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School of Veterinary Medicine, Department of Anatomy, Physiology and Cell Biology, University of California, Davis, California 95616
ABSTRACT
Sperm undergo extreme variations in temperature and osmolality during cryopreservation, resulting in cell damage that includes plasma membrane defects, changes in cell volume, decreased motility, and flagellar defects. However, the fundamental biologic mechanisms underlying these events are poorly understood. We investigated the effects of osmotic stress and cytochalasins b (CB) and d (CD), naturally occurring toxins that disrupt actin organization, on the actin cytoskeleton and motility of Rhesus macaque sperm (Macaca mulatta). Sperm were diluted in media of low, medium, or high osmolality, or medium-osmolality media containing CB or CD, were stained with phalloidin-fluorescein isothiocyanate, and were processed for microscopy. The majority of sperm incubated in medium-osmolality media exhibited postacrosomal stain, whereas the minority displayed banding patterns of F-actin stain in the head. High-osmolality media, as well as CB and CD incubation, resulted in reorganization of F-actin into bands of stain in the majority of sperm heads. Cytochalasin b treatment also resulted in curled and looped tails, a phenomenon of hyposmotic stress, and CB and CD caused significant, dose-dependent decreases in motility determined by computer-assisted sperm assessment. Rho A cell populations were determined using flow cytometry, and immunocytochemistry analysis demonstrated that Rho A localization was altered after osmotic stress. Together, our results support a mechanism in which reorganization of the actin cytoskeleton induced by osmotic stress and potentially mediated by a Rho A signaling pathway contributes to sublethal sperm flagellar and motility defects.
gamete biology, sperm, sperm motility and transport, stress
Cryopreservation causes extreme fluctuations in osmolality and temperature in sperm, resulting in sublethal and lethal damage to the cells. During freezing, cells are exposed to extracellular ice crystallization that results in concentration of solutes in the unfrozen aqueous channels between ice crystals [1–3]. The cell responds by the osmotic movement of water out of the cell, shrinking in volume until the solute concentrations between intracellular and extracellular compartments equilibrate. Conversely, as cells are exposed to a hypotonic extracellular environment during thawing, cell volume is increased by osmotic movement of water. The extent of cell volume excursions depends on several factors, including water permeability of the cell membrane, cell type, cryopreservative presence, and temperature [4, 5]. Cryopreservation-induced damage to cells includes plasma membrane defects, tail morphologic defects, and decreased mitochondrial membrane potential (MMP), sperm motility, viability, and fertility [2, 3]. It has been suggested that plasma membrane defects may be a manifestation of the close association of the actin cytoskeleton with the plasma membrane [1, 3, 6, 7]. Despite decades of work on improving cryopreservation protocols to increase sperm survival and maintain function, the fundamental biologic mechanisms responsible for cell damage are still not fully understood.
The actin cytoskeleton is organized in a dynamic intracellular network important in maintaining normal cell shape and coordinated cell movement [8], as well as regulating cell volume [9–13]. The actin network undergoes reorganization in response to osmotic stress in organisms from yeast to mammals via rapid activation of MAPK cascades [13, 14]. Hyperosmotic stress was reported to induce reorganization of the cortical actin network in yeast and Dictyostelium [15, 16]. Cytoskeleton reorganization also occurs in cultured mammalian cells and in neutrophils under hyperosmotic conditions, and it involves the Rho GTPase family of signaling molecules [12, 17]. Reorganization of the cytoskeleton following cell swelling and shrinking has been identified as a signal to initiate physiologic cell volume recovery [18]. Sperm also undergo cell volume regulation in response to changes in osmolality during maturation and upon release in the female reproductive tract [19]. The actin network has been implicated in this process. In one study, boar sperm that were incubated in low-osmolality capacitation media containing cytochalasin d (CD), a disruptor of the actin cytoskeleton, exhibited reduced initial swelling [20]. However, it is not clear to what extent the metabolically controlled process of cell volume recovery could occur during the extremes in temperature that accompany cryopreservation.
The actin cytoskeleton is also important for cell motility in many cell types during chemotaxis, wound healing, and proliferation, as well as in response to stress, and actin-based motility requires activation of Rho signaling cascades [8, 21–23]. Actin has been identified in the sperm head, neck, and flagellum in many mammalian species [24–26]. A role for a Rho GTPase, and possibly actin, in mammalian sperm motility was previously suggested by Hinsch et al. [27], who showed that inhibition of Rho proteins resulted in decreased bovine sperm motility over an 8-h incubation period. Furthermore, actin was suggested to be involved in guinea pig sperm motility based on a study using various actin modulators, although the results reported were somewhat ambiguous [28]. In contrast, other studies have reported no effect on total or progressive sperm motility after incubation with actin disruptors, although in one study an effect on sperm velocity and hyperactivation was detected [29]. Clearly, the role of actin in sperm motility and other sperm functions warrants further research.
In this study, we have focused on understanding the biologic mechanisms underlying sperm cell damage in response to osmotic stress. The hypothesis that osmotic stress induces actin cytoskeleton reorganization, potentially activating Rho GTPase signaling and affecting sperm motility and morphology, was investigated. We used an osmotic stress model and cytochalasins to determine the extent of the Macaque sperm actin network, whether the actin cytoskeleton is altered by these stressors, and whether changes in the actin network contribute to other aspects of cell injury.
SYBR14/propidium iodide (PI) and JC-1 were purchased from Molecular Probes (Eugene, OR). All other chemicals were obtained from Sigma Chemical Co. (St. Louis, MO) unless otherwise indicated. Media used were low, medium, or high osmolality (100, 300, or 600 mmol/kg, respectively) Dulbecco PBS (DPBS), containing 0.5 mg/ml BSA, pH 
7.4, to represent hyposmotic stress, control and hyperosmotic stress conditions. Final osmolalities for low and high solutions were achieved by making an adjusted set of anisosmolal solutions (25 and 675 mmol/kg, respectively) to avoid dilution when sperm were mixed with media. Media for measuring mean cell volume, MMP, and viability were the same as above, except no BSA was added. Low-osmolality solutions were prepared by diluting DPBS (to 25 or 100 mmol/kg), and high-osmolality solutions were made by diluting 10x strength DPBS (to 600 or 675 mmol/kg). Final osmolalities were determined using a freezing point osmometer (model 5004; Precision Systems Inc., Natick, MA) that was calibrated against standards for accuracy within ±7 mmol/kg. Biggers, Whitten, and Whittingham medium containing 0.5 mg/ml BSA (BWW; pH 7.4, 290 mmol/kg) was used for sperm processing.
Animal Housing, Semen Sample Collection, and Processing
Monkeys (Macaca mulatta) were housed at the California National Primate Research Center and maintained according to Institutional Animal Care and Use Committee protocols at the University of California. Experiments were conducted in accordance with the National Research Council publication "Guide for Care and Use of Laboratory Animals" (copyright 1996, National Academy of Sciences). Semen samples were obtained by electroejaculation from male macaques as described [30] into 50-ml centrifuge tubes. BWW (4 ml) was added to samples, the coagulum was removed, and sperm suspensions were evaluated for initial motility as described below. Sperm were enriched for live, motile cells, and debris was removed by Percoll (80%) density centrifugation at 350 x g for 25 min. Sperm pellets were washed twice by centrifugation, 5 min each, and were resuspended in DPBS without BSA for cell volume, MMP, and viability measurements, or BWW for cytochalasin and osmotic stress incubations. Cell concentration was determined using a hemacytometer to make final sperm suspensions, and motility was evaluated again after processing.
Sperm cell volumes were measured after incubation in low-, medium-, or high-osmolality media using an electronic particle counter (Coulter Counter, model Z2; Beckman Coulter Corp., Miami, FL) with a standard 50-µm aperture tube and calibrated using 5-µm polystyrene latex beads (Beckman Coulter). The Coulter Counter was interfaced with a computer, and data were acquired with Accucomp software (Beckman Coulter). The protocol used for measuring cell volume has been described previously [31]. Briefly, 15-µl aliquots of sperm suspensions (200 million cells/ml) were diluted in 20 ml of 100, 300, or 600 mmol/kg DPBS and incubated for 10 min at room temperature. A histogram displaying particle count versus volume (cell volume distribution) was recorded, and statistical parameters (mean, mode, median, and standard deviation of volume) were collected from a minimum of 6800 cells per sample.
Mitochondrial Membrane Potential and Viability Evaluation by Flow Cytometry
Aliquots of 25 µl from sperm suspensions (200 million cells/ml) in DPBS were added to 775 µl osmolal-adjusted DPBS for final osmolalities of 100, 300, and 600 mmol/kg and were incubated for 10 min. For mitochondrial evaluation, 0.5 µl of a JC-1 solution (final concentration, 0.5 µM) was added and incubated 10 min, and cells were analyzed using a FACScan flow cytometer (BD Biosciences, San Jose, CA) based on the method of Gravance et al. [32]. JC-1 forms either multimers (j-aggregates) in mitochondria with high membrane potential or monomers in mitochondria with low membrane potential [33]. A 488-nm argon laser was used for excitation of JC-1, with green (JC-1 monomers) and orange (JC-1 multimers) fluorescence detected with emission filters of 535 nm and 595 nm, respectively. Color compensation was applied to correct for spill-over between the two fluorescence detection channels. High and low MMP percentages were determined using histogram data from the flow cytograms by determining the proportion of cells with orange fluorescence (representing JC-1 aggregates with high MMP) and green fluorescence (representing JC-1 monomers with low MMP).
For evaluation of viability, cells were incubated as above, except SYBR14 and PI solutions were added to the sperm suspensions (final probe concentrations were 0.02 and 9.6 µM, respectively). Following incubation, sperm samples were analyzed on a FACScan cytometer. SYBR14 permeates all sperm cells, but PI is only permeable to cells with damaged plasma membranes, making it possible to distinguish live from dead or damaged cells. The SYBR14 and PI probes were excited at 488 nm, and the fluorescence emission spectra were 516 and 617 nm, respectively. A minimum of 8700 sperm-specific events were collected for each experimental treatment.
Osmotic Stress and Cytochalasin Incubations
Sperm suspensions (110 million to 150 million sperm/ml) were divided into 50-µl aliquots; mixed with an equal volume of DPBS containing cytochalasin to yield concentrations of 0, 25, 50, or 100 µM cytochalasin b (CB) or CD; and preincubated on a rocker for 10 min at room temperature. Stock solutions of cytochalasins were made in dimethyl sulfoxide (DMSO; final concentration after dilution with samples was 
1% DMSO), and no effects from this solvent were detected when compared to controls without solvent (data not shown). A second set of sperm aliquots was prepared as above, but without the addition of cytochalasin. Sperm samples with cytochalasin were diluted 1:5 with DPBS (300 mmol/kg) to yield final concentrations of 0, 5, 10, or 20 µM CB or CD. Alternatively, the second set of sperm samples was diluted 1:5 with appropriate DPBS to yield final osmolalities of 100, 300, and 600 mmol/kg. All samples were incubated on a rocker for 20 min at room temperature and then divided for use in the following experiments.
Sperm motility was analyzed after the 20-min incubation on at least 200 cells/sample in a minimum of four fields by computer-assisted sperm analysis (CASA) using HTM Ceros (Version 12.2g; Hamilton-Thorne Research, Beverly, MA). Sperm sample (5 µl) was placed on a prewarmed microscope slide and overlaid with a 22-mm2 prewarmed cover slip, and the slide was maintained at 37°C during analysis by a heated slide holder (Hamilton-Thorne Research). Instrument settings for motility analysis were as previously described [34]. Percent total motility (TM), percent progressive motility (PM), straight line velocity (VSL), curvilinear velocity (VCL), average path velocity (VAP), linearity (LIN), amplitude of lateral head displacement (ALH), beat cross frequency (BCF), and straightness (STR) were determined. Progressive motility was defined as follows: a motile sperm must have a minimum STR of 80% and a minimum VAP of 25 µm/s; STR is the ratio of VSL/VAP x 100 (percentage value). VAP is the average velocity of the smoothed cell path. In some experiments, sperm samples were incubated 20 min more (total 40-min incubation) and reanalyzed as above. Some samples were returned to near control (300 mmol/kg) conditions (by centrifugation at 300 x g in an Eppendorf centrifuge for 3–4 min followed by resuspension in BWW), and motility was reanalyzed.
Phalloidin conjugated to fluorescein isothiocyanate (FITC) was used to stain F-actin [35] using a staining protocol modified from the method of Brenner et al. [36]. After the 20-min incubation, 5-µl aliquots of sperm suspension were removed from samples, spread on clean cover slips (size 1 1/2; 20 x 20 mm), and allowed to dry. Cover slips were fixed with 5% paraformaldehyde (PFA) in Tris-buffered saline (TBS), pH 7.4, for 10 min and washed three times for 2 min each with TBS. Fixed cells then were permeabilized with ice-cold acetone for 10 min, air dried, and stored overnight at 4°C. Samples on cover slips were brought to room temperature and rehydrated, blocked with 1% BSA in TBS for 10 min, and washed with TBS. In a dark, humid chamber, cover slips were stained with 3 µM phalloidin-FITC for 1 h and then washed four times for 2 min each. Cover slips were mounted onto glass slides with mounting medium (Vector Laboratories Inc., Burlingame, CA); the edges were sealed with nail enamel and dried. Slides were stored at 4°C, and each experiment was evaluated in 1 day (with no more than 2 days of storage).
Rho A Analysis by Flow Cytometry and Immunocytochemistry
Sperm were prepared and incubated in media of low, medium, or high osmolality as described above, then immediately fixed and permeabilized with 2% PFA and 0.25% Triton X-100 in DPBS, pH 7.4. Following two washes with DPBS containing 1% BSA, sperm were blocked with 3% BSA in DPBS for 20 min, and then washed again. Samples were then incubated with Rho A monoclonal antibody (Cytoskeleton Inc., Denver, CO) in DPBS and 1% BSA overnight at 4°C. Samples were washed again, incubated with secondary antibody conjugated to FITC for 1 h, washed, and evaluated by FACScan analysis using a 488-nm argon laser; a minimum of 10 000 events were collected for each treatment. For immunocytochemistry, sperm were fixed and permeabilized on cover slips as described for F-actin staining, blocked with 5% BSA in DPBS, and washed. Cover slips were incubated overnight at 4°C with Rho A monoclonal antibody, diluted 100-fold in 1.5% BSA in DPBS. Secondary detection was with goat-anti-mouse immunoglobulin G (IgG) conjugated to FITC in DPBS and1.5% BSA, followed by four washes with DPBS; slides were prepared as described above.
Microscopy and Image Processing
Images were collected with a Delta Vision System (Applied Precision, Issaquah, WA), a cooled charge-coupled device (CCD) camera (CH350; Roper Scientific/Princeton Instruments Inc., Trenton, NJ), and an Olympus IX70 microscope using a 100x/1.35 oil immersion lens (UPlanApo), 1.5x auxiliary lens, and softWoRx 3.22 software suite (Applied Precision, Issaquah, WA). Z sections were obtained at 0.2-µm intervals; all images in this paper represent one z section at the midpoint of the sperm. All data acquired from one experiment (1 day) were thresholded to the same minimum/maximum values and therefore can be directly compared (as in Figs. 3 and 5C, panels 1, 3, and 4), whereas figures representing a composite of different experiments (as in Fig. 5C, panel 2, and Figs. 6 and 7) cannot. Images were colorized using Olympus MicroSuite Biological Suite FIVE, Build 1089, by applying a color look-up key that assigned specific RGB values to each intensity range (representing 20 intensity units). The resulting colors varied from black (lowest intensity, 0–20 units) to blue, green, yellow, orange, red, and white (highest intensity, 241–255; see color look-up key in Fig. 3F). It should be noted that fluorescence intensity was not quantitated; instead, we used the colorized images to better distinguish the pattern of F-actin stain within sperm heads. Images were processed in Adobe Photoshop CS2. Images were cropped, and some were rotated for visual consistency; no other changes to the original images were made except for adjustment of brightness applied equally to all four images in Figure 5C. Slides were analyzed and sperm were counted by one person based on the observed F-actin head staining patterns and tail morphology. At least 75 cells were counted per slide.
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Treatments were compared using one-way ANOVA models followed by Tukey posttest comparisons when data were determined to be distributed normally. Percentage data that were not normally distributed were analyzed using Kruskal-Wallis tests. An unpaired t test was used for comparison of two groups, followed by a Mann-Whitney test (Fig. 4, control vs. CB). GraphPad Prism InStat 3 software (San Diego, CA) was used for all statistical analyses, with the significance level for P defined as <0.05. All data are presented as arithmetic means ± SEM.
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Images of sperm obtained after incubation in media of low, medium, or high osmolality, or with cytochalasin, are shown in 3D to display F-actin structure and staining patterns throughout the sperm. The images obtained were deconvolved (using the deconvolution module in softWoRx) and reconstructed into 3D volumes. The movies shown are 3D representations of the specimens and are not time-lapse videos. Note that the contrast in videos was optimized to demonstrate the F-actin cytoskeletal architecture along the sides of the sperm head, which stained quite intensely; therefore, the head staining patterns are not as apparent as in the 2D images.
An Osmotic Stress Model for Evaluation of Cell Damage
In order to investigate the mechanisms of sublethal cellular injury caused by osmotic stress, an in vitro model was first developed to mimic the osmotic stress component of cryopreservation, without the parallel temperature stress component. Sperm suspensions were incubated in media of low, medium, or high osmolality, and cell volume, MMP, viability, and motility were assessed. Sperm mean cell volume (MCV ± SEM) in control medium (medium-osmolality DPBS; 300 mmol/kg) was determined to be 23.08 ± 2.41 µm3 under the described conditions (Fig. 1A). Sperm in low-osmolality medium (100 mmol/kg) showed a statistically significant increase in MCV (31.04 ± 0.51 µm3; P < 0.05), whereas the MCV for sperm in high-osmolality medium (600 mmol/kg) did not differ significantly (20.17 ± 1.3 µm3) when compared to control values. The increase in sperm cell volume following low-osmolality incubations was expected based on previous work on the osmotic tolerance limits for macaque sperm [34, 37] and indicate that the sperm in our model were undergoing osmotic stress.
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Mitochondrial function was evaluated using JC-1 as a probe to detect high and low MMP (Fig. 1B). A decrease in the proportion of cells exhibiting high potential indicates an increase in damaged and/or dead mitochondria. The JC-1-stained cells were divided into either high-MMP (orange fluorescence) or low-MMP (green fluorescence) populations, according to natural partitioning and appropriate gating parameters for monkey sperm [34]. Sperm incubated in high-osmolality medium (600 mmol/kg) exhibited a significant decrease in the population with high MMP (59.3% ± 10.3%) and a concomitant increase in the population with low MMP (40.8%) compared with sperm incubated in control medium (90.6% ± 1.3%, and 9.4%, respectively; P < 0.05). There was not a significant change in the proportion of high (95.7% ± 1.7%) or low MMP (4.3%) detected in sperm incubated in low-osmolality medium compared with controls (Fig. 1B).
Viability of sperm cells was determined with probes that can differentiate membrane-intact, live cells from sperm that have lost their membrane integrity and are considered to be damaged and/or dead. The percentage of membrane-intact cells after 20 min of incubation in control or anisosmolal media is shown in Figure 2A. The sperm populations with intact membranes did not decrease after low- or high-osmolality incubations, indicating that the sperm remained viable during the conditions used in this study. The apparent increase in viable cells after incubation in high-osmolality media may reflect a population of cells that exhibited sublethal plasma membrane damage or may be due to variability among monkeys.
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Sperm motility was also determined after osmotic stress incubations. Sperm incubation in media of low and high osmolality resulted in an almost complete loss of total motility and progressive motility (TM/PM, 3/1% and 1/0%, respectively), in contrast to the high percentage of motile sperm observed in medium-osmolality controls (Fig. 2B). Decreased sperm motility is a well-established hallmark of anisosmolal conditions. Results from this set of experiments indicate that sperm cells exposed to extremes in osmolality demonstrate increased cell volume in low-osmolality media, sublethal decreases in MMP in high-osmolality media, and dramatic decreases in motility, yet are still viable.
The Sperm Actin Cytoskeleton Undergoes Remodeling During Osmotic Stress
To begin to decipher the mechanisms of cellular injury associated with cryopreservation, we used the osmotic stress model to characterize subcellular changes in sperm cytoarchitecture and physiology. Sperm were incubated in media of low, medium, and high osmolality, followed by phalloidin labeling of F-actin and microscopy to determine the effects on the actin cytoskeleton. As shown in Figure 3, it is possible to detect differences in the overall staining patterns in sperm heads and in midpieces and principal pieces in the flagellum in the black and white images (Fig. 3, A, C, and E). However, to discern more detailed staining patterns and specific distribution of stain, images collected by fluorescence detection were colorized (Fig. 3, B, D, and F) based on their staining intensity by applying a color look-up key (inset in Fig. 3F). The same images displayed in Figure 3 also were deconvolved and reconstructed into 3D volumes, and these are shown in the Supplemental Data (available online at www.biolreprod.org).
Two main patterns of F-actin stain were observed in sperm heads: 1) postacrosomal (PA), which often extended and gradually decreased toward the apical portion of the head (this category varied in intensity level yet maintained the pattern); and 2) bands of stain, with more pronounced stain in the PA region, less or unstained at the equatorial region, and one or more "bands" or regions of stain in the apical sperm head. The PA staining pattern was observed in the majority (72% ± 12%) of sperm in control (medium osmolality) media (Fig. 3, A and B), whereas the banding pattern was observed in 28% of sperm in control samples and varied from subtle to more obvious bands of F-actin stain. In addition, virtually all sperm in control media displayed very bright stain in the neck and midpiece, and stain also was observed in the principal piece of the flagellum that gradually decreased toward the end of the tail. In the 3D views, the sperm from Figure 3, A and B (Fig3A3D1.mov in Supplemental Data) exhibited areas of brightly stained cytoskeletal structure along the side of the head and an intricate staining pattern in the midpiece and principal piece of the flagellum that could not be fully appreciated in the 2D representation.
After hyposmotic stress, almost all sperm (90% ± 7%) exhibited the PA pattern of F-actin stain (Fig. 3, C and D). The cytoskeletal structure observed along the side of the sperm head was hard to detect compared to control sperm in the 3D view (Fig3C3D2.mov). Low-osmolality incubation also caused curling and/or bending of the flagellum in most sperm (see also Figs. 7 and 8, discussed below), similar to that observed in other mammalian species [38], and appeared to cause swelling of sperm heads. The differences in staining pattern distribution in sperm incubated in low versus medium osmolality media were not statistically significant due to a relatively large degree of variation between monkeys. In contrast, the majority of sperm subjected to hyperosmotic stress exhibited the banding pattern of F-actin stain (69% ± 12%), that ranged from subtle to dramatic bands of fluorescence in the apical sperm head (Fig. 3, E and F; Fig3E3D3.mov in Supplemental Data). A portion of sperm displaying the banding pattern had intense F-actin stain at the apical tip of the sperm head (top arrow in Fig. 3F). Sperm subjected to hyperosmotic stress also exhibited intensely stained cytoskeletal structure along the sides of the head, and an intricate staining pattern in the midpiece and proximal piece of the tail in 3D views (Fig3E3D3.mov). The percentages of sperm displaying the PA and Banding F-actin head staining patterns were summarized in Figure 4. The differences in head staining pattern distribution were statistically significant in sperm subjected to hyperosmotic stress versus sperm in control media (Fig. 4; P < 0.05), suggesting that remodeling of the actin cytoskeleton occurred in response to hyperosmotic stress.
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Reorganization of the actin cytoskeleton in other cell systems requires signaling through the Rho family of GTPases [8, 22]; therefore, the hypothesis that the Rho A pathway is involved in an osmotic stress response in sperm was investigated using flow cytometry and immunocytochemistry (Fig. 5). Two distinct sperm cell populations were detected in medium-osmolality samples and after subjection to hyperosmotic stress, as shown in a flow diagram (Fig. 5A). The flow cytometry results are summarized in Figure 5B. Overall, the Rho A-positive cell population detected by flow cytometry represented a relatively small percentage of sperm. This population increased, albeit not significantly, in low- and high-osmolality media compared with sperm in control media. Rho A protein was localized by immunocytochemistry in the PA region of the sperm head (Fig. 5C), where it was evenly distributed in sperm incubated in control media (Fig. 5C, panels 1 and 2). Sperm that were subjected to low-osmolality media exhibited fluorescence in the PA region that was sequestered on the sides of this area in approximately half of the labeled cells (Fig. 5C, panel 3). Sperm subjected to high-osmolality media exhibited variable Rho A labeling patterns, with some sperm exhibiting fluorescence throughout the PA and some displaying sequestered or very little PA fluorescence (Fig. 5C, panel 4). There also was a low level of fluorescence observed in the midpiece and tail of sperm subjected to osmotic stress that was not apparent in sperm incubated in control media.
Cytochalasins Induce Actin Reorganization, Flagellar Defects, and Decreased Sperm Motility
To further characterize the remodeling of the sperm actin cytoskeleton, sperm were incubated in medium-osmolality media containing cytochalasin, followed by microscopy (Fig. 6). The percentages of sperm displaying the two F-actin head staining patterns were summarized in Figure 4. Cytochalasin B treatment induced the banding pattern of F-actin stain in 69% of sperm heads (Fig. 6, B and D; Fig6B3D4.mov in Supplemental Data), a proportion identical to that induced by high-osmolality media, whereas the remaining 31% displayed the PA pattern of actin stain. These differences were statistically significant when compared to the actin staining patterns in sperm incubated in control media (no cytochalasin; P < 0.05). Actin microfilaments also stained brightly along the sides of the head and in a discrete pattern in the midpiece and proximal piece of the flagellum in the 3D view (Fig. 6D represents a snapshot of the sperm shown in Fig6B3D4.mov), which is similar to the results described for sperm incubated in control and high-osmolality media (Fig. 3). Treatment with CD caused virtually identical F-actin staining pattern distributions (data not shown).
In addition to reorganization of F-actin in the sperm head, CB incubation also induced morphologic defects in sperm tails (Fig. 7). Sperm incubated in control (medium osmolality) media exhibited straight tails, with F-actin stain in the midpiece that decreased toward the tail tip (Fig. 7, A and E). In 3D view, the F-actin stain in the flagellum was distributed in a punctate pattern (Fig. 7E and Fig7A3D5.mov in Supplemental Data). In contrast, incubation in medium-osmolality media containing CB induced curls and loops in sperm tails (Fig. 7, B and C), which appeared essentially the same as that observed after incubation in low-osmolality media (Fig. 7D). Tail curling was observed independently of actin head staining patterns (compare Fig. 7B to 7C). Cytochalasin B incubation induced abnormal tail morphology in 35% of sperm compared with 90% of sperm exhibiting abnormal tails after low-osmolality incubation. The results from these experiments are summarized in Figure 8. Similar sperm tail defects were observed after CD treatment, although in a lower percentage of sperm (Fig. 8B).
Cytochalasins are also disruptors of actin-based motility in many somatic cell types; therefore, the effects of cytochalasins on sperm motility were tested. Sperm were incubated in medium-osmolality media containing CB or CD, and motility parameters were determined by CASA (Fig. 9). After a 20-min incubation (Fig. 9A), 10 µM CB caused a small decrease in motility, whereas 20 µM CB caused significant decreases in both total and progressive motility compared with sperm motility in control (no cytochalasin) media (P < 0.01). Similarly, 40-min incubations (Fig. 9B) in 20 µM CB resulted in statistically significant decreases in both total and progressive motility when compared to sperm motility in control (no cytochalasin) media. Incubation with CD for both time periods caused similar decreases in sperm motility (P < 0.05 for total motility). Sperm samples that were returned to medium-osmolality conditions (300 mmol/kg) after 40-min incubations with CB exhibited a return to control levels of total and progressive motility (Fig. 9C; no significant differences when compared to sperm incubated in control media), indicating that the decrease in motility was reversible and was not simply due to toxic effects.
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In this study, we determined the direct effects of osmotic stress on sperm cell physiology in order to investigate potential mechanisms for cell damage. In the osmotic stress model, macaque sperm exhibited damage similar to that previously reported in studies of osmotic tolerance [34, 37]: increased cell volume in low-osmolality media, decreased mitochondrial function in high-osmolality media [34], and decreased motility in both low- and high-osmolality media. Viability did not decrease after incubation in either of these media, indicating that sperm cell damage was primarily sublethal. This model was then used to investigate the extent and distribution of the actin network and its role in contributing to cell damage following osmotic stress.
We have shown detailed microscopic images of actin microfilament staining patterns in macaque sperm; these images were obtained using phalloidin-FITC as a specific probe for F-actin, a cooled CCD camera, and Delta Vision colorizing software. The CCD camera allows a one-time exposure (usually several seconds) with relatively low level illumination, which works well with thin specimens [35], such as sperm. Furthermore, deconvolution of the acquired images (see Supplemental Material) offers a 3D view of the F-actin cytoskeletal framework throughout an entire sperm. Therefore, the use of the CCD camera and Delta Vision system allowed us to achieve high-resolution images that were superior to those we obtained using confocal microscopy or traditional light-level microscopy (data not shown).
In medium-osmolality (control) media, the majority of sperm exhibited F-actin stain in the PA region of the head, whereas a minority of sperm exhibited one or more bands of stain rostral to the PA. Actin has been previously detected in the PA, equatorial, subacrosomal, and/or acrosomal regions of the head and in the midpiece and flagellum in studies with different species and using different localization techniques [24–26]. The two different head staining patterns of actin filaments detected in this study may reflect subpopulations of macaque sperm. Ejaculated sperm are thought to contain a heterogeneous mix of sperm differing in their fertilization capability [39]; thus, the presence of distinct actin-staining subpopulations may potentially reflect subsets of sperm that are more or less fertilization competent. We suggest that the banding pattern observed in sperm heads may represent the population of sperm that are less fertilization competent (see discussion below).
Osmotic stress was shown to induce specific changes in F-actin staining that we interpret as reorganization of the actin cytoskeleton in the sperm head. Based on microscopy results in both 2D (Fig. 3) and 3D (Video Supplement to Fig. 3) images, hyposmotic stress tended to result in more sperm exhibiting the PA pattern of stain compared with control sperm, although there was not a significant difference. It has been previously hypothesized that the actin cytoskeleton might be damaged during sperm cryopreservation [1, 3] because cold temperatures and increased intracellular calcium can depolymerize actin filaments [40, 41]. Studies using ram sperm have provided indirect evidence for this idea, since upon cooling [42] or freeze/thaw [43], an increase in G-actin was detected in the PA region. Since monomeric (G) and polymeric (F) actin are maintained in a dynamic equilibrium in cells [44], this would correspond with a net decrease in actin filaments, which might explain why no F-actin was detected in these studies. The present study provides microscopic data suggesting that hyposmotic stress alone may cause subtle changes in actin organization in macaque sperm, with very few sperm exhibiting F-actin structure in the apical head region.
Hyperosmotic stress resulted in a reorganization of F-actin in the apical sperm head in the majority of sperm, causing formation of bands and localized areas of stain based on microscopy data. To our knowledge, this is the first evidence of actin cytoskeleton remodeling in sperm in response to hyperosmotic stress. The actin cytoskeleton has been shown to undergo polymerization and depolymerization during capacitation and the acrosome reaction in sperm from several mammalian species [26, 36, 45–47], which would be analogous to reorganization of the actin cytoskeleton in the sperm head. Remodeling of the actin network, therefore, appears to be an important component of sperm activation and development of fertilization competence. We suggest the possibility that hyperosmotic-induced reorganization of actin filaments in macaque sperm contributes to the sublethal cell damage that occurs during the cryopreservation process, and may possibly be the basis for the phenomenon of capacitation-like changes observed in frozen-thawed sperm [1, 3]. Capacitation is a complex phenomenon that has been described as a priming step for subsequent sperm acrosome reaction and sperm-egg plasma membrane fusion [46], but it is also thought that once sperm are capacitated they have a very limited lifetime in which they either fertilize or die [48]. Perhaps remodeling of the actin cytoskeleton during cryopreservation prematurely activates sperm or a subset of sperm for fertilization (those exhibiting the banding pattern of F-actin), rendering them incapable of "re-activating" when necessary (i.e., for artificial reproduction techniques).
Cytochalasin B treatment of macaque sperm in medium-osmolality media resulted in virtually identical reorganization of the actin cytoskeleton in the sperm head, as did hyperosmotic stress. The cytochalasins have been described as actin filament disruptors, since they can affect many aspects of actin polymerization and depolymerization, although they do not depolymerize actin per se [49]. Their actions depend on the congener and concentration used; for example, 0.5 µM CB can cause depolymerization of F-actin, whereas 10 µM, similarly to the concentration used in this study, causes disruption of the cortical F-actin network [11]. Therefore, the results obtained from cytochalasin experiments support our finding that hyperosmotic stress induces remodeling of the F-actin network in sperm. Actin reorganization induced by hyperosmotic stress has been described in many organisms and cell types. Hyperosmotic stress induced reorganization of the cortical actin network and redistribution of actin-binding proteins to the cortex in yeast and Dictyostelium [15, 16], and it also induced actin reorganization in cultured cells [12]. In addition, osmotic stress-induced actin reorganization has been linked to regulatory volume decrease (RVD) and increase in a variety of cell types [9–11, 13, 14, 18, 50]. Cell volume regulation also has been reported in boar [19, 20], mouse, human, and monkey sperm [19]. Petrunkina and colleagues reported an association between F-actin and cell volume recovery during capacitation using CD, although no direct effect on the actin cytoskeleton was determined [20]. Our results support a linkage between changes in cell volume induced by osmotic stress and reorganization of the actin cytoskeleton in macaque sperm.
Osmotic stress-induced actin reorganization is known to involve the Rho family of GTPases, which includes Rho, Rac, and Cdc42, in many cell types [11, 12, 17]. Rho family G proteins are small signaling molecules that are regulated upstream by proteins, including the guanine nucleotide exchange factors, and they exert their effects downstream based on their interactions with multiple effector proteins [8, 22, 51]. In the present work, a trend toward an increase in Rho A-positive sperm cell populations was detected after hyposmotic and hyperosmotic stress, suggesting that Rho A signaling may be affected by extremes in osmolality. Additionally, we have shown by immunocytochemistry that Rho A is localized in the PA area of sperm heads in medium-osmolality media, and its distribution was altered after subjection to osmotic stress. The increase in Rho A-positive populations may reflect a redistribution of Rho A protein in the sperm head and an overall increase of Rho A labeling throughout sperm upon changes in osmolality, although determining the precise mechanism(s) involved would require further investigation. It has been reported that actin polymerization during capacitation of ram sperm can be blocked by incubation with C3 exoenzyme, an ADP-ribosyltransferase that inhibits Rho activity [52], suggesting a role for Rho in capacitation [36]. Similar results have been reported for guinea pig sperm, implicating Rho A and B in actin polymerization during the acrosome reaction using activators and inhibitors [53]. Based on the results reported in this study, we suggest that osmotic stress-induced reorganization of the sperm actin cytoskeleton may involve a Rho A signaling-based mechanism.
In addition to the result that osmotic stress has a severe impact on sperm motility, we also demonstrated that CB and CD incubation in medium-osmolality media inhibited sperm total and progressive motility. The inhibition of motility occurred to a lesser degree than in osmotic stress conditions, and the effect was reversible for CB after return to near-control osmolality (Fig. 9). In somatic cells, the actin cytoskeleton is necessary for cell motility and requires cycles of actin polymerization and depolymerization [54]. Cytochalasins are reported to inhibit motility by disrupting the actin network, in addition to having other effects on motile processes [55]. At relatively low concentrations (0.1–10 µM), CB has been reported to inhibit the uptake of monosaccharides in mammalian cells, whereas at higher concentrations (1–100 µM), CB affects morphology and inhibits motility [55, 56]. It has been reported that disruption of actin filaments by CD can lead to activation of p53, a key player in the control of the cell cycle that has a central role in a cell's response to stress signals [57]. However, the finding that both cytochalasins inhibited motility equally, together with microscopy results demonstrating that CB and CD disrupted the actin cytoskeleton, supports a role for actin in mammalian sperm motility. In a recent report, it was suggested that actin plays a role in guinea pig sperm motility, although contradicting results were reported for the various actin modulators used [28]. Inhibition of actin polymerization by latrunculins was reported to disrupt the initiation of motility in maturing wallaby sperm [58]. In a study using human sperm, CB and CD incubation for 2 h decreased sperm velocity and hyperactivation parameters, yet had no effect on total motility [29]. Our results demonstrate that the maximum inhibition of motility occurred within 20 min and CB effects were reversible; therefore, it is possible that sperm in the human study had time to recover to control motility levels in the 2-h incubation [29]. In another study, two ion channel inhibitors were shown to block RVD in response to hyposmotic stress and to decrease progressive motility in ejaculated monkey sperm [59]. Since the actin cytoskeleton is thought to be involved in RVD [18], these results suggested a connection between cell swelling, progressive motility and, indirectly, the actin cytoskeleton. The motility of sperm is based on sliding and bending of the axoneme, a flagellar structure composed of microtubules in a "9 + 2" arrangement plus many accessory proteins [60]. Activation of the dynein ATPases in the axoneme inner arms cause the sliding of outer doublet microtubules. In Chlamydomonas, a unicellular green alga often used as a model for studying eukaryotic flagellar motility [61, 62], and in sea urchin sperm [63], dynein inner arms contain actin [64, 65]. Therefore, it is conceivable that the F-actin localized in mammalian sperm flagella ([24–26] and the present study) may be located in or around dynein inner arms and be susceptible to disruption.
Since actin-based cell motility in somatic cells is initiated by the activation of Rho GTPase signaling molecules [8, 22, 23], a role for Rho in sperm motility also has been suggested. In one study, ribosylation of putative Rho proteins resulted in inhibition of bovine sperm motility [27]. Rhophilin, a Rho-binding protein, and ropporin, a sperm-specific binding protein of rhophilin, have been localized on the principal piece and fibrous sheath of mouse sperm [66–68]. These proteins may interact with AKAP3 in a PKA-dependent mechanism [60, 68, 69], and they have been suggested to regulate sperm motility [68]. Rho proteins have previously been identified in the head and flagellum of sea urchin sperm [63], and in the present study, Rho A was localized to the PA region of the macaque sperm head. Together, our results that cytochalasins induced reorganization of the macaque sperm actin cytoskeleton, a reversible decrease in motility, and that Rho A distribution is altered in response to osmotic stress, suggest that actin reorganization and sperm motility may be regulated by a Rho A signaling mechanism. Alternatively, or additionally, actin disruption may indirectly lead to decreased sperm motility by disorganizing the axoneme or supporting structures of the flagellum. Future investigations will be necessary to develop a clear understanding of the role of F-actin and Rho A in mammalian sperm motility.
We have shown that cytochalasin incubation in medium-osmolality media resulted in morphologic tail defects in 35% of sperm compared with sperm incubated in the same media without cytochalasin. In comparison, low-osmolality incubation resulted in tail curling and bending defects in 90% of sperm in this study. These results support a mechanism in which disruption of the F-actin network contributes partially to the tail coiling defects observed as a result of hyposmotic stress [38]. Furthermore, the cytochalasin-induced reorganization of F-actin in the head—similar to the hyperosmotic effect—and the coiled tails—similar to the hyposmotic effect—occurred independently, suggesting that two different mechanisms may be involved. Overall, our results support a direct connection between cell volume regulation, flagellar morphology, motility, and the actin cytoskeleton in the sublethal damage that occurs during osmotic stress and, potentially, during cryopreservation.
ACKNOWLEDGMENTS
The authors wish to thank Megan McCarthy for technical assistance, and Frank Ventimiglia for help with the Delta Vision System and advice with image processing.
FOOTNOTES
1Supported by a National Institutes of Health grant to S.A.M. (National Center for Research Resources grant # 5R01RR016581) and a NIH supplemental award to L.M.C. 
Correspondence: 2Liane M. Correa, Veterinary Medicine: Anatomy, Physiology, and Cell Biology, University of California, One Shields Ave., Davis, CA 95616. FAX: 530 752 7690; e-mail: lmcorrea{at}ucdavis.edu
Received: 1 February 2007.
First decision: 8 March 2007.
Accepted: 8 August 2007.
REFERENCES
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