Fertilization sets in motion a series of structural and physiological changes in the egg that initiates the developmental program in animal cells. In echinoderm eggs, sperm binding induces a bifurcated signaling cascade that both blocks further sperm binding and reactivates protein synthesis and the cell cycle (Runft et al.,2002). One of the central initiating events after sperm binding in all animal oocytes is the propagation of a calcium wave across the cell that induces cortical granule exocytosis and contributes to the activation of protein kinase C (PKC; Stricker,1999). And while the exact molecular mechanisms by which the calcium wave is initiated remains an area of active investigation (Kuo et al.,2000; Jaffe et al.,2001; Voronina and Wessel,2004), it is clear that cytoskeletal remodeling is one downstream consequence of the sperm signal and calcium wave.
The egg cortex in unfertilized echinoderm and ascidian eggs is composed of a thin layer of cortical actin filaments, short microvilli, and unpolymerized actin monomers (Schroeder,1979; Henson and Begg,1988; Spudich et al.,1988; Bonder et al.,1989; Sardet et al.,2002). After sperm binding, the egg cortex undergoes a remodeling of the actomyosin cytoskeleton that includes a rapid elongation of microvilli and thickening of the cortical shell (Vacquier,1981; Bonder and Fishkind,1995). This remodeling is initiated at the site of sperm binding where a cone of actin filaments rapidly elongates to engulf the sperm head (Schatten and Schatten,1980; Tilney and Jaffe,1980; Cline and Schatten,1986). A wave of cortical actin polymerization then spreads over the surface of the egg by 2 min postfertilization (Cline and Schatten,1986; Yonemura and Mabuchi,1987). Although actin polymerization follows the calcium wave, the increase in actin polymerization has been tied closely to the rise in intracellular pH that follows PKC activation (Begg and Rebhun,1979; Carron and Longo,1982). However, the regulatory molecules that respond to cytoplasmic alkalinization or the potential roles of rho family GTPases such as Rac1 or Cdc42 in modulating actin polymerization have not been identified.
Myosin II (MII) represents one of the most intensely studied contractile proteins, and its regulation during cell division and morphogenesis in nonmuscle cells remains an active area of investigation. However, relatively little is known about myosin's role during fertilization. In ascidian oocytes, a cortical contraction follows the calcium wave across the egg's surface that reorganizes the subcortical cytoplasm and confines the germ plasm into a constricted pole (Sawada and Osanai,1985; Speksnijder et al.,1990; McDougall et al.,1995; Roegiers et al.,1995,1999). In mouse oocytes, both MIIA and IIB isotypes are expressed in oocytes and can be found localized in the cortex adjacent to the meiotic spindle, polar body, and fertilization cone after fertilization (Simerly et al.,1998). Furthermore, expression of nonphosphorylatable mutants of myosin regulatory light chain resulted in defects in meiotic progression, fertilization cone retraction, and cytokinesis (Simerly et al.,1998). Together, these studies suggest that myosin II activation may represent a very early cytoskeletal-remodeling event during development.
Studies of myosin II in echinoderm eggs have focused primarily on the role of myosin II during cell division. Studies in starfish eggs were the first to define the essential role of nonmuscle MII in force generation during cytokinesis (Mabuchi and Okuno,1977; Kiehart et al.,1982). Myosin II contractile activity can be detected in both egg extracts (Kane,1980,1983) and isolated cortices (Walker et al.,1994), and biochemical studies of MII phosphorylation have implicated both heavy chain-based and regulatory light chain-based regulation during cell division (Mabuchi and Takano-Ohmuro,1977; Chou and Rebhun,1986; Yabkowitz and Burgess,1987; Larochelle and Epel,1993; Mishima and Mabuchi,1996; Totsukawa et al.,1996; Komatsu et al.,1997; Walker et al.,1997; Shuster and Burgess,1999). However, a role for myosin II during fertilization in echinoderms is less well established. Biophysical studies of eggs during fertilization noted a dramatic rise in cortical stiffness during fertilization (Mitchison and Swann,1955; Hiramoto,1970), but these studies could not discriminate whether these changes were due to an increase in cortical contractility or the increase in cortical actin. Similarly, early video analyses of fertilization noted a deformation in the cell surface originating at the point of sperm entry (Hamaguchi and Hiramoto,1980; Eisen et al.,1984), but it could not be determined whether the surface deflections were an actual contractile event or a secondary effect of the fertilization envelope's separation from the plasma membrane.
Several lines of evidence have implicated a role for myosin II contractility as one of the earliest cytoskeletal remodeling events of development, and we took advantage of the optical clarity, physiological and physical malleability of sea urchin eggs to further characterize myosin II contractility during fertilization. Time-lapse analysis of fertilization using differential interference contrast (DIC) optics revealed the presence of a sperm-induced contractile wave that was dependent on myosin II activity and the calcium wave. This contractile response could be replicated by mobilizing calcium in unfertilized eggs, suggesting that myosin II–based contractility was directly responsive to calcium. Lastly, inhibition of myosin II activity during and immediately after fertilization slowed retraction of the fertilization cone. Together, these results suggest that myosin II activation is the initial cytoskeletal remodeling event during development and whose activity is required for further cytoskeletal remodeling during the first cell cycle.
Myosin II Organization in Sea Urchin Eggs
It has been demonstrated previously that myosin II, as well as a light chain kinase activity, are present and active in isolated cortical cytoskeletons from unfertilized eggs (Walker et al.,1994,1997), and the fraction of cortical myosin II increases significantly by the time of first cleavage (Walker et al.,1994). To examine myosin localization during fertilization, sea urchin eggs were fixed at various times before or after insemination and processed for actin and myosin II localization. Unfertilized eggs contain a thin shell of F-actin, mostly in the form of short microvilli (Fig. 1A). After sperm binding, a well-characterized wave of actin filament polymerization swept across the egg, resulting in a well-developed cortical shell by 3 min postfertilization (Fig. 1B). Myosin II distribution largely mirrored F-actin (Fig. 1C,D) but was not enriched in the growing fertilization cone (not shown). Additionally, there was a population of MII-labeled linear elements beneath the 1-μm cortical shell and aligned perpendicularly to the membrane (Fig. 1C) that was devoid of F-actin and diminished after fertilization (Fig. 1D). Although the exact identity of these structures was unclear, myosin II localization confirmed earlier biochemical studies indicating that myosin II was present in the cortex before fertilization (Walker et al.,1994).
A Wave of Myosin II Contractility Follows the Calcium Wave During Fertilization
Classic biophysical studies of the sea urchin egg cortex during fertilization noted a sharp, transient increase in cortical stiffness after sperm binding (Mitchison and Swann,1955; Hiramoto,1970). However, from these studies, it was unclear whether the observed changes in cortical stiffness resulted from an increase in cortical contractility or from the increase in cortical F-actin that accompanies the increase in cytoplasmic pH (Begg and Rebhun,1979; Carron and Longo,1982). To examine sperm-induced surface contractile activity in sea urchins, Lytechinus pictus eggs were fertilized and followed by time-lapse microscopy. Within 8 sec of sperm binding, a slight ripple occurred at the sperm entry point (Fig. 2B, asterisk) that developed into a broad flattening at the site of sperm entry that propagated as a wave across the egg surface (Fig. 2B, arrow). When displayed as a kymograph, the contractile wave could be seen not only as a deflection of the surface but also by the displacement of yolk platelets and pigment granules as the wave progressed across the cell (Fig. 2D, bracket). Whereas the degree of deformation was variable from cell to cell, sperm-induced surface deflections were consistently detected in all cells observed where the site of sperm entry was clearly visible (n = 45).
The deformation of the cell surface traveled across the egg concomitantly with the elevation of the fertilization envelope (Fig. 2), but it was unclear whether the alteration of the cell surface was due to a change in the contractile state of the cortex or was a secondary effect of the envelope separating from the plasma membrane. To discriminate between these possibilities, unfertilized eggs were pretreated with dithiothreitol (DTT) or 1 M urea (not shown) to prevent formation of the fertilization envelope (Fig. 3). Whereas the marked flattening of the cell surface observed in eggs with an intact fertilization envelope (Fig. 3D) was not as evident in DTT-treated eggs, a contractile wave was clearly observed traversing the cell (Fig. 3B′). Sperm binding resulted in a contractile wave that initiated at the point of sperm entry (Fig. 3B′, black bracket) and traveled across the cell surface, ending in a small ingression of the membrane after the wave had traversed the egg diameter (Fig. 3B′, white bracket). Similar results were obtained with urea-treated eggs, but urea treatment was less efficient in stripping the fertilization envelope (not shown). To confirm that the contractile wave was due to myosin II, and not some other membrane activity (such as cortical granule release), cells were treated with blebbistatin, a small molecule inhibitor of nonmuscle myosin II ATPase activity (Straight et al.,2003). Recent studies in nonmammalian species revealed that blebbistatin blocks polar body formation in Spisula oocytes (Pielak et al.,2004), inhibits scallop and Dictyostelium myosin II (Limouze et al.,2004; Shu et al.,2005), and cytokinesis in sea urchin eggs at the same concentrations that block myosin II in cultured cells (not shown). To inhibit myosin II function during fertilization, eggs were pretreated with 100 μM blebbistatin for 30 min and the contractile response was monitored after sperm binding (Fig. 3C,C′). Blebbistatin-treated eggs fertilized normally, and in cells not treated with DTT, greater than 92% of eggs had elevated fertilization envelopes (n = 300). However, as shown in Figure 3C′, a minor deflection could be detected after sperm binding, but no surface contractile activity could be detected traversing the cell (n = 15). Together, these results suggest that the deformations observed during fertilization were due to a transient wave of myosin II contractility and was not a secondary effect of the fertilization envelope separating from the cell membrane.
The early events of fertilization in echinoderm eggs include a rapid membrane depolarization (Jaffe,1976; Chambers and de Armendi,1979; Whitaker and Steinhardt,1983) and a “cortical flash” of free calcium (McDougall et al.,1993; Shen and Buck,1993), both of which precede the initiation and propagation of the calcium wave (Jaffe et al.,2001). Earlier descriptions of surface activity ascribed the cell deformation to membrane depolarization (Eisen et al.,1984). However, the apparent contractile wave traversing the egg closely paralleled the elevation of the fertilization envelope, raising the possibility that the contraction was following the calcium wave. To determine whether there was a causal relationship between the calcium wave and the contractile activity after fertilization, eggs were pretreated with the phospholipase C inhibitor U73122, which has been shown to block the calcium transient in sea urchin eggs (Lee and Shen,1998; Leckie et al.,2003), its inactive analog U73343, or blebbistatin. The spatiotemporal relationship between the calcium wave and membrane surface activity after fertilization was imaged using the calcium indicator calcium green–dextran, and kymographs were generated from the resulting image stacks. As shown in Figure 4A, in eggs pretreated with the inactive analog U73343, a noticeable deflection in the cell surface was detected within 10 sec from the earliest detectable rise in calcium green fluorescence signal. As the calcium transient crossed the egg (from left to right), a second more visible deflection of the cell surface followed the increase in calcium green fluorescence. In contrast, suppression of PLC activity with U73122 inhibited both the calcium wave and the contractile response (Fig. 4B). In blebbistatin-treated eggs, neither the initial surface deflection nor the sustained contraction could be detected (Fig. 4C), although the calcium transient progressed normally. Together, these results suggest that the myosin II–dependent deflection of the cell surface closely followed and was dependent on the calcium wave after sperm binding.
Induction of a Myosin II Contractile Response in Cells Under Compression
The cytoskeleton of a spherical cell is isotropic, and subtle changes in contractile activity are difficult to detect by microscopy alone. To further examine the calcium responsiveness of unfertilized eggs, an assay was developed where changes in cortical contractility could be detected simply by monitoring resistance to compression. Compression has been used to monitor contractile changes in echinoderm eggs during cell division (Yoneda and Schroeder,1984; Hiramoto,1990; Rappaport,1996) as well as in unfertilized eggs treated with the phosphatase inhibitor calyculin A (Asano and Mabuchi,2001). Unfertilized eggs stripped of their fertilization envelopes were settled onto glass-bottomed culture dishes, and the seawater was rapidly replaced with a gas-permeable fluorocarbon oil that has been used routinely to compress dividing eggs (Fig. 5A; Sluder et al.,1999). The weight of the oil typically compressed eggs to a thickness of 15–28 μm, and fertilized eggs flattened in this manner remained for 8 hr (not shown). When flattened eggs were injected with 1 mM adenophostin A, a potent IP3 receptor agonist (Sato et al.,1998; Bird et al.,1999), cells began to round up within 10 sec of injection, as evidenced by a retraction of the cell margins (Fig. 5C′,D′,E′). Alternatively, direct mobilization of intracellular calcium by local application of 50 μM ionomycin to the cell margin induced a strong contractile response in both unfertilized and fertilized eggs (Fig. 6B′,C′, respectively), whereas application of 0.1% dimethyl sulfoxide (DMSO) in seawater (Fig. 6A,A′) had no effect. Lastly, the contractile response occurred in both the presence (Fig. 6) and absence (not shown) of Ca++ in the seawater, suggesting that mobilization of internal Ca++ stores was sufficient to induce a contractile response.
Injection of IP3 receptor agonists induced a strong contractile response (Fig. 5) and, together with the local application of calcium ionophore (Fig. 6), suggested that the cortical cytoskeleton could directly respond to a calcium stimulus in the absence of other sperm-induced signals. In an effort to better temporally resolve the initiation of a calcium transient and the subsequent contractile response, unfertilized eggs were injected before flattening with caged-inositol 1,4,5-triphosphate or caged cADP-ribose along with calcium green–dextran. Cells were then flattened and then exposed to a single pulse of ultraviolet (UV) light to uncage the injected reagents (Fig. 7). Whereas the UV flash was able to uncage a control substance (caged fluorescein dextran, Fig. 7A), there was no contractile response from UV exposure alone (Fig. 7A). However, uncaging either caged IP3 or cADP resulted in both an increase in intracellular calcium and a contractile response (Fig. 7B,C). Simultaneous mapping of the calcium transient and the contractile response indicated that contractile response followed the initial calcium transient on average by 4 sec for IP3 and 7 sec for cADP-ribose. From these results, it appeared that the cell cortex could respond within 5 sec of stimulation, which agreed with the earliest contractile responses observed in spherical, fertilized eggs (Fig. 4).
Mobilization of intracellular calcium either using adenophostin, ionomysin, or caged compounds resulted in a rapid contractile response (Figs. 5–7), suggesting that calcium alone may be responsible for the contractile wave after sperm binding. Nonmuscle myosin II is regulated in part by light chain phosphorylation (Bresnick,1999), and to determine whether egg activation resulted in an increase in light chain phosphorylation, unfertilized eggs were treated with ionomycin and analyzed for serine 19 phosphorylation by immunoblotting (Fig. 8). Consistent with earlier in vivo labeling studies (Walker et al.,1997), a basal level of regulatory light chain phosphorylation could be detected in whole cell lysates from untreated eggs (Fig. 8, lane 2). Ionomycin treatment resulted in detectable increases in serine 19 phosphorylation by 1 min after activation, with a 2.3-fold increase after 5 min (Fig. 8, lane 6). Possibly due to the transient nature of the contractile wave (approximately 15 sec) or the differential regulation of soluble and cytoskeletal myosin II (Shuster and Burgess,1999), we were unable to detect a similar increase in myosin light chain phosphorylation after fertilization in whole cell lysates (not shown).
Inhibition of Myosin II Affects Cytoskeletal Remodeling After Fertilization
Time-lapse microscopy of both spherical and manipulated cells suggested that there was a calcium-induced, myosin II–dependent contractile response during egg activation (Figs. 1–7). To determine whether the increase in myosin II activity during fertilization plays an immediate or downstream role in cytoskeletal remodeling during the first cell cycle, eggs were preincubated in blebbistatin or DMSO carrier control, fertilized, and monitored over time for the major morphological characteristics of the first cell cycle (Fig. 9). Blebbistatin treatment had no effect on sperm binding, cortical granule release (as evidenced by elevation of the fertilization envelope), or engulfment of the sperm head or tail (not shown). However, blebbistatin-treated eggs retained their fertilization cones long after the structure had been absorbed in controls. Examination of F-actin organization after fertilization revealed that, by 20 min postfertilization, microvilli were prominent in both control and blebbistatin-treated eggs (Fig. 9C,D). However, by 10 min postfertilization control eggs had absorbed the fertilization cone, and by 20 min, only 3% of cells had a detectable fertilization cone (n = 330 cells). In contrast, 83% of blebbistatin-treated cells still had a detectable fertilization cone (Fig. 9B,D).
Egg activation has been an intense focus of study for over a century, and whereas differences have been noted among the different metazoan groups (Stricker,1999), all eggs and oocytes undergo a similar set of responses after sperm binding. A mobilization of intracellular calcium, establishment of a block to polyspermy, resumption of the cell cycle, and reorganization of the cortical cytoskeleton all occur to differing degrees within the first minute after insemination. Echinoderm eggs have proven an invaluable experimental system for studying fertilization and early development, and we have taken advantage of the optical clarity as well as the physical and physiological malleability of sea urchin eggs to study the activation of myosin II during fertilization. Using time-lapse microscopy, we describe a myosin II-dependent contractile wave that supports and extends the classic biophysical studies of cortical stiffness during fertilization (Mitchison and Swann,1955; Hiramoto,1970). Moreover, this contractile response could be replicated by mobilizing calcium in unfertilized eggs, suggesting that calcium was responsible for activating myosin II activity after fertilization. Lastly, inhibition of myosin II activity retarded the retraction of the fertilization cone. Together, these results suggest that the fertilization-induced activation of myosin II represents another downstream effector of the sperm–egg interaction that participates in the remodeling of the actin cytoskeleton.
Cytoskeletal Remodeling During the First Zygotic Cell Cycle
In comparison to somatic cells, the cortical actomyosin cytoskeleton of unfertilized eggs and oocytes lends relatively little structural integrity to the cell. Ultrastructural analyses identified two populations of actin in the cortical cytoskeletons of unfertilized sea urchin eggs: a filamentous fraction and an amorphous layer of unpolymerized actin (Spudich,1992; Bonder and Fishkind,1995). F-actin was found primarily in the form of short microvilli, with a sparse population of actin filaments lying directly beneath the membrane that often terminated at the microvillar base (Henson and Begg,1988). Localization studies also described a 1-μm layer subjacent to the microvilli that was immunoreactive with anti-actin antibodies but not with fluorescent phallotoxins (Spudich et al.,1988; Bonder et al.,1989), and this population of actin appeared to be associated with cortical granules. Myosin II is also localized in the cortical cytoskeleton of unfertilized eggs (Fig. 1), and as shown in Figures 2–7, a contractile response could be induced by mobilizing intracellular calcium within seconds of activation, suggesting that myosin II was able to generate force against this relatively disorganized cortical actin cytoskeleton. It is possible that new actin filament polymerization is being stimulated by calcium mobilization, but all data to date in both eggs and acrosomal processes point toward cytoplasmic alkalinization as the major stimulus for actin polymerization (Tilney et al.,1978; Begg and Rebhun,1979; Carron and Longo,1982). A detailed ultrastructural analysis of myosin II organization in the egg cortex similar to what has been performed in sea urchin coelomocytes, fish keratocytes, and mammalian fibroblasts (Verkhovsky and Borisy,1993; Svitkina et al.,1997; Henson et al.,1999) will better resolve the linear structures found in unfertilized egg (Fig. 1C) as well as clarify how myosin II generates contractile force against this relatively unorganized substrate.
We detected no visible effects on the actin polymerization in blebbistatin-treated cells, which is consistent with earlier findings of cells with compromised myosin II activity (DeLozanne and Spudich,1987; Knecht and Loomis,1987; Honer et al.,1988; Straight et al.,2003). However, we did find a significant delay in the absorption or retraction of the fertilization cone. Derived from microvilli directly adjacent to the bound sperm head, the fertilization cone rapidly elongates to engulf the sperm nucleus (Tilney and Jaffe,1980; Cline and Schatten,1986). In Lytechinus pictus, the fertilization cone grows to approximately 10 μm with retraction beginning by 9 min postfertilization (Terasaki,1996). However, in blebbistatin-treated cells, the fertilization cone remains visible for more than 20 min after insemination (Fig. 9D), and in 28% of cells examined, the fertilization cone was still detectable 30 min after insemination. Similar results were observed in mouse eggs injected with nonphosphorylatable mutants of myosin regulatory light chain, where both polar body extrusion and fertilization cone absorption were inhibited (Simerly et al.,1998). Live cell analyses of actin filament dynamics in sea urchin eggs revealed that actin in the fertilization cone undergoes a retrograde-like transportation from the cell cortex toward the cell interior (Terasaki,1996), which is consistent with the involvement of a + end-directed actin motor. Moreover, studies of retrograde transport in coelomocyte lamellapodia (Henson et al.,1999,2003) as well as actin dynamics in the contractile ring (Guha et al.,2005; Murthy and Wadsworth,2005) have also implicated myosin II in the regulation of actin filament transport and turnover. Taken together, it appears that one role for myosin II in early development may include the turnover of the earliest actin filament structures.
Regulation of Contractility in Eggs and Oocytes
Nonmuscle myosin II is regulated by both heavy and light chain phosphorylation (Bresnick,1999), and microinjection of nonphosphorylatable mutants of regulatory light chain leads to errors in fertilization cone retraction in mouse oocytes and cytokinesis (Simerly,1998). A low level of myosin regulatory light chain phosphorylation has been detected in unfertilized eggs in vivo, and in isolated cortical cytoskeletons, a light chain kinase activity can also be detected (Walker et al.,1997). Moreover, phosphatase inhibitor treatment of unfertilized eggs induces increased light chain phosphorylation and a strong contractile response, lending further support to the notion that there exists a basal light chain kinase activity before fertilization (Tosuji et al.,1992; Asano and Mabuchi,2001). Our results indicate that the contractile wave follows the calcium transient at fertilization (Fig. 4) and that mobilization of calcium alone was sufficient to induce a contractile response (Figs. 5–7). Ionomycin activation of unfertilized eggs resulted in an increase of serine 19 phosphorylation (Fig. 8), but after normalization, there was only a 2.3-fold increase over DMSO controls. This finding may be due to the fact that only a small fraction of myosin II is cytoskeletal in eggs and blastomeres (∼10%) and that soluble and cortical populations are regulated differentially (Shuster and Burgess,1999). Thus, examination whole cell lysates may underestimate regulatory events at the cell cortex. The Ca++/calmodulin-dependent myosin light chain kinase is a likely candidate to mediate myosin II activation in unfertilized eggs (Bresnick,1999), but studies using established myosin light chain kinase or Rho-kinase inhibitors gave equivocal results (not shown), so we cannot eliminate the possibility that Rho-kinase is involved in the activation myosin II after fertilization. Indeed, Rho GTPase has been localized to cortical granules in unfertilized eggs (Cuellar-Mata et al.,2000), and inhibition of Rho with C3 transferase inhibits cortical granule translocation to the cortex and exocytosis during fertilization (Covian-Nares et al.,2004). Further experimentation will further discriminate the respective roles of Ca++- and Rho-dependent functions in cytoskeletal remodeling during fertilization.
Unless noted otherwise, all chemicals were purchased from Sigma Co (St. Louis, MO). (±)-blebbistatin, adenophostin A, Ionomycin, U73122, and U73343 were purchased by Calbiochem (La Jolla, CA). Calcium green–dextran (molecular weight, 10,000), DMNB-caged fluorescein dextran, NPE-caged inositol (1,4,5) trisphosphate (IP3), NPE-caged cyclic ADP-ribose (cADPR), and rhodamine dextran (molecular weight, 10,000) were purchased from Molecular Probes (Eugene, OR).
Embryo Culture, Microinjection, and Manipulation
The sea urchins Lytechinus pictus and Strongylocentrotus purpuratus were obtained from Marinus, Inc. (Long Beach, CA), and maintained in artificial seawater (ASW) at 15°C. Gametes were obtained by intracoelemic injection of 0.5 M KCl. The egg jelly was removed by washing the eggs in calcium-free seawater and passage through 150 μm Nitex membranes, and in some cases, vitelline envelopes were removed from unfertilized eggs by a brief treatment with 1 M urea or by incubation in ASW containing 10 mM DTT for 10 min. For all live cell experiments, a minimum of 10 recordings were taken for each condition.
To visualize changes in cortical contractility, eggs were placed under compression (Yoneda and Schroeder,1984; Asano and Mabuchi,2001). Dejellied and demembranated eggs were placed in protamine sulfate–coated, glass-bottomed 35-mm culture dishes (World Precision Instruments, Inc., Sarasota, FL), and the seawater was carefully removed and replaced with FC-40 fluorocarbon oil as previously described (Sluder et al.,1999). Cells compressed under in this manner were flattened to a thickness of approximately 15–28 μm, and remained viable for up to 8 hr at 15°C.
For injection experiments, cells were injected using a Parker-Hannifin Picospritzer II pressure injection system before fertilization or flattening under FC-40 oil. To visualize calcium transients in living cells, unfertilized eggs were injected with 2 mM calcium green–dextran (molecular weight, 10,000) in injection buffer (10 mM HEPES, 150 mM potassium aspartate, pH 7.0) to a final concentration of 20 μM. To mobilize calcium in cells under compression, 25 μM DMNB-caged fluorescein dextran, 2 mM NPE-caged inositol 1,4,5-triphosphate or 2 mM NPE-caged cADP-ribose (Molecular Probes) were injected into unfertilized cells in injection buffer in the presence or absence of calcium green–dextran to image the resultant calcium transient at an injection volume of approximately 0.5% of cell volume. Cells were then flattened under fluorocarbon oil as described above. Ten seconds after the initiation of image acquisition, cells were exposed to a 1 sec pulse of UV light to uncage the injected compounds.
Calcium Ionophore Treatment
Unfertilized eggs were treated with 1 M urea for ∼2 min to remove the jelly coat and fertilization envelope, washed in calcium-free seawater, and flattened under fluorocarbon oil. The 0.1% DMSO or 50 μM ionomycin diluted in ASW was loaded into pulled capillary pipettes, and a 20- to 30-pl dose was applied directly to the surface of the flattened cell (Smith et al.,2000).
To detect changes in myosin light chain phosphorylation in response to ionophore treatment, eggs were treated with either 0.05% DMSO or 1 μM ionomycin and samples were collected at time points after ionophore addition. Samples were resolved by sodium dodecyl sulfate-polyacrylamide gel electrophoresis, transferred to Immobilon membranes, and probed with rabbit anti–phospho-serine19 myosin light chain antibodies (Cell Signaling Technologies, Beverly, MA). Blots were stripped and reprobed with monoclonal anti-α tubulin antibodies (Sigma) as a loading control. Blots were imaged using a Bio-Rad Chemidoc XRS system (Hercules, CA), and bands were quantified using Quantity One and Image J software.
Imaging Acquisition and Processing
All live cell experiments were performed on an inverted Carl Zeiss Axiovert 200M DIC and epifluorescence microscope equipped with a computer-driven Uniblitz (Vincent Associates, Rochester, NY) and optical fluorescence shutters to control brightfield and epifluorescence light sources, respectively. The temperature was maintained at 15°C using a Brook Industries heating and cooling stage (Lake Villa, IL). DIC and fluorescence images were recorded using a 12-bit Axiocam CCD camera driven by Axiovision 4.2, and figures were prepared using ImageJ, and Adobe Photoshop 6.0.1 software.
Immunofluorescence and Confocal Microscopy
The 0.1% DMSO carrier control or blebbistatin-treated eggs were fertilized, stripped of their fertilization envelopes, and settled on protamine-coated coverslips. Cells were then fixed and made permeable in 40 mM PIPES, 1 M glycerol, 5 mM ethyleneglycoltetraacetic acid, 5 mM MgCl2, 0.5% Triton X-100, and 3.7% formaldehyde pH 6.9 and fixed for 3 hr at 4°C. Cells were then washed, blocked with 3% BSA in PBS, and processed for actin and DNA localization with Alexa fluor 546–phalloidin and Hoescht 33342 (Molecular Probes), respectively. Images were acquired using an Olympus Fluoview confocal microscope at the Central Microscopy Facility at the Marine Biological Laboratory. To analyze F-actin and myosin organization in S. purpuratus, eggs were fertilized, stripped of their fertilization envelopes, and fixed and then made permeable according to Wong et al. (1997). Eggs were then analyzed for F-actin and myosin II with rhodamine phalloidin and anti–myosin II (Yabkowitz and Burgess,1987) antibodies, respectively. Primary antibodies were detected with Alexa fluor–labeled secondary antibodies, and observed by wide-field epifluorescence microscopy.
The authors thank David Burgess and Ted Chambers for thoughtful discussions that inspired this line of investigation.