Hemisecting Xenopus Oocytes and Eggs

The large size of Xenopus oocytes and eggs hampers the penetration of antibodies, particularly after fixation with aldehydes. To aid antibody penetration, we routinely

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Fig. 2. Triple-fluorescence labeling of mitotic and meiotic spindles in Xenopus oocytes and embryos. Oocytes (A-C) and blastula-stage embryos (D—F) were fixed in methanol as described and stained with rabbit anti-XKCMl and Alexa 488-conjugated goat antirabbit immunoglobulin G (A and D), rat anti-a-tubulin (YL1/2) and Alexa 546 goat antirat immunoglobulin G (B and E), and TO-PRO-3 (C and F). Three channel images were collected with a Zeiss LSM-510 and were merged in Photoshop. Scale bar is 10 |lm.

Fig. 2. Triple-fluorescence labeling of mitotic and meiotic spindles in Xenopus oocytes and embryos. Oocytes (A-C) and blastula-stage embryos (D—F) were fixed in methanol as described and stained with rabbit anti-XKCMl and Alexa 488-conjugated goat antirabbit immunoglobulin G (A and D), rat anti-a-tubulin (YL1/2) and Alexa 546 goat antirat immunoglobulin G (B and E), and TO-PRO-3 (C and F). Three channel images were collected with a Zeiss LSM-510 and were merged in Photoshop. Scale bar is 10 |lm.

"hemisect" oocytes with a sharp scalpel prior to bleaching or staining. We cut oocytes and eggs either equatorially to observe the animal or vegetal cortex or laterally along the animal-vegetal axis. When properly mounted, hemisected oocytes and eggs also allow visualization of regions of interest deep within the oocytes or eggs that would otherwise be inaccessible with high numerical aperture objectives (see Subheading 3.6.). In addition, hemisecting allows separate processing of the two oocyte or egg halves with different antibodies. In our experience, it is best to change the scalpel blade often to ensure as smooth a cut surface as possible. Although tedious, we have hemisected oocytes as small as approx 400 |im in diameter. Smaller oocytes (<200 |im) are processed whole, and antibodies generally diffuse throughout the cytoplasm, allowing optical sectioning of the complete cytoplasmic volume (6,8,12,14).

The procedure is as follows:

1. Rehydrate/wash samples with either TBS or TBSN.

2. Transfer oocytes to a Petri dish or glass agglutination slide with a large amount of buffer surrounding the oocytes to reduce movements caused by surface tension.

3. Hemisect oocytes or eggs either in the plane of the equator or along the animal-vegetal axis.

4. After cutting, return oocytes to microcentrifuge tubes filled with TBS or TBSN. If desired, transfer the halves (animal vs vegetal or right vs left) to different tubes for processing.

3.3. Bleaching of Xenopus Oocytes and Eggs

The cortical pigment of stages IV to VI Xenopus oocytes and eggs attenuates laser illumination and obscures fluorescence from the underlying cytoplasm. Pigmentation of fixed Xenopus oocytes and eggs can be reduced or eliminated by bleaching them with 10% H2O2 in methanol (1 part 30% H2O2/2 parts 100% methanol; see Note 5) (21). The animal and vegetal hemispheres of bleached Xenopus oocytes and eggs are almost indistinguishable. If desired, oocytes should be hemisected prior to bleaching. Peroxide bleach is very reactive with clothing and causes painful chemical "burns" on contact with skin. Gloves should be worn and care taken when using bleach.

Oocytes and eggs from albino frogs (supplied by most distributors of Xenopus) can be used when the antibodies/epitopes are incompatible with peroxide bleach or methanol. However, distinguishing the animal and vegetal axis of albino oocytes and eggs can be very difficult.

The procedure is as follows:

1. Carefully aspirate the TBSN or TBS.

2. Bleach samples by adding approx 1.5 mL of 10% H2O2 in methanol.

3. Incubate samples with the tubes on their sides for 12 to 48 h at room temperature under fluorescent illumination.

4. Carefully aspirate the bleaching solution and rinse samples three times (~1 h each) with TBSN (or TBS if NaBH4 reduction is required).

3.4. Borohydride Reduction of Glutaraldehyde-Fixed Oocytes and Eggs

Xenopus oocytes and eggs that have been fixed with glutaraldehyde should be treated with sodium borohydride to reduce unreactive aldehydes and the autofluorescence of Schiff bases generated during fixation (22). Borohydride reduction is unnecessary for samples fixed in methanol or FT. We typically treat FGT-fixed oocytes with 50 mM NaBH4 in TBS overnight at 4°C or 100 mM for 4 to 6 h at room temperature. Borohydride solutions are effervescent, and the buffers used should lack all detergents. In addition, tubes or vials should be left open and upright during this treatment to prevent pressure buildup that may result in the caps popping open, which can send the tubes flying.

For the procedure,

Rinse samples three times with TBS to remove bleach or detergent from previous steps. Aspirate off the TBS.

Add approx 1 mL of 50 or 100 mM NaBH4 in TBS.

Incubate either overnight at 4°C or 4 to 6 h at room temperature in uncapped tubes. Add 1 to 2 drops of TBSN to break the surface tension of the bubbles formed from NaBH4. Tap tubes to help break the surface tension and allow the oocytes and eggs to sink to the bottom.

Carefully aspirate off the overlying NaBH4 solution.

Wash samples in several changes of TBSN for 1 to 3 h at room temperature.

3.5. Processing of Xenopus Oocytes and Eggs for Immunofluorescence

The large size of Xenopus oocytes and eggs limits penetration of antibodies to approx 75 to 100 |im during an overnight incubation. Therefore, processing Xenopus oocytes and eggs for immunofluorescence requires that the incubation of antibodies be substantially increased compared to cultured cells or sections, with corresponding increase in the duration of intermediate washes. In our experience, antibody incubation times of 24 to 48 h and washes of 48 to 72 h are satisfactory for hemisected oocytes and eggs.

We have used a number of antibodies against cytoskeletal proteins with varying degrees of success (for a complete list, see Table 1 and ref. 15). The appropriate fixation conditions and the working dilutions for each primary and secondary antibody must be determined empirically. We typically start with a 1/100 dilution of the stock provided in TBSN + 2% BSA (a final concentration of 10 to 50 |g/|L) and adjust the dilution to minimize background or increase brightness. NaN3 can be added to 0.1% to retard bacterial growth during storage at 4°C (see Note 6). In our experience, we have found that preblocking Xenopus oocytes and eggs is not required. In some cases, we have reused diluted primary antibodies several times, observing that the background or nonspecific staining is reduced in subsequent uses.

The choice of a fluorescently conjugated secondary antibody must satisfy several criteria: The fluorochrome must match the excitation wavelengths available, and the fluorochrome must be sufficiently photostable to allow the collection of many optical sections, including frame averaging. Moreover, fluorochromes used for dual or triple labeling must be spectrally separable (either the absorption or emission spectra; see Fig. 2 and Note 6).

Our early studies relied heavily on Texas Red-, tetramethylrhodamine-, and fluo-rescein-conjugated antibodies for both single- and multiple-labeling strategies (see Table 2). More recently, we have used Alexa-conjugated secondary antibodies available from Molecular Probes Inc. (Eugene, OR) with excellent results. The excitation and emission spectra of Alexa 488 and Alexa 546 are readily separable with filters available on most common confocal microscopes (consult the specifications of your confocal to determine which excitation/emission wavelengths are compatible), and the Alexa dyes exhibit excellent photostability.

Many dyes are available for counterstaining nuclei and chromosomes. Hoechst and 4,"6-Diamidino-2-phenyl indole (DAPI), commonly used to stain nuclei and chromosomes for conventional fluorescence microscopy, are not compatible with laser lines in most confocal laser scanning microscopes (they can be used with instruments capable of multiphoton excitation). We have used propidium iodide, ethidium homodimer, YO-PRO-1, BO-PRO-3, and TO-PRO-3 with good results (see Table 2). Propidium iodide and ethidium homodimer work best on oocytes and eggs fixed with methanol. The cyanine dyes YO-PRO-1, BO-PRO-3, and TO-PRO-3 work well with aldehyde-fixed samples. The spectrum of TO-PRO-3 is readily separated from both Alexa 488 and Alexa 546, allowing triple-fluorescence labeling of two antibodies and chromosomes/nuclei (see Fig. 2 and ref. 1).

The protocol outlined here is used for triple labeling with two antibodies and the far-red chromosome dye TO-PRO-3. For single- or double-labeling strategies, omit the appropriate steps. This protocol has been adapted from refs. 1, 2,11, and 15:

1. Fix, hemisect, bleach, and reduce samples with NaBH4 as desired in 1.7-mL microcentrifuge tubes (see above).

2. Wash samples with several changes of TBSN over 4 to 6 h.

3. Incubate samples with 100 to 200 |L of the first primary antibody (diluted in TBSN + 2% BSA) for 24 to 48 h at 4°C with gentle rotation using a rotary mixer (see Note 7).

4. Wash samples with TBSN at 4°C with gentle rotation for 48 to 72 h, changing the buffer every 8 to 12 h (see Note 8).

5. For dual fluorescence, incubate samples with 100 to 200 | L of the second primary antibody (diluted in TBSN + 2% BSA) for 24 to 48 h at 4°C with gentle rotation using a rotary mixer (see Note 7).

6. Wash samples as described in step 4.

7. Incubate samples with 100 to 200 | L fluorescent-conjugated secondary antibodies (diluted in TBSN + 2% BSA) at 4°C for 24 to 48 h with gentle rotation using the rotary mixer (see Note 9 and Table 2).

8. Wash samples as in step 4.

9. If staining for chromosomes with TO-PRO-3, wash samples in TBS for 1 to 2 h with three or four changes at room temperature.

10. Stain samples with approx 1 mL of 5 |M TO-PRO-3 in TBS for 30 to 60 min at room temperature with gentle rotation on a nutator or orbital shaker with samples lying flat (cover samples to keep dark after this step).

11. Wash samples two or three times overnight with TBS at 4°C with gentle rotation.

3.6. Clearing and Mounting Xenopus Oocytes and Eggs for Confocal Microscopy

The accumulation of yolk during oogenesis renders oocytes and eggs opaque and prevents visualization of structures no more than a few micrometers below the cell surface of uncleared oocytes and eggs. Fluorescence imaging of "intact" Xenopus oocytes and eggs was greatly facilitated by the discovery that a 1:2 mixture of benzyl alcohol and benzylbenzoate (BA:BB) closely matches the refractive index of yolk, thereby rendering Xenopus oocytes and eggs nearly transparent (Murray and Kirschner, as cited in ref. 21). In addition, the refractive index of BA:BB (~1.55) closely matches that of common immersion oils (~1.52) and glass cover slips, reducing the image-degrading effects of spherical aberration.

Although clearing with BA:BB dramatically improves imaging deep within Xenopus oocytes and eggs, the use of BA:BB is not without its drawbacks. First, because BA:BB is immiscible in aqueous solutions, samples must be completely dehydrated with methanol or ethanol prior to clearing (see Subheading 3.7. for staining of actin microfilaments). In addition, samples cleared in BA:BB tend to be brittle and can easily be damaged during mounting. BA:BB also dissolves many plastics, including polystyrene, cellulose acetate, and the plastics used for computer keyboards. Polyethylene and polypropylene tubes are resistant and should be used for all steps utilizing BA:BB. Finally, caution should be used when handling BA:BB because benzyl ben-zoate is an eye and skin irritant.

A Slides with spacers cut from coverslips. B Glass slides with steel spacers

A Slides with spacers cut from coverslips. B Glass slides with steel spacers

Fig. 3. Slides for mounting Xenopus oocytes and eggs for confocal microscopy. (A) Pieces of ovary from juvenile frogs and small (stage I) oocytes are mounted on slides with spacers cut from no. 1 or 2 cover slips. Spacers are "glued" to the slides with fingernail polish and allowed to dry thoroughly before mounting samples. (B) Larger stages I to II and hemisected stages III to IV oocytes are mounted in chambers formed by 0.25-mm steel spacers glued to glass slides with fingernail polish. (C) Hemisected stages V to VI oocytes and eggs are mounted in commercially available well slides (0.5- to 0.6-mm wells). (D) Intact oocytes and eggs are mounted in 1.0 to 1.2 mm thick slides machined from aluminum, with cover slips glued to both faces with fingernail polish. In all cases, cover slips are sealed to slides with liberal applications of fingernail polish and allowed to dry thoroughly before viewing. Reprinted from Methods in Cell Biology, Vol. 70, 2002, pp. 299-416, with permission from Elsevier.

Fig. 3. Slides for mounting Xenopus oocytes and eggs for confocal microscopy. (A) Pieces of ovary from juvenile frogs and small (stage I) oocytes are mounted on slides with spacers cut from no. 1 or 2 cover slips. Spacers are "glued" to the slides with fingernail polish and allowed to dry thoroughly before mounting samples. (B) Larger stages I to II and hemisected stages III to IV oocytes are mounted in chambers formed by 0.25-mm steel spacers glued to glass slides with fingernail polish. (C) Hemisected stages V to VI oocytes and eggs are mounted in commercially available well slides (0.5- to 0.6-mm wells). (D) Intact oocytes and eggs are mounted in 1.0 to 1.2 mm thick slides machined from aluminum, with cover slips glued to both faces with fingernail polish. In all cases, cover slips are sealed to slides with liberal applications of fingernail polish and allowed to dry thoroughly before viewing. Reprinted from Methods in Cell Biology, Vol. 70, 2002, pp. 299-416, with permission from Elsevier.

Because of their size and physical properties, mounting oocytes for high-resolution microscopy poses challenges not faced with smaller cells or sections. Xenopus oocytes and eggs are not adherent to cover slips or slides. Therefore, the oocytes or eggs must be securely mounted or "wedged" between the cover slip and slide to prevent movement during image collection on the confocal microscope. In addition, the region of interest must be sufficiently close to the cover slip to allow image collection with high numerical aperture objectives, which typically have working distances less than 200 |im.

Small oocytes (<150 |im in diameter) or pieces of ovary tissue are mounted on standard slides in wells formed by spacers cut from no. 1 or 2 cover slips (Fig. 3A) and affixed to the slide with fingernail polish. Larger stages I to II oocytes and hemisected stages III to IV oocytes are mounted in glass slides to which stainless steel 0.25-mm spacers have been attached with fingernail polish (Fig. 3B). Hemisected stages V to VI oocytes and eggs are mounted in commercially available glass-well slides (0.5- to 0.6-mm wells; Fig. 3C), with the region of interest (cortex or cut surface) mounted up. Finally, whole oocytes and eggs are mounted in double-sided chambers fabricated from aluminum sheet (0.7-1.0 mm thick), with cover slips attached to both sides (Fig. 3D). These slides allow the imaging of both sides of uncut oocytes and eggs.

1. Dehydrate samples 1 to 2 h with three or four changes of 100% methanol (samples can be stored several months in methanol).

2. Remove the methanol and add approx 1.5 mL BA:BB to the oocytes or eggs. Do not mix.

3. Allow oocytes and eggs to sink slowly to the bottom of the tube (takes 5 to 15 min). Oocytes and eggs will clear and sink as they are infiltrated by BA:BB (see Note 10).

4. Carefully aspirate the BA:BB from the samples to be mounted.

5. Carefully resuspend the samples in a small volume of new BA:BB.

6. Transfer the samples to the appropriate slides in a drop of BA:BB.

7. Aspirate most of the BA:BB.

8. Orient the oocytes and eggs in the desired manner in the slides with forceps or scalpels, if necessary.

9. Fill the well with BA:BB.

10. Drop a no. 1 cover slip quickly on top of the well with gentle pressure.

11. Gently press down on the cover slip to seat it properly on top of the slide. Aspirate any excess BA:BB expelled from the well.

12. Seal the cover slip to the slide with a liberal application of fingernail polish.

13. Allow the slides to dry thoroughly overnight at room temperature or several hours in a fume hood prior to viewing on the confocal microscope.

14. Well-sealed slides are stable for several months in the dark at room temperature (stained with Alexa dyes and TO-PRO-3).

3.7. Fixation and Phalloidin Staining of Actin Microfilaments in Xenopus Oocytes and Eggs

The large pool of monomeric actin (G-actin) in Xenopus oocytes and eggs precludes the use of antiactin antibodies for staining F-actin. We have successfully used fluorescently conjugated phalloidin to stain F-actin in previtellogenic Xenopus oocytes and have had limited success staining F-actin in stage VI oocytes and eggs (5,7). In our experience, even brief treatments with organic solvents such as methanol abolish phalloidin staining of actin microfilaments in oocytes and eggs, prohibiting postfixation with methanol and bleaching with peroxide-methanol. Therefore, albino female frogs should be used to examine the actin cytoskeleton in postvitellogenic oocytes and eggs.

Moreover, dehydration of phalloidin-stained samples in methanol or other BA:BB-compatible solvents extracts any bound phalloidin, prohibiting use of BA:BB as a clearing agent. Although previtellogenic oocytes can easily be stained for F-actin, the absence of bleaching and clearing limits the visualization of phalloidin-stained F-actin to only approx 5 to 10 |im of the cell cortex of postvitellogenic oocytes (stages II-VI) or eggs. Hemisection can be used to examine structures deep within the cytoplasm. However, the inability to section optically below the region of knife damage hinders collection of high-quality images.

1. Fix 5 to 15 oocytes (see Note 11), dejellied eggs, or pieces of ovary tissue from juvenile frogs in 1.0 to 1.5% formaldehyde in fix buffer for 4 to 18 h at room temperature in 1.7-mL microcentrifuge tubes (place tubes on their side to increase exposure of samples to fix and avoid clumping).

2. Wash overnight with several changes of TBSN at room temperature with gentle rotation using an orbital shaker or nutator. If needed, the samples can be hemisected with a sharp scalpel (no. 15) as described above (see Note 12).

Fig. 4. Optical sectioning eliminates the out-of-focus fluorescence that hampers conventional fluorescence imaging of Xenopus oocytes. Conventional images (A and B) and confocal optical sections (C and D) of stage I (A and C) and stage VI (B and D) Xenopus oocytes fixed in FGT as described and stained with mouse anti-a-tubulin and rhodamine-conjugated antimouse immunoglobulin G. C and D are single optical sections collected with a Biorad MRC-600. Reprinted from Methods in Cell Biology, Vol. 70, 2002, pp. 279-416, with permission from Elsevier.

Fig. 4. Optical sectioning eliminates the out-of-focus fluorescence that hampers conventional fluorescence imaging of Xenopus oocytes. Conventional images (A and B) and confocal optical sections (C and D) of stage I (A and C) and stage VI (B and D) Xenopus oocytes fixed in FGT as described and stained with mouse anti-a-tubulin and rhodamine-conjugated antimouse immunoglobulin G. C and D are single optical sections collected with a Biorad MRC-600. Reprinted from Methods in Cell Biology, Vol. 70, 2002, pp. 279-416, with permission from Elsevier.

3. Replace last TBSN wash with fluorescently labeled phalloidin (2.4 units/mL in TBSN) and incubate for 6 to 18 h at room temperature (see Note 13).

4. Wash samples with TBSN with several changes for 24 to 48 h.

5. Mount pieces of ovaries (from juvenile frogs) and small stage I oocytes in glycerol-mounting buffer. For larger oocytes, mount in appropriate slides with TBSN, pH 8.0) containing NaN3 or n-propyl gallate (25-50 mg/mL) to retard photobleaching, if necessary (see Note 14).

3.8. Confocal Immunofluorescence Microscopy of the Cytoskeleton in Xenopus Oocytes and Eggs

The "optical sectioning" afforded by confocal microscopy dramatically reduces the out-of-focus light that degrades the quality and sharpness of images collected from oocytes or other large samples (see Fig. 4). In addition, the collection of serial optical sections and three-dimensional (3D) reconstructions has allowed unparallel views of the organization of the cytoskeleton of Xenopus oocytes and eggs in 3D space (6,14). The following discussion of confocal microscopy is based on our experience with BioRad MRC600 and Zeiss LSM510 confocal laser scanning microscopes. However, aspects of this discussion should be useful with other commercially available confocal microscopes (see your instrument's instructional manual).

The apparent brightness of the image is a function of several parameters. First, the intrinsic brightness of the sample is affected by the quality of the primary antibody or stain, the choice of fluorochrome, and the abundance of the antigen or structure. In addition, the apparent brightness is dependent on the laser power used for illumination and the photomultiplier or detector gain. In practice, the last two parameters must be determined empirically to optimize image quality. In general, the lowest laser power possible should be used to reduce photobleaching. Reduced laser power may be offset by increasing the detector gain, which can result in increased image noise, resulting in images that appear grainy. Image noise can be reduced or eliminated by line or frame averaging, which in turn increases exposure of the sample to laser illumination, resulting in increased photobleaching or photodamage. Laser power, detection gain, and frame averaging should be set to optimize brightness while minimizing image noise, bleaching, and photodamage.

The apparent brightness can also be increased by increasing the thickness of the section collected (by increasing the pinhole size). This in effect increases the volume of the sample from which photons are collected and thus image brightness. However, the superposition of addition cytoplasmic structure and increased fluorescent signal from above and below the focal plane may affect image clarity.

Most commercially available confocal microscopes collect images with 8, 12, or 16 bits of information per pixel in each channel, corresponding to 256, 4096, or 65,536 gray levels per channel. We have found that 8-bit images, which include 256 gray levels per channel, are sufficient for most purposes and require only half the disk storage space required by 16-bit images. The increased number of gray levels provided by 12- or 16-bit images is useful when collecting images with extreme contrast (very bright and very faint details).

The black level and gain of the confocal detection should be adjusted to make maximum use of the full dynamic range of the instrument. This is accomplished by adjusting the black level (also known as baseline or amplifier offset, depending on the instrument) and gain such that the darkest pixels have an intensity value of 0, and the brightest pixels have a value of 255 (in an 8-bit image). This process is facilitated by using a false-color lookup table that maps extremes in intensity (0 and 255) to contrasting colors such as green/blue and red (see Note 15). Adjusting the black levels and gain so that the image includes both green/blue and red pixels ensures that the image utilizes the full range of gray levels available (see Note 15).

One of the major benefits or an advantage of confocal microscopes is the ability to project serial optical sections, resulting in images with extended depth of focus. Most software bundled with confocal microscopes includes tools for projection of optical sections and may include tools for simple 3D reconstructions. In our experi ence, to prevent discontinuities or "banding" in projected images or 3D reconstructions, the section interval or spacing must be matched to the section thickness. In practice, we have found that slight oversampling is required to prevent "gaps" or banding. Greater overlap (oversampling) is required to reduce banding in images collected with low-power objectives (10x and 20x) objectives. When collecting multichannel images, pinholes for each channel should be adjusted to provide uniform section thickness.

Finally, the digital nature of confocal images facilitates postprocessing to optimize brightness, contrast, and other image parameters. Many software packages that allow viewing, processing, and printing of confocal images are available (see Note 16). In general, we limit postprocessing to minor adjustments of image brightness and contrast and to resizing. These processed images are saved as copies, and the original unprocessed images are archived. Proprietary image or data formats used by some confocal microscopes may require that the image be exported into one of the standard digital image file formats (TIFF or JPEG) before they can be opened into an image-processing application. For further discussion of postprocessing of confocal images, see refs. 14 and 15.

In conclusion, the advent and widespread availability of confocal microscopes and development of techniques for fluorescence imaging of Xenopus oocytes and eggs allows visualization of cellular organization heretofore impossible in these large cells. Although optimum processing conditions for each structure or antibody will vary and must be determined empirically, these protocols should prove to be useful starting points.

4. Notes

1. The rotary mixers designed for infiltration of embedding plastics and waxes can easily be modified for this purpose by attaching one or more microfuge tube racks with epoxy, silicon sealant, or other adhesives to the rotor plate.

2. Collagenase A is available from several vendors. However, we have found that collagenase from Roche Diagnostics Inc. gives consistent results at 10 mg/mL in MBSH without the variability noted with collagenase A from other sources. Treatment with collagenase A for 60 to 90 min removes most of the follicular tissue; however, the remaining layer of follicle cells must be manually removed with watchmaker's forceps.

3. Fingernail polish must be compatible with BA:BB. In our experience, Sally Hansen's Hard as Nails clear nail polish (with or without nylon) works well.

4. To minimize accidental aspiration of oocytes or eggs, we often use a micropipet tip to reduce the bore of the Pasteur pipet used on the aspirator. The micropipet tip will be held securely by the vacuum.

5. Bleaching seems to be enhanced by illumination from fluorescent lights or sunlight.

6. Commercially available antibodies are reconstituted (if needed), aliquoted, quickly frozen in liquid nitrogen, and stored at -80°C. Secondary antibodies are first preabsorbed with liver acetone powder for approx 2 to 4 h prior to aliquoting and freezing. To preabsorb the secondary antibodies, a. Transfer dry liver acetone powder to a 1.7-mL microfuge tube, filling the tube about one-third full.

b. Fill the tube with TBSN and mix thoroughly by vortexing (use a spatula to break apart clumps of powder).

c. Sediment acetone powder in microcentrifuge (3 min at full speed) and remove/discard supernatant.

d. Wash acetone powder several more times with TBSN until the supernatant is clear and colorless.

e. After the final wash, remove all TBSN, leaving wet liver acetone powder, and add reconstituted secondary antibody to the washed powder.

f. Mix thoroughly with a spatula and then incubate for several hours at 4°C.

g. Centrifuge for 10 min in microfuge to pellet acetone powder.

h. Carefully remove antibody solution.

i. Aliquot, rapid freeze in liquid nitrogen, and store secondary antibodies at -80°C.

7. Attention should be paid to the order of primary antibody addition because the first antibody may sterically hinder binding of the second antibody to its epitope. For example, we generally bind antibodies to microtubule-associated proteins (i.e., XMAP215, XMAP230, etc.) first to prevent masking by subsequent binding of tubulin antibodies.

8. To minimize background, samples must be thoroughly washed free of any unbound primary antibodies before adding secondary antibodies. Wash efficiency is a factor of both the total time of washing and the number of changes.

9. When feasible, we add secondary antibodies as a cocktail at the appropriate dilutions in TBSN + 2% BSA. Secondary antibodies need to be chosen carefully to avoid unintentional crossreaction. For example, primary antibodies against rat and mouse cannot be used together because the secondary antibodies will crossreact with both primary antibodies. Similarly, you should not use secondary antibodies generated in the same or closely related species (i.e., sheep and goat) as either of the primary antibodies.

10. If oocytes or eggs retain a milky appearance after clearing, they may not have been completely dehydrated. Pass them through several changes of methanol to remove BA:BB and the remaining aqueous buffer and reclear with BA:BB.

11. Collagenase digestion disrupts the organization of actin microfilaments in Xenopus oocytes. Therefore, oocytes must be manually isolated or dissected from ovaries of juvenile frogs.

12. In the absence of methanol postfixation, the nucleoplasm is not well fixed with formaldehyde and may be lost from oocytes when they are hemisected.

13. In our experience, fluorescently labeled phalloidin can also be added to the fix without any differences in F-actin staining.

14. Despite fixation and permeabilization in buffers containing detergents, larger oocytes shrink appreciably in glycerol-based mounting solutions and must be mounted in aqueous solutions such as TBSN or TBS (antifade chemicals can be added).

15. We have used several false-color lookup tables, including the "setcol" table for the BioRad MRC600, which shows bright pixels as red and black pixels as green. The "range indicator" lookup table furnished with the Zeiss LSM510 shows bright pixels as red and black pixels as blue.

16. We usually use Adobe Photoshop (http://www.adobe.com) for most of our postprocessing. Other free software packages useful for postprocessing of confocal images include NIH Image for McIntosh computers (http://rsb.info.nih.gov/nih-image/Default.html), Scion Image for personal computers (http://www.scioncorp.com), and the Java script ImageJ (http://rsb.info.nih.gov/ij/).

Acknowledgments

We would like to thank Ed King for all of his help with the confocal microscopes and Claire Walczak for providing the anti-XKCM1 antibodies used to generate the images for Fig. 2. We also would like to thank present and past members of the lab for their contributions to the methods and protocols discussed. Work presented in this chapter was supported by National Science Foundation grants MCB-950651 and MCB-9904504 and equipment grant DBI-9977204 for the Zeiss 510 confocal microscope.

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