both EM-grade from Ted Pella), 100 mM KCl, 3 mM MgCl2, 10 mM HEPES, 150 mM sucrose, and 0.1% Triton X-100, pH 7.6. Oocytes can be kept in this fixative for several days before embedding. 5.6. Embedding and sectioning Fixed oocytes are stained in 0.5% uranyl acetate without osmium tetraoxide treatment, dehydrated in increasing concentrations of ethanol, and infiltrated and embedded in LX-112 medium (Epon substitute). Ultrathin sections (70–100 nm) are stained for 5 min in 1% aqueous uranyl acetate and 2 min in 1% aqueous lead citrate at room temperature and examined in a transmission electron microscope (Fig. 3B). 6. Digital visualization of Xenopus oocyte/organelles 6.1. Introduction Digital electron microscopy offers remarkable quantitative and qualitative advantages derived from computational visualization, such as a more nuanced assessment of Xenopus threedimensional (3D) structure. Prior to digital microscopy, noncomputational techniques used photographic film to capture the two-dimensional (2D) image projections of stained specimen sections. Modern techniques allow for direct image capture using CCDs or high-quality image scanning of photomicrographs. Due to the complexity of cellular structure and the inherent limitations of 2D imagery, it was difficult for microscopists to convey their remarkable understanding of three-dimensional structure prior to the advent of digital microscopy. Additionally, the management of such photomicrographs made collaboration difficult because of logistical limitations in organizing, indexing, and disseminating these 2D images. Significant technical advances in the design and manufacture of electron microscopes, most notably digital image acquisition and computer control of instrumentation, have allowed for sectional microscopy and electron tomography techniques. These methods provide greater insight into 3D cellular structure by establishing the distribution of lipids, proteins, and RNAs within the cytoplasm (Fig. 4). The ability to digitally assemble a sectional stack of 2D images allows for a greater understanding of cellular arrangement through interactive 3D visualization. Moreover, the application of computed axial tomography in electron microscopy allows for even finer 3D ultrastructure (2–10 nm resolution). At these resolutions, methods of specimen preparation become critical because of
Fig. 4. (A) Three-dimensional reconstruction. The mitochondrial cloud (light blue) extending between the nucleus (yellow) and the vegetal cortex contains germinal granules (red) at its vegetal tip (the METRO region). The nucleus is surrounded by mitochondria (dark blue). See [19] for details. (B) Three-dimensional reconstruction of a germinal granule showing the distribution of Xcat2 mRNA (green dots). See [18] for details. (C) Three-dimensional reconstruction of the Xenopus female germline cyst; three different views of the same cyst. Cytoplasm is gray, nuclei are red, mitochondria of primary mitochondrial cloud are green, centrioles blue, and ring canals are yellow. See [13] for details.
the likelihood of introducing artifacts that corrupt biological structure. For example, the use of heavy-metal stains frequently induces protein cross-linking that causes the creation of artificial protein
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complexes or lipid membrane destruction, which are observable at high resolution. Electron tomography repeatedly irradiates a small area of the specimen from a number of angles, requiring close monitoring of radiation damage at the time of 2D image acquisition. These raw tomographic tilt-series projections are computationally transformed by 3D reconstruction algorithms (e.g., direct Fourier inversion, filtered back projection, iterative algebraic). Each image is assembled as a function of the Euler angles of the projection, resulting in the 3D spatial image. A notable instrument limitation is the inability to fully rotate the goniometric specimen stage 90°, resulting in a ‘‘missing cone” of data that reduces the 3D resolution and produces smearing artifacts along the optical axis. Generally, specimens are no thicker than 500 nm, but this is a function of the operating voltage of the electron microscope and the composition of the specimen. Thinner specimens allow for greater tilt angles, and dual-axis tilt specimen stages can be used to improve resolution. Corresponding technical advancements in rapid cryo-freezing provide significant benefits to researchers who require the highest resolution and fidelity; in such cases, unstained cryo-hydrated specimens should be used. Such methods produce specimens that are highly correlated to in vivo samples because the speed of freezing essentially halts molecular and atomic activity, providing in situ snapshots of biological dynamics. These quickly frozen atomic positions present accurate glimpses of the chemical interactions and signaling within the cell. Another important advantage of cryo-microscopy is that these low-temperature specimens are better able to tolerate radiation damage caused by the electron beam; this is critical in the use of electron tomography because the same specimen region is irradiated numerous times. A number of hybrid approaches can be used, such as sectional cryo-tomography, freeze substitution, and cryo-negative staining. Researchers should familiarize themselves with the various methods and associated limitations during the development of a project plan, choosing an optimal set of protocols that will provide reproducible results in an efficient manner. The use of stains or antibodies at low resolution frequently provides quick satisfactory results that can be extended to higher-resolution methods. 6.2. Methods to obtain three-dimensional images A number of methods exist for researchers to obtain 3D imagery, the easiest being the Internet. The recent establishment of the Electron Microscopy Data Bank (EMDB, http://emdatabank.org) enables researchers to download an increasing number of 3D images of macromolecular complexes and cells. EMDB was established by the Protein Databank in Europe (PDBe) at the European Bioinformatics Institute, the Research Collaboratory for Structural Bioinformatics (RCSB) at Rutgers, and the National Center for Macromolecular Imaging (NCMI) at Baylor College of Medicine. EMDB provides a global deposition and retrieval network for cryo-EM 3D images, models, and associated metadata. Additionally, EMDB is a portal for software tools for standardized map format conversion; map, segmentation, and model assessment; visualization; and data integration. In tandem, a number of scientific journals are requiring the deposition of 3D imagery into open-access repositories as a condition of publication; this allows global communities of researchers to personally study the digital specimens, apply new computational techniques for verification, and initiate new tangential avenues of investigation, further enhancing the value of the initially funded research. The data format of choice is MRC2000, developed in 1982; it is the oldest and most widely used 3D electron microscopy image format. When the desired imagery does not exist at EMDB, researchers may contact global centers that explicitly provide these services,
requiring the researcher to vet and prepare specimens to be imaged. The National Center for Research Resources (NCRR, http:// ncrr.nih.gov) provides laboratory scientists and clinical researchers with the tools and training needed for research. NCRR funds a large number of biomedical technology research centers, such as the National Center for Macromolecular Imaging (http://ncmi.bcm.edu), specializing in high resolution cryo-electron microscopy and electron tomography; the National Center for Microscopy and Imaging Research (http://ncmir.ucsd.edu), specializing in cellular electron and optical microscopy; and the Resource for the Visualization of Biological Complexity (http://www.wadsworth.org/rvbc/), specializing in sectional time-resolved cryo-electron tomography. Similar centers exist in Europe, Japan, and China. Researchers are encouraged to contact the centers directly to determine the suitability of a center’s skills and resources to the researcher’s objectives and needs. Frequently the centers can accommodate a researcher’s skill set; for example, an experienced microscopist might allow a researcher direct access to the facilities, or a researcher who has no electron microscopy experience could merely provide the specimen to the center and outline any unique quality-control issues in handling their specimen. Depending on the frequency of specimen production and on quality control, it may be desirable to purchase a plunge-freezing apparatus and ship cryogenically frozen specimens to the center on electron microscopy grids. After microscopic image acquisition, the center can usually provide the researcher raw electron microscopic data, assist the researcher in using 2D-to-3D image reconstruction tools, or generate a final 3D structure that could be uploaded to the EMDB. For researchers and institutions considering the purchase of an electron microscope, these centers can also provide valuable consultation on implementation. For researchers who prefer to perform the 3D image reconstruction, there are a number of software tools available. Some software programs are integrated with the instrument, such as with FEI electron microscopes. There are commercial scientific visualization software products that may be purchased, such as Amira (http:// www.amiravis.com). Amira has some remedial tools designed for confocal microscopy that can be adapted to align sectional electron micrographs. Alternatively, NCRR-funded open-source software tools such as IMOD (http://bio3d.colorado.edu), developed by the Boulder Lab for 3D Electron Microscopy, are used throughout the structural biology community because of Boulder’s rigorous research and development at the forefront of electron tomography. The IMOD package is able to reconstruct serial sections or electron tomograms. It has sophisticated features, such as compensation for specimen deformation due to microtomy shearing and automated detection of gold particles used as fiducial markers in electron tomography. The Particle Estimation for Electron Tomography (PEET, http://bio3d.colorado.edu) package provides alignment and averaging of subvolumes extracted from tomograms, useful in studying identical sub-cellular structures such as the nuclear pore complex. 6.3. Methods of segmenting three-dimensional images Segmentation is a labor-intensive process requiring the researcher to be actively involved in annotating the boundaries between regions of interests. No comprehensive automated system exists at this time, but research is ongoing to develop reliable 3D pattern recognition. An inherent problem is automatically identifying components that have never been structurally observed before; this is further compounded by the interaction of multiple proteins in novel or poorly understood combinations. Low signal-to-noise ratios are another important limitation. The Amira, Shape, and Seg3D software packages are frequently used for segmentation of data sets, but other methods exist, such as The Watershed
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Transformation, eigenvector techniques, and multiphase level set segmentation. Amira, being primarily a generic visualization package, offers a number of software tools to analyze and dissect 3D images. Most important are the image filters, image segmentation editor, isosurfacing, and volume rendering. Image filters can be applied to the 3D reconstructed image prior to segmentation, allowing for greater contrast when annotating the boundaries. For tomograms, median and Gaussian filters are used to reduce the noise and minimize the missing-cone artifacts. The generic segmentation editor within Amira allows a researcher to manually go through the 3D image slice-by-slice along a preferred orthogonal axis. The researcher must make critical decisions about the annotation of biological regions of interest (ROI). The segmentation editor can be utilized with a mouse, but it is highly recommended that a graphics tablet and pen be used; the pen permits hand control that is more accurate, with significantly faster throughput. The editor has a number of segmentation tools, the most basic being Brush, similar to the brush tool in Adobe Photoshop. Other tools such as Magic Wand and Blowtool can significantly expedite annotation but require considerable effort to master because these tools are highly sensitive to image composition, particularly noise. The segmentation editor allows for the annotation of 256 ‘‘materials,” and multiple editors can be applied simultaneously to the same dataset. Shape (http://bio3d.colorado.edu) is an open-source package designed for automated segmentation of tomograms. It provides for denoising using 3D median filtering, extraction of features using Line Filter Transform and Orientation Filter Transform, and the extraction of contours along thin structures. Its major limitations involve datasets with large numbers of ribosomes, low-contrast filters, and microtubules that are out of plane. Researchers frequently use multiple packages, combining the best features of each by translating intermediate 3D images and segmented subvolumes between the packages. Although automated methods can significantly enhance the quality and speed of segmentation, ultimately the researcher must provide the final quality control either visually and/or manually. Seg3D (http://seg3d.org) is an open-source package developed by the University of Utah’s Center for Integrative Biomedical Computing, funded by the NCRR. Among its many features, it implements the National Library of Medicine’s Insight Tool Kit (ITK, http://itk.org), an advanced algorithmic software library developed to segment the Visible Human. Seg3D integrates automatic and manual features and at the same time is capable of managing very large images.
6.4. Methods to visualize three-dimensional images There are a number of software tools to visualize datasets using two methods of graphics: isosurfacing and volume rendering. Isosurfacing creates 3D polygons by contouring an image using a common contour threshold. These polygons can be individually colored with a 3D color field and by mapping a color-lookup table (CLUT) onto the color field. This can provide additional visual cues, such as radial distance or the highlighting of specific structures. Multiple isosurfaces may be displayed, each having independent thresholds, either on the same dataset or separate datasets. Volume rendering computes a 2D projection of a 3D image as a function of camera position and the 3D image voxels; the voxels are
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defined from 3D images convolved with a color transfer function, typically a red-green-blue transparency (CLUT). The primary computational limitations to visualization are the size of the 3D image, followed by the graphical computing power of the hardware. For example, if the computer memory is 4 gigabytes and the 3D image is 500 gigabytes, then visualization may not be possible without advanced computational techniques and resources; for the majority of datasets, the 3D image size is less than computer memory. Also, if the 3D image contains a significant amount of details or noise, the number of polygons may increase to the point that the graphics become sluggish or potentially cause an operating system failure. The use of isosurfacing provides convenience at computational expense, and the use of volume rendering can reduce graphical overload. Although there is a large number of visualization software packages, a few are frequently used by the electron microscopy community: Amira, Chimera, and SciRun. Chimera (http:// www.cgl.ucsf.edu/chimera/) is an open-source molecular-microscopy visualization tool developed by the Resource for Biocomputing, Visualization, and Informatics and is funded by NCRR. SciRun (http://scirun.org) is an open-source package developed by the University of Utah’s Center for Integrative Biomedical Computing, also funded by the NCRR. All three packages were developed in academic settings and have numerous technical features. Each offers a number of tutorials to demonstrate basic capabilities and illustrate methods of constructing highly complex visualizations and animations. The examples of 3D reconstruction of Xenopus oocytes and organelles are shown in Fig. 4. Acknowledgments We are grateful to Mrs. W. Jankowska and B. Szymanska for excellent technical support and Ms. E. Kisiel for helping with the figures. Panel A in Fig. 4 was reproduced with the permission of the publisher from the Fig. 4 panel A in the article published in Int. J. Dev. Biol. 48 (2004) 17-21. References [1] [2] [3] [4] [5] [6] [7] [8] [9] [10] [11] [12] [13] [14] [15] [16] [17]
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J.G. Gall, Z. Wu, C. Murphy, H. Gao, Exp. Cell Res. 296 (2004) 28–34. K.A. Al-Mukhtar, A.C. Webb, J. Embryol. Exp. Morphol. 26 (1971) 195–217. L.W. Coggins, J. Cell Sci. 12 (1973) 71–93. W.M. Bement, D.G. Capco, J. Electron, Microsc. Tech. 16 (1990) 202–234. H. Stenmark, Nature Rev. Mol. Cell Biol. 10 (2009) 582. H.J. Geuze, Trends Cell Biol. 9 (1999) 92–93. G. Griffiths, Trends Cell Biol. 11 (2001) 153–154. G. Griffiths, Eur. J. Cell Biol. 83 (2004) 245–251. S.M. Bilinski, M.K. Jaglarz, B. Szymanska, L.D. Etkin, M. Kloc, Exp. Cell Res. 299 (2004) 171–178. S.M. Bilinski, M. Jaglarz, Cell Motil. Cytoskeleton 43 (1999) 213–220. P.B. Schiff, J. Fant, S.B. Horwitz, Nature 277 (1979) 665–667. P.B. Schiff, S.B. Horwitz, Proc Natl. Acad. Sci. U.S.A. 77 (1980) 1561–1565. M. Kloc, S. Bilinski, M.T. Dougherty, E.M. Brey, L.D. Etkin, Dev. Biol. 266 (2004) 43–61. A.P. Chan, M. Kloc, C.A. Larabell, M. LeGros, L.D. Etkin, Mech. Dev. 124 (2007) 350–363. M. Kloc, S. Bilinski, M.T. Dougherty, Exp. Cell Res. 313 (2007) 1639–1651. M. Kloc, S. Bilinski, A.P. Chan, L.D. Etkin, Differentiation 67 (2000) 80–83. M. Kloc, L.D. Etkin, in: J.D. Richter (Ed.), Advances in Molecular Biology. A Comparative Methods Approach to the Study of Oocytes and Embryos, Oxford University Press, 1999, pp. 256–278. M. Kloc, M.T. Dougherty, S. Bilinski, A.P. Chan, E. Brey, M.L. King, C.W. Patrick Jr, L.D. Etkin, Dev. Biol. 241 (2002) 79–93. K. Wilk, S. Bilinski, M.T. Dougherty, M. Kloc, Int. J. Dev. Biol. 48 (2004) 17–21.
Contents lists available at ScienceDirect
Methods journal homepage: www.elsevier.com/locate/ymeth
Review Article
Electron microscopy, immunostaining, cytoskeleton visualization, in situ hybridization, and three-dimensional reconstruction of Xenopus oocytes Szczepan M. Bilinski a, Mariusz K. Jaglarz a, Matthew T. Dougherty b, Malgorzata Kloc c,* a
Department of Systematic Zoology, Institute of Zoology, Jagiellonian University, Krakow, Poland Biomedical Computation and Visualization Laboratory, Baylor College of Medicine, National Center for Macromolecular Imaging, Houston, TX 77030, USA c Department of Surgery, The Methodist Hospital and The Methodist Hospital Research Institute, Houston, TX 77030, USA b
a r t i c l e
i n f o
Article history: Accepted 9 December 2009 Available online 16 December 2009 Keywords: Oogenesis Xenopus Ultrastructure Immunostaining 3D reconstruction
a b s t r a c t Although the overwhelming development of molecular techniques in recent decades has made ultrastructural studies less popular, to the point that ultrastructural interpretation is becoming a dying art, it still remains an indispensable tool for cell and developmental biologists. The introduction of EM-immunocytochemistry and three-dimensional visualization methods allows us to complement the knowledge gained from ultrastructural and molecular approaches. Because the first clues about the functions of newly discovered genes often come from the subcellular localization patterns of their proteins or RNAs, in this chapter we describe the methods that allow for precise ultrastructural localization and visualization of protein and RNA molecules within the compartments, organelles, and cytoskeleton of Xenopus oocytes. Ó 2009 Elsevier Inc. All rights reserved.
1. Introduction The electron microscope (EM) was invented in 1931 by German engineers Ernst Ruska (awarded Nobel Prize for this invention in 1986) and Max Knoll. Flourishing during 1950s–1970s, the application of EM in cell and developmental biology resulted in a plethora of discoveries, including descriptions of the ultrastructure of the nucleus (reviewed in [1]) and various cytoplasmic organelles in Xenopus oocytes and embryos ([2,3], reviewed in [4]). Although ultrastructural studies became less popular (and certainly less publishable) with the extraordinary progress and development of molecular techniques, they still remain an indispensable tool for cell and developmental biologists. Therefore, it is worth emphasizing, as has been done most recently by Stenmark [5], how much of our current understanding of cellular and molecular processes we owe to ultrastructural studies. While the introduction of EM-immunocytochemistry allowed us to complement the knowledge gained from ultrastructural and molecular approaches, the use of EM, the number of accomplished electron microscopists who are able to interpret ultrastructural data, the number of people who are adept at electron microscopy, and the number of electron microscope facilities are declining at an alarming rate both in USA and in * Corresponding author. Address: Immuno-Biology Laboratory, The Methodist Hospital Research Institute, 6565 Fannin St., SM 8-034 Houston, TX 77030, USA. Fax: +1 713 793 7151. E-mail address: [email protected] (M. Kloc). 1046-2023/$ - see front matter Ó 2009 Elsevier Inc. All rights reserved. doi:10.1016/j.ymeth.2009.12.003
Europe [6–8]. This is especially unfortunate considering the fact that very often the first clue as to the function of a newly discovered gene comes from the localization pattern of its protein or RNA within cellular organelles or cell/embryo subcellular compartments. In this chapter we describe the methods that enable precise ultrastructural localization and visualization of protein and RNA molecules within the compartments, organelles, and cytoskeleton of Xenopus oocytes.
2. Post-embedding immunogold EM In the immunogold EM technique, the localization of an antigen is revealed indirectly by the application of primary antibody followed by secondary antibody conjugated to colloidal gold (5– 30 nm) or nanogold (0.8–1.4 nm) particles (Figs, 1A–D, 2A). The advantage of using the gold tags lies in their small size, electrondense nature, and strong scattering of electrons, which allow for enhanced contrast between the label and the tissue. In Xenopus oocytes, this technique has been used to study, for example, the formation of germinal granules [9]. The post-embedding immunogold EM procedure can be divided into five steps: (a) (b) (c) (d) (e)
oocyte/ovary fixation, oocyte/ovary embedding, ultrathin sectioning, immunogold labeling of sections, staining/contrasting of immunolabeled sections.
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2.1. Oocyte/ovary fixation Xenopus oocytes or small pieces of the ovaries are fixed in 2–4% formaldehyde, freshly made from paraformaldehyde (Sigma–Aldrich) in 0.1 M Sörensen’s phosphate buffer (pH 7.2–7.4), or, alterna-
tively, 0.5–2% glutaraldehyde (EM grade, Ted Pella, Inc.) in the same buffer. Glutaraldehyde fixation generates better preservation of ultrastructure but usually has a deleterious effect on tissue immunoreactivity. Glutaraldehyde is a strong denaturating agent whose protein cross-linking properties may result in modifications
Fig. 1. (A–D) Post-embedding immunogold EM. A. XVLG (vasa homologue, primary antibody: mouse anti- XVLG, gift from K. Ikenishi, Osaka City University, Osaka, Japan) in nuage material (arrow) attached to the nuclear envelope of the germinal vesicle (gv); c – cytoplasm. B. Accumulations of the nuage material (arrows) labeled with anti-Sm antibody (primary antibody: Y-12, gift from J. Gall, Carnegie Institution for Science, Baltimore, USA). gv – germinal vesicle, c – cytoplasm. (C) Mitochondrial cement between mitochondria (m) labeled with anti-Sm antibody. (D) Nucleolus (nu) in the germinal vesicle labeled with K-121 antibody (Oncogene Research Products, Calbiochem), which recognizes the 5’ terminal 2,2,7-TMG cap of certain small nuclear and nucleolar RNAs. A–D. Secondary antibody conjugated with18 nm gold particles. (A–D) Stage I Xenopus oocytes, see [9] for details. Bars: (A–C) 200 nm, (D) 500 nm.
Fig. 2. (A) Post-embedding immunogold EM. Bundles of intermediate filaments (arrows) in the cytoplasm of the prefollicular cells surrounding young oocytes/cystocytes in the Xenopus ovary labeled with anti-Par3 antibody (Zymed). (B) Extraction method. Fragment of the Triton-extracted oocyte, arrows point to microtubules. (C) Taxol enhanced microtubular cytoskeleton. Electron micrograph of one pole of the mitotic spindle in the dividing cystocyte within the Xenopus ovary. Arrowhead – centriole, arrows – microtubules of the mitotic spindle, asterisks – chromosomes. Bars: (A) 200 nm, (B) 400 nm, (C) 500 nm.
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of the epitope(s) or affect the binding and/or accessibility of the antibody. The choice of fixative and its concentration should be determined for a particular antigen and the corresponding antibody by trial and error. In our experience, fixation of Xenopus oocytes/ovaries in 1% glutaraldehyde or a mixture of 2% formaldehyde and 1% glutaraldehyde works best (depending on the nature of the antigen and antibody). The advantage of using the mixture of fixatives is that formaldehyde penetrates the tissue more rapidly and stabilizes cell structures, which are subsequently fixed more firmly by glutaraldehyde. As a result, a good balance is struck between maintaining cell ultrastructure and, at the same time, retaining high immunoreactivity. In contrast to the standard EM fixation protocol, in immuno-EM there is usually no post-fixation with osmium tetroxide, which may have an adverse effect on antigenicity and may prevent proper UV-induced resin polymerization (e.g., Lowicryl). The fixation step is crucial to the success of the immuno-EM technique. Therefore one should spare no effort in ensuring the structure of the oocytes/ovaries is preserved as closely to its natural state as possible, while keeping antigen properties uncompromised. The viability of an antigen should be tested in a range of conditions (different fixatives, resins, buffers, etc.). It is important to remember that at each step many factors may influence the final result, so keeping an accurate and detailed record of the procedures used is strongly recommended. This should include the fixative type, concentration, and time-course. Because so many variables may affect the final outcome, it is often necessary, especially in the case of previously untested antibodies, to perform preliminary immunocytochemical experiments using light and/or fluorescence microscopy. This enables a large number of variables to be tested in a more efficient, time- and cost-saving manner. If conducted properly, such preliminary experiments can also provide crude information about the distribution of the antigen(s) in the analyzed cell/tissue, which subsequently can be further refined in the samples processed for electron microscopy. 2.2. Oocyte/ovary embedding Xenopus oocytes can be embedded in practically any resin used in EM studies, but Epon 812 or its equivalents [e.g., Poly/Bed 812 (Polysciences, Inc.), EMbed 812 (Electron Microscopy Sciences)], and Lowicryl K4 M (Electron Microscopy Sciences) are used most frequently. A word of caution should be added regarding Epon embedding. This epoxy resin polymerizes best at 60 °C for 64 h, and there are some indications that this may reduce or even completely abolish immunoreactivity. Therefore, for immuno-EM studies it is advisable to conduct the polymerization at a lower temperature (50 °C) and reduce its time to 48 h. In general, acrylic resins such as Lowicryl K4 M and HM20, L.R. White, and L.R. Gold are better suited for immuno-EM as they do not polymerize into three-dimensional crosslinked networks, in contrast to epoxy resins. However, embedding in these resins is more cumbersome because they require anaerobic conditions for polymerization and usually prolong exposure to indirect UV light (360 nm) at low temperature (e.g. Lowicryl K4 M).
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dle than Lowicryl sections. However, prior to antibody labeling, they require gentle etching with sodium hydroxide and/or hydrogen peroxide to remove a thin layer of the resin and to reveal antigenic determinants/epitopes (Section 2.6). 2.4. Immunogold staining of sections The optimal concentrations of primary and secondary antibodies should be determined by trial and error (the useful trial concentration is usually 1:50–1:200 for primary antibodies and 1:200– 1:500 for the secondary ones, see also below). It is good practice, or even necessity, to perform simultaneous control labeling experiments with each localization attempt. Such experiments should be carefully designed to avoid potential artifacts that may lead to false conclusions. The results of labeling experiments should be regarded with a certain degree of skepticism until the results are verified by control experiments. However, it is worth noting that some control experiments are not always necessary or feasible (e.g., adsorption of antibodies with an excess of a studied antigen or with homogenized tissue (ovary/oocytes) containing the antigen), especially if commercial or previously tested antibodies are used. Both negative and positive controls should be utilized whenever possible. The negative control is done by simply omitting the primary antibody while keeping the rest of the labeling procedure unchanged. This permits the estimation of the ‘‘background” level of labeling due to non-specific interactions between the secondary antibody and the cell constituents, the resin, or both. The results of the negative control also facilitate adjustments to the secondary antibody concentration. The positive control, on the other hand, requires the use of a primary antibody that has previously been demonstrated to cross-react with antigen(s) of the given species (in this case X. laevis). By observing tissue labeling in the positive control, one can be more confident that the absence of labeling with a newly tested antibody did not result from technical problems. Another possible control experiment is to use pre-immune serum (or normal serum from a non-immunized animal of the same species in which the antibody was generated) instead of the primary antibody. Secondary antibodies conjugated to gold particles (5–30 nm in diameter) are available from several vendors, including Sigma–Aldrich, Jackson ImmunoResearch Lab., Inc., and Nanoprobes, Inc. It should be mentioned that it is also possible to visualize the distribution of two (or more) antigens in the same section. This procedure is termed ‘‘multiple labeling” and requires the application of secondary antibodies with gold particles of sufficiently different sizes (e.g., 5 and 20 nm). 2.5. Staining/contrasting of immunolabeled sections Because osmium is not used in this procedure (Section 2.1), tissue contrast is often very low. Therefore, staining procedures with Reynold’s solution (lead citrate) and uranium acetate solutions are usually slightly longer than in the standard EM protocol. However, the length of staining depends on the fixation procedure and the resin type used for embedding. Extended staining, even though it improves contrast, may obscure the gold particles.
2.3. Ultrathin section preparation 2.6. Protocol for post-embedding immunogold EM Gray or silver sections (40–70 nm thick) are cut with an ultramicrotome and transferred onto nickel or gold grids (copper grids are not recommended due to increased risk of artifactual precipitates). To prevent grid bars from potentially obscuring places of interest, it is best to use single-slot grids or, alternatively, grids with very thin cross-bars and a high open area. However, singleslot grids require prior coating with formvar (polyvinyl formal; Polysciences, Inc.) or carbon films to give extra support to ultrathin sections. In our experience, Epon sections are easier to cut and han-
(1) Fix the oocytes/ovaries in 2–4% formaldehyde, freshly made from paraformaldehyde in phosphate buffer (pH 7.2–7.4) for up to 1 h at room temperature or in 0.5–2% glutaraldehyde in the same buffer for 30 min; alternatively, use a mixture of 2% formaldehyde and 1% glutaraldehyde in the same buffer for 30 min. (2) Rinse several times in phosphate buffer (usually 3 10 min) at room temperature.
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(3) Dehydrate, infiltrate, and embed the oocytes in the resin of choice (according to the manufacturer’s protocol). (4) Cut sections with an ultramicrotome and collect them on coated/uncoated nickel grids. (5) Etching: incubate1 Epon sections (the etching step is not necessary for the Lowicryl sections) in a drop of 0.25% sodium hydroxide in 100% ethanol for 1–5 min, followed by 3 washes in 100% ethanol. (Hold grid by the edge with fine forceps and dip several times in each of three small glass beakers containing 100% ethanol, or, alternatively, grids can be gently rinsed using a squirting wash bottle.) After rinsing 3 times in distilled water, immerse the grids separately in a drop of 5–10% hydrogen peroxide (freshly made from a concentrated stock solution) for 5–15 min. Rinse 5 times in distilled water.
(6)
(7)
(8) (9)
(10)
(11)
(12)
The incubation of grids can be carried out in small drops of solution placed on a piece of clean dental wax (each grid in a separate drop) or in small depressions made with the cap of an Eppendorf microtube in a piece of Parafilm (Sigma–Aldrich). The grids can be either fully immersed in the solution or placed on the top of the drop (depending on individual preference). Great care should be taken not to damage the film coating the grids with forceps. Block the non-specific binding sites. Incubate the grids in a drop of freshly made 2–5% bovine serum albumin (BSA) in phosphate-buffered saline (PBS) with the addition of 0.05% sodium azide (NaN3) for 20–30 min at room temperature. A similar concentration of non-fat powdered milk or 1–5% fish gelatin (Sigma–Aldrich) in PBS can be used instead of BSA, depending on the type of antibody and the manufacturer’s recommendations. In some instances, it may be beneficial to briefly expose the grids to glycine (50 mM, 10–30 min) to quench free (unreacted) aldehyde groups. Incubate sections in primary antibody diluted in 1% BSA + PBS + NaN3 overnight at 4 °C or, alternatively, for 2– 4 h at room temperature. The optimal antibody concentration can only be found by trial and error. A good starting point is a 1:50 dilution of monoclonal antibodies and 1:200 for polyclonal antisera, but this varies greatly depending on the antibody and the manufacturer. Rinse 5 times in PBS (several dips each). Label the sections with secondary antibody complexed to colloidal gold particles of required size and diluted in BSA + PBS + NaN3 for 2 h at room temperature. Care should be taken to choose a secondary antibody that is fully compatible with the primary antibody. Again, the optimal antibody dilution should be discovered by trial and error; one can start with a 1:200 dilution. Rinse twice in PBS followed by three washes in distilled water. At this point, leave the grids out to dry or proceed with contrasting. Contrast sections with uranium acetate and lead citrate according to standard EM procedures. The optimum time for each contrasting step has to be established empirically. Finally, after rinsing several times with distilled water and drying, sections are now ready for viewing in TEM.
oocytes. This is due primarily to the presence of numerous cytoplasmic organelles, nutrient/reserve materials accumulated in yolk granules, and lipid droplets that mask the cytoskeletal components/filaments. One way to overcome this difficulty is to remove the ‘‘unwanted” cell constituents with detergent (Triton X-100) extraction. We have adopted a gentle extraction method, originally developed for insect oocytes [10], to Xenopus oocytes. This method is particularly suitable for visualization and analysis of microtubule distribution (Fig. 2B). The most critical step in this method is the Triton extraction. Care should be taken not to prolong this step, as this may result in complete destruction of the cytoskeletal structures. The whole procedure requires 12–13 steps, including three fixation steps: (1) Extraction. (2) Fixation in fixative 1, 15–30 min at room temperature. (3) Rinsing in 0.1 M phosphate buffer pH 7.4, 3 15 min at room temperature. (4) Fixation in fixative 2, 1–1.5 h at 4 °C. (5) Rinsing in 0.1 M phosphate buffer pH 7.4, 3 30 min at room temperature. (6) Fixation in fixative 3, 1–1.5 h at room temperature. (7) Rinsing in distilled water, 3 30 min at room temperature. (8) En bloc staining in 2% aqueous uranyl acetate for 2 h or overnight at room temperature. (9) Rinsing in tap water, 3 5 min. (10) Dehydration in increasing concentrations of ethanol (30% – 2 10 min, 50% – 2 10 min, 70% – 3 10 min, 96% – 3 15 min, 100% – 3 30 min) and in acetone (3 10 min); all steps at room temperature. (11) Embedding in Epon 812 or its equivalents (e.g. Poly/Bed 812 from Polysciences, Inc., EMbed 812 from Electron Microscopy Sciences) according to the manufacturer’s protocol. (12) Cutting block into ultrathin sections. (13) Additional section contrasting (this step is optional). 3.1.1. Extraction buffer Defolliculated Xenopus oocytes or small pieces of ovaries are immersed in the following extraction buffer:
137 mM NaCl 5 mM KCl 1.1 mM Na2HPO4 0.4 mM KH2PO4 4 mM NaHCO3 12 mM MgCl2 2 mM EGTA 50 mM 2-(N-morpholino) ethanesulfonic acid (MES) 5.5 mM glucose 1% Triton X-100
(all chemicals available from Sigma–Aldrich, St. Louis, MO, USA). Extraction time: 5–15 min, depending on the specimen size. It is worthwhile to extract 2–3 bunches of oocytes at 5-min intervals to determine the optimal timing.
3. Methods for cytoskeleton visualization 3.1.2. Fixative 1 3.1. Extraction method 2.5% glutaraldehyde in the extraction buffer. It is challenging to analyze cytoskeleton distribution and spatial organization at the ultrastructural level in cells as large as Xenopus
3.1.3. Fixative 2
1
It is recommended to do this step and the following incubation of sections with the antibodies in a moist chamber; one can be easily prepared using a covered Petri dish lined with moist filter paper.
2% osmium tetroxide, 0.8% potassium ferrocyanide,
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in 0.1 M phosphate buffer pH 7.4. The mixture should be prepared just before use and kept at 4 °C. 3.1.4. Fixative 3 0.15% tannic acid in 0.1 M phosphate buffer pH 7.4. 3.2. Fixation with Taxol Another method of improving visualization of the microtubule cytoskeleton is the addition of taxol to the fixative. Taxol binds tightly to microtubules and stabilizes them in cells and in vitro [11,12]. As no osmium tetroxide is used during the fixation step, the cell membranous structures are only lightly stained, while microtubules and centrioles are highly contrasted/enhanced (Fig. 2C). This method has been used successfully to study the organization of Xenopus germline cysts [13].
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3.3.1. Microtubule stabilizing fixative 2% formaldehyde, 3% glutaraldehyde (EM grade, Ted Pella, Redding, CA), in 0.1 M sodium cacodylate buffer pH 7.3 (Polysciences, Warrington, PA), 10 lM taxol (Cytoskeleton, Denver, CO). 4. Whole-mount (pre-embedding) EM immunostaining of Xenopus oocytes The pre-embedding (whole-mount) immunostaining technique is much faster than performing immunostaining on sections. The only potential limitation is the permeability of the tissue. Thus, this method is great for staining the isolated oocytes and the surface layers of the whole embryo, but it may fail to label deeper embryonic layers (Fig. 3A, C–E). 4.1. Fixation
3.3. Fixation to improve microtubule visualization Fix oocytes in microtubule stabilizing fixative for 30 min at room temperature. Wash in cacodylate buffer 3 10 min. Stain in 0.5% uranyl acetate for 30–60 min. Dehydration and embedding according to the standard EM protocol.
Manually or enzymatically (with collagenase, cat # C6885, Sigma–Aldrich, St. Louis, MO, USA) defolliculated stages-I–VI oocytes are fixed as follows: 4.1.1. For non-cytoskeletal protein immunostaining Oocytes are fixed in 2% formaldehyde plus 0.5% glutaraldehyde (both-EM grade from Ted Pella Inc., Redding, CA, USA:
Fig. 3. Whole mount (pre-embedding) nanogold immunostaining and in situ hybridization. (A) Fatvg protein is present on the surface of vesicles (v) in the stage III Xenopus oocyte (see [17] for details), y – yolk granule. Secondary antibody conjugated with 1.4 nm gold. (B) In situ hybridization, Xcat2 mRNA (arrows) is present within the germinal granule; m – mitochondria. Anti-digoxigenin antibody, conjugated with 0.8 nm gold, and silver enhanced. (C–E) Oocytes labeled with anti-cytokeratin C11 antibody (Sigma– Aldrich) and nanogold-conjugated secondary antibody, and silver enhanced (see [15] for details). Arrows indicate cytokeratin filaments distributed in the vegetal cortex of the stage VI Xenopus oocyte, y – yolk granule. (C) Low magnification view of the fragment of the vegetal cortex of labeled oocyte. Semithin section (0.7 lm) stained with 1% methylene blue in 1% borax. (D and E) Ultrathin sections stained with uranyl acetate and lead citrate. Bars: (A and B) 500 nm, (C) 5 lm, (D and E) 500 nm.
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glutaraldehyde 8% stock, cat# 18421; formaldehyde 16% stock, cat # 18505) in 1 phosphate-buffered saline (PBS) in moleculargrade water for 2 h at room temperature or overnight at 4 °C. Using this concentration of glutaraldehyde is a trade-off between the relatively good preservation of ultrastructure and higher immunoreactivity. Increases in the glutaraldehyde concentration (within a 1–3% range) will improve ultrastructure but will usually decrease immunoreactivity (Fig. 3A) [14]. 4.1.2. For cytokeratin immunostaining Oocytes are fixed in 1–4% formaldehyde (either EM-grade from Ted Pella, Inc. or from 37% formaldehyde stock, cat # F-1635 from Sigma–Aldrich, St. Louis, MO, USA) in 100% methanol (GC resolve grade, cat # A457 from Fisher Scientific, Pittsburgh, PA, USA) overnight at 20 °C. The preservation of the ultrastructure of the components of the oocyte and of the cytokeratin epitope are identical in samples fixed with formaldehyde concentrations ranging from 1% to 4% [15]. 4.2. Post-fixation washes Fixed oocytes are rehydrated and washed with slow shaking on a rotating platform as follows: 100% ethanol 2 10 min, 70% ethanol 15 min, 50% ethanol 15 min, 2 5 min washes in phosphatebuffered saline (PBS) containing 0.05% Tween 20 (cat #166–2404, Bio-Rad Laboratories, Hercules, CA), 10 min in PBS–0.25% Triton X-100 (cat # X100, Sigma–Aldrich, St. Louis, MO, USA), and 2 5 min with PBS–0.05% Tween 20. 4.3. Blocking, antibody incubation, and washing All steps are performed with slow shaking on a rotating platform. Non-specific binding is blocked by incubating 6 h in casein blocking buffer (cat #161–0782, Bio-Rad Laboratories, Hercules, CA) with 0.05% Tween 20 at room temperature. This is followed by incubation with the appropriate primary antibody. For cytokeratin immunostaining, oocytes are incubated with anti-pan cytokeratin clone C-11 mouse monoclonal antibody (cat # C293, Sigma– Aldrich, St. Louis, MO, USA). All primary antibodies should be used at 1:50 dilution in casein blocking buffer containing 0.05% Tween 20, overnight at 4 °C. Oocytes are then washed 4–6 h at room temperature or overnight at 4 °C with PBS–0.05% Tween 20. The overnight washing significantly decreases the background. After washing, oocytes are incubated overnight at 4 °C with secondary antibody (appropriate for the given primary antibody, i.e., anti-mouse, anti-rabbit, etc.) conjugated to 1.4 nm gold (Nanoprobes, Yaphank, NY, USA) and diluted to 1:50 with casein blocking buffer containing 0.05% Tween 20. The next day, oocytes are washed 3 15 min with PBS–0.05% Tween 20, post-fixed for 20 min in 1% glutaraldehyde in PBS– 0.05% Tween 20, washed 3 15 min in molecular-grade water, and silver- or gold-enhanced.
enhancement, oocytes are washed 3 10 min in molecular-grade water and then post-fixed in fixative that contains 2% glutaraldehyde (EM-grade from Ted Pella, Inc., Redding, CA, USA) in 1 PBS. Samples can be kept in this fixative at 4 °C for several days before embedding. 4.5. Embedding and sectioning Whole-mount nanogold-immunostained and silver- or gold-enhanced oocytes are contrasted with 0.5% uranyl acetate only (without osmium tetraoxide, which damages the metal label) and, after dehydration in increasing concentrations of ethanol, infiltrated and embedded in LX-112 medium (Epon substitute; Ladd Research Industries, Burlington, VT, USA). The samples are polymerized in a 70 °C oven for 2 days. Semithin sections (0.7–1 lm) are stained with 1% methylene blue in 1% borax (Fig. 3C). Ultrathin sections (70–100 nm) are cut, stained for 5 min in 1% aqueous uranyl acetate, and 2 min in 1% aqueous lead citrate at room temperature in a Leica EM Stainer (Leica, Deerfield, IL), and examined in a JEM 1010 transmission electron microscope (JEOL, USA, Inc., Peabody, MA) at an accelerating voltage of 80 kV (Fig. 3D–E). Digital images are obtained using AMT Imaging System (Advanced Microscopy Techniques Corp, Danvers, MA). 5. Whole-mount (pre-embedding) EM in situ hybridization of Xenopus oocytes 5.1. Fixation Fixative for electron microscopy in situ hybridization of oocytes (identical to that used for Xenopus embryos and described in Kloc et al. [16]) contains 4% formaldehyde, 0.1% glutaraldehyde (both EM-grade from Ted Pella), 100 mM KCl, 3 mM MgCl2, 10 mM HEPES, 150 mM sucrose, and 0.1% Triton X-100, with a pH of 7.6. The main difference between this fixative and the fixative used for light microscopy in situ hybridization is the presence of glutaraldehyde, MgCl2, and sucrose, which have a stabilizing effect on the membranes and organelles. Defolliculated oocytes are fixed at room temperature for 1–2 h with shaking. 5.2. Post-fixation washes All subsequent treatments are performed with shaking. Fixed oocytes are washed 3 5 min with PBS–0.1% Tween 20 and then treated for 10 min in 10 mg/ml proteinase K in PBS–0.01% Tween 20. This step facilitates the penetration of the probe through the cortical layer of the oocyte. After proteinase K treatment, oocytes are washed for 5 min in 0.1 M triethanolamine (TEA) pH 8.5, then incubated for 10 min in 0.1 M TEA with 0.5% acetic anhydride, washed 3 5 min in PBS-Tween and post-fixed for 20 min in the fixative described above. After post-fixation oocytes are washed 3 5 min in PBS-Tween, and then hybridized overnight at 50 °C.
4.4. Silver or gold enhancement 5.3. Hybridization and post-hybridization washes Silver enhancement is performed with Light Insensitive Silver Enhancer (cat # 2013) and gold enhancement with GoldEnhance (cat# 2113) (both from Nanoprobes, Yaphank, NY, USA), according to the manufacturer’s protocol, for approximately 45 min. Both procedures are done without shaking. Although the general belief is that gold enhancement gives much lower background than silver enhancement, in our hands both types of enhancement work comparably well with very low background. To prevent over-staining, the development of the signal is monitored under the dissecting microscope and stopped (by transferring to molecular-grade water wash) when the oocytes reach a light-brown color. After the
Hybridization and all post-hybridization washes are performed exactly as described previously for light microscopy in situ hybridization [17]. In this citation we also gave a comprehensive list of all chemicals and solutions needed to perform in situ hybridization. 5.4. Antibody incubation After the last post-hybridization wash in PBS-Tween, oocytes are incubated for 15 min in G1 buffer (Boehringer Mannheim/ Roche) and then incubated overnight at 4 °C with a 1:30 dilution
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of anti-DIG 0.8 nm gold (Boehringer Mannheim/Roche) in G2 buffer. As far as we know, Boehringer Mannheim/Roche stopped producing anti-DIG 0.8 nm gold antibody. Although we never tried a different antibody, we believe that this antibody can be successfully replaced with any available anti-digoxigenin antibody conjugated with nanogold (0.8–1 nm) or 5-nm gold (using gold larger than 5 nm will probably create penetration problems). 5.5. Silver or gold enhancement After incubation with the antibody, oocytes are washed 415 min in PBS-Tween, post-fixed in 2% glutaraldehyde in PBSTween, and washed 3 15 min in molecular-grade water. Subsequently, they are silver- or gold-enhanced in Light Insensitive Silver Enhancer or GlodEnhance (Nanoprobes Inc, NY) for approximately 45 min. To prevent over-staining, the development of the signal should be monitored under a dissecting microscope. After the enhancement, oocytes are washed 3 10 min in molecular-grade water and then post-fixed in fixative that contains 4% formaldehyde, 2% glutaraldehyde (both EM-grade from Ted Pella), 100 mM KCl, 3 mM MgCl2, 10 mM HEPES, 150 mM sucrose, and 0.1% Triton X-100, pH 7.6. Oocytes can be kept in this fixative for several days before embedding. 5.6. Embedding and sectioning Fixed oocytes are stained in 0.5% uranyl acetate without osmium tetraoxide treatment, dehydrated in increasing concentrations of ethanol, and infiltrated and embedded in LX-112 medium (Epon substitute). Ultrathin sections (70–100 nm) are stained for 5 min in 1% aqueous uranyl acetate and 2 min in 1% aqueous lead citrate at room temperature and examined in a transmission electron microscope (Fig. 3B). 6. Digital visualization of Xenopus oocyte/organelles 6.1. Introduction Digital electron microscopy offers remarkable quantitative and qualitative advantages derived from computational visualization, such as a more nuanced assessment of Xenopus threedimensional (3D) structure. Prior to digital microscopy, noncomputational techniques used photographic film to capture the two-dimensional (2D) image projections of stained specimen sections. Modern techniques allow for direct image capture using CCDs or high-quality image scanning of photomicrographs. Due to the complexity of cellular structure and the inherent limitations of 2D imagery, it was difficult for microscopists to convey their remarkable understanding of three-dimensional structure prior to the advent of digital microscopy. Additionally, the management of such photomicrographs made collaboration difficult because of logistical limitations in organizing, indexing, and disseminating these 2D images. Significant technical advances in the design and manufacture of electron microscopes, most notably digital image acquisition and computer control of instrumentation, have allowed for sectional microscopy and electron tomography techniques. These methods provide greater insight into 3D cellular structure by establishing the distribution of lipids, proteins, and RNAs within the cytoplasm (Fig. 4). The ability to digitally assemble a sectional stack of 2D images allows for a greater understanding of cellular arrangement through interactive 3D visualization. Moreover, the application of computed axial tomography in electron microscopy allows for even finer 3D ultrastructure (2–10 nm resolution). At these resolutions, methods of specimen preparation become critical because of
Fig. 4. (A) Three-dimensional reconstruction. The mitochondrial cloud (light blue) extending between the nucleus (yellow) and the vegetal cortex contains germinal granules (red) at its vegetal tip (the METRO region). The nucleus is surrounded by mitochondria (dark blue). See [19] for details. (B) Three-dimensional reconstruction of a germinal granule showing the distribution of Xcat2 mRNA (green dots). See [18] for details. (C) Three-dimensional reconstruction of the Xenopus female germline cyst; three different views of the same cyst. Cytoplasm is gray, nuclei are red, mitochondria of primary mitochondrial cloud are green, centrioles blue, and ring canals are yellow. See [13] for details.
the likelihood of introducing artifacts that corrupt biological structure. For example, the use of heavy-metal stains frequently induces protein cross-linking that causes the creation of artificial protein
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complexes or lipid membrane destruction, which are observable at high resolution. Electron tomography repeatedly irradiates a small area of the specimen from a number of angles, requiring close monitoring of radiation damage at the time of 2D image acquisition. These raw tomographic tilt-series projections are computationally transformed by 3D reconstruction algorithms (e.g., direct Fourier inversion, filtered back projection, iterative algebraic). Each image is assembled as a function of the Euler angles of the projection, resulting in the 3D spatial image. A notable instrument limitation is the inability to fully rotate the goniometric specimen stage 90°, resulting in a ‘‘missing cone” of data that reduces the 3D resolution and produces smearing artifacts along the optical axis. Generally, specimens are no thicker than 500 nm, but this is a function of the operating voltage of the electron microscope and the composition of the specimen. Thinner specimens allow for greater tilt angles, and dual-axis tilt specimen stages can be used to improve resolution. Corresponding technical advancements in rapid cryo-freezing provide significant benefits to researchers who require the highest resolution and fidelity; in such cases, unstained cryo-hydrated specimens should be used. Such methods produce specimens that are highly correlated to in vivo samples because the speed of freezing essentially halts molecular and atomic activity, providing in situ snapshots of biological dynamics. These quickly frozen atomic positions present accurate glimpses of the chemical interactions and signaling within the cell. Another important advantage of cryo-microscopy is that these low-temperature specimens are better able to tolerate radiation damage caused by the electron beam; this is critical in the use of electron tomography because the same specimen region is irradiated numerous times. A number of hybrid approaches can be used, such as sectional cryo-tomography, freeze substitution, and cryo-negative staining. Researchers should familiarize themselves with the various methods and associated limitations during the development of a project plan, choosing an optimal set of protocols that will provide reproducible results in an efficient manner. The use of stains or antibodies at low resolution frequently provides quick satisfactory results that can be extended to higher-resolution methods. 6.2. Methods to obtain three-dimensional images A number of methods exist for researchers to obtain 3D imagery, the easiest being the Internet. The recent establishment of the Electron Microscopy Data Bank (EMDB, http://emdatabank.org) enables researchers to download an increasing number of 3D images of macromolecular complexes and cells. EMDB was established by the Protein Databank in Europe (PDBe) at the European Bioinformatics Institute, the Research Collaboratory for Structural Bioinformatics (RCSB) at Rutgers, and the National Center for Macromolecular Imaging (NCMI) at Baylor College of Medicine. EMDB provides a global deposition and retrieval network for cryo-EM 3D images, models, and associated metadata. Additionally, EMDB is a portal for software tools for standardized map format conversion; map, segmentation, and model assessment; visualization; and data integration. In tandem, a number of scientific journals are requiring the deposition of 3D imagery into open-access repositories as a condition of publication; this allows global communities of researchers to personally study the digital specimens, apply new computational techniques for verification, and initiate new tangential avenues of investigation, further enhancing the value of the initially funded research. The data format of choice is MRC2000, developed in 1982; it is the oldest and most widely used 3D electron microscopy image format. When the desired imagery does not exist at EMDB, researchers may contact global centers that explicitly provide these services,
requiring the researcher to vet and prepare specimens to be imaged. The National Center for Research Resources (NCRR, http:// ncrr.nih.gov) provides laboratory scientists and clinical researchers with the tools and training needed for research. NCRR funds a large number of biomedical technology research centers, such as the National Center for Macromolecular Imaging (http://ncmi.bcm.edu), specializing in high resolution cryo-electron microscopy and electron tomography; the National Center for Microscopy and Imaging Research (http://ncmir.ucsd.edu), specializing in cellular electron and optical microscopy; and the Resource for the Visualization of Biological Complexity (http://www.wadsworth.org/rvbc/), specializing in sectional time-resolved cryo-electron tomography. Similar centers exist in Europe, Japan, and China. Researchers are encouraged to contact the centers directly to determine the suitability of a center’s skills and resources to the researcher’s objectives and needs. Frequently the centers can accommodate a researcher’s skill set; for example, an experienced microscopist might allow a researcher direct access to the facilities, or a researcher who has no electron microscopy experience could merely provide the specimen to the center and outline any unique quality-control issues in handling their specimen. Depending on the frequency of specimen production and on quality control, it may be desirable to purchase a plunge-freezing apparatus and ship cryogenically frozen specimens to the center on electron microscopy grids. After microscopic image acquisition, the center can usually provide the researcher raw electron microscopic data, assist the researcher in using 2D-to-3D image reconstruction tools, or generate a final 3D structure that could be uploaded to the EMDB. For researchers and institutions considering the purchase of an electron microscope, these centers can also provide valuable consultation on implementation. For researchers who prefer to perform the 3D image reconstruction, there are a number of software tools available. Some software programs are integrated with the instrument, such as with FEI electron microscopes. There are commercial scientific visualization software products that may be purchased, such as Amira (http:// www.amiravis.com). Amira has some remedial tools designed for confocal microscopy that can be adapted to align sectional electron micrographs. Alternatively, NCRR-funded open-source software tools such as IMOD (http://bio3d.colorado.edu), developed by the Boulder Lab for 3D Electron Microscopy, are used throughout the structural biology community because of Boulder’s rigorous research and development at the forefront of electron tomography. The IMOD package is able to reconstruct serial sections or electron tomograms. It has sophisticated features, such as compensation for specimen deformation due to microtomy shearing and automated detection of gold particles used as fiducial markers in electron tomography. The Particle Estimation for Electron Tomography (PEET, http://bio3d.colorado.edu) package provides alignment and averaging of subvolumes extracted from tomograms, useful in studying identical sub-cellular structures such as the nuclear pore complex. 6.3. Methods of segmenting three-dimensional images Segmentation is a labor-intensive process requiring the researcher to be actively involved in annotating the boundaries between regions of interests. No comprehensive automated system exists at this time, but research is ongoing to develop reliable 3D pattern recognition. An inherent problem is automatically identifying components that have never been structurally observed before; this is further compounded by the interaction of multiple proteins in novel or poorly understood combinations. Low signal-to-noise ratios are another important limitation. The Amira, Shape, and Seg3D software packages are frequently used for segmentation of data sets, but other methods exist, such as The Watershed
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Transformation, eigenvector techniques, and multiphase level set segmentation. Amira, being primarily a generic visualization package, offers a number of software tools to analyze and dissect 3D images. Most important are the image filters, image segmentation editor, isosurfacing, and volume rendering. Image filters can be applied to the 3D reconstructed image prior to segmentation, allowing for greater contrast when annotating the boundaries. For tomograms, median and Gaussian filters are used to reduce the noise and minimize the missing-cone artifacts. The generic segmentation editor within Amira allows a researcher to manually go through the 3D image slice-by-slice along a preferred orthogonal axis. The researcher must make critical decisions about the annotation of biological regions of interest (ROI). The segmentation editor can be utilized with a mouse, but it is highly recommended that a graphics tablet and pen be used; the pen permits hand control that is more accurate, with significantly faster throughput. The editor has a number of segmentation tools, the most basic being Brush, similar to the brush tool in Adobe Photoshop. Other tools such as Magic Wand and Blowtool can significantly expedite annotation but require considerable effort to master because these tools are highly sensitive to image composition, particularly noise. The segmentation editor allows for the annotation of 256 ‘‘materials,” and multiple editors can be applied simultaneously to the same dataset. Shape (http://bio3d.colorado.edu) is an open-source package designed for automated segmentation of tomograms. It provides for denoising using 3D median filtering, extraction of features using Line Filter Transform and Orientation Filter Transform, and the extraction of contours along thin structures. Its major limitations involve datasets with large numbers of ribosomes, low-contrast filters, and microtubules that are out of plane. Researchers frequently use multiple packages, combining the best features of each by translating intermediate 3D images and segmented subvolumes between the packages. Although automated methods can significantly enhance the quality and speed of segmentation, ultimately the researcher must provide the final quality control either visually and/or manually. Seg3D (http://seg3d.org) is an open-source package developed by the University of Utah’s Center for Integrative Biomedical Computing, funded by the NCRR. Among its many features, it implements the National Library of Medicine’s Insight Tool Kit (ITK, http://itk.org), an advanced algorithmic software library developed to segment the Visible Human. Seg3D integrates automatic and manual features and at the same time is capable of managing very large images.
6.4. Methods to visualize three-dimensional images There are a number of software tools to visualize datasets using two methods of graphics: isosurfacing and volume rendering. Isosurfacing creates 3D polygons by contouring an image using a common contour threshold. These polygons can be individually colored with a 3D color field and by mapping a color-lookup table (CLUT) onto the color field. This can provide additional visual cues, such as radial distance or the highlighting of specific structures. Multiple isosurfaces may be displayed, each having independent thresholds, either on the same dataset or separate datasets. Volume rendering computes a 2D projection of a 3D image as a function of camera position and the 3D image voxels; the voxels are
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defined from 3D images convolved with a color transfer function, typically a red-green-blue transparency (CLUT). The primary computational limitations to visualization are the size of the 3D image, followed by the graphical computing power of the hardware. For example, if the computer memory is 4 gigabytes and the 3D image is 500 gigabytes, then visualization may not be possible without advanced computational techniques and resources; for the majority of datasets, the 3D image size is less than computer memory. Also, if the 3D image contains a significant amount of details or noise, the number of polygons may increase to the point that the graphics become sluggish or potentially cause an operating system failure. The use of isosurfacing provides convenience at computational expense, and the use of volume rendering can reduce graphical overload. Although there is a large number of visualization software packages, a few are frequently used by the electron microscopy community: Amira, Chimera, and SciRun. Chimera (http:// www.cgl.ucsf.edu/chimera/) is an open-source molecular-microscopy visualization tool developed by the Resource for Biocomputing, Visualization, and Informatics and is funded by NCRR. SciRun (http://scirun.org) is an open-source package developed by the University of Utah’s Center for Integrative Biomedical Computing, also funded by the NCRR. All three packages were developed in academic settings and have numerous technical features. Each offers a number of tutorials to demonstrate basic capabilities and illustrate methods of constructing highly complex visualizations and animations. The examples of 3D reconstruction of Xenopus oocytes and organelles are shown in Fig. 4. Acknowledgments We are grateful to Mrs. W. Jankowska and B. Szymanska for excellent technical support and Ms. E. Kisiel for helping with the figures. Panel A in Fig. 4 was reproduced with the permission of the publisher from the Fig. 4 panel A in the article published in Int. J. Dev. Biol. 48 (2004) 17-21. References [1] [2] [3] [4] [5] [6] [7] [8] [9] [10] [11] [12] [13] [14] [15] [16] [17]
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