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MYOSIN II REDISTRIBUTION DURING REAR RETRACTION AND THE ROLE OF FILAMENT ASSEMBLY AND DISASSEMBLY
Cell Biology International 2002, Vol. 26, No. 3, 287–296 doi:10.1006/cbir.2001.0855, available online at http://www.idealibrary.com on
MYOSIN II REDISTRIBUTION DURING REAR RETRACTION AND THE ROLE OF FILAMENT ASSEMBLY AND DISASSEMBLY GUDRUN KOEHL* and JAMES G. MCNALLY Laboratory of Receptor Biology and Gene Expression, National Cancer Institute, Bethesda, MD 20892, U.S.A. Received 17 July 2001; accepted 14 November 2001
The literature to date suggests a role for myosin II in rear retraction, including evidence that myosin undergoes a characteristic ‘C’-to-spot redistribution at the cell posterior which is associated with retraction. Here we investigate the mechanism of both retraction and the ‘C’-to-spot using Dictyostelium cells containing mutant forms of myosin that affect its polymerization. 3Asp-myosin forms few if any filaments. When 3Asp cells are added to a wild-type mound, the mutant cells move directionally, but rear retraction is markedly delayed, demonstrating that myosin II filaments are essential for efficient retraction. In addition, using a GFP-tagged 3Asp-myosin, we observed a posterior spot pattern associated with retraction, but no cortical ‘C’ pattern preceding it. This suggests that filamentous myosin is required to produce the ‘C’, and that its failure to form results in defective rear retraction. In contrast, an alternate mutant myosin that forms filaments constitutively, 3Ala-myosin, forms ‘Cs’ and then spot patterns at the posterior, but in the interim the spots do not disintegrate. This suggests that spot dissolution occurs by filament depolymerization. In summary our data demonstrate a role for myosin II and the ‘C’-to-spot in efficient rear retraction, and define filament assembly as critical for formation of the ‘C’ and filament disassembly as critical for dissolution of the spot. Published by Elsevier Science Ltd.
K: myosin II; retraction; motility; GFP; time-lapse. A: GFP, green fluorescent protein; MHC, myosin heavy chain
INTRODUCTION The simplest conceptual model for amoeboid cell locomotion involves extension of a cell’s leading edge followed by retraction of its rear (Lauffenburger and Horwitz, 1996; Mitchison and Cramer, 1996). Much data suggest that expansion of the leading edge involves actin polymerization. Less is known about the molecular mechanism of rear retraction, although several lines of evidence have implicated a role for myosin II (hereafter referred to as myosin). *Present address: Klinik und Poliklinik fu¨r Chirurgie, Universita¨tsklinikum Regensburg, Forschungsbau H4, Franz-Josef-Strauss Allee 11, 93053 Regensburg, Germany. To whom correspondence should be addressed: James G. McNally, Laboratory of Receptor Biology and Gene Expression, National Cancer Institute, Bethesda, MD 20892, U.S.A. E-mail: [email protected] 1065–6995/02/$-see front matter
In Dictyostelium, myosin knockout cells cannot translocate when placed on an adhesive substrate (Jay et al., 1995), or within a cell mass where adhesive forces are also high (Eliott et al., 1993; Doolittle et al., 1995). However, the same cells can translocate (slowly) on a glass or agar substrate (Wessels et al., 1988). This substrate-dependent disparity in myosin-null cell motility could reflect a specific requirement for myosin in overcoming high adhesive forces (Jay et al., 1995; Doolittle et al., 1995), which is likely to be a principal task accomplished during retraction of the cell rear (Palecek et al., 1999). But if myosin plays a key role in such retraction, how can myosin-null cells still translocate on less adhesive substrates? This question has led to the suggestion that retraction of the cell rear involves two components, one myosindependent and the other myosin-independent (Jay Published by Elsevier Science Ltd.
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et al., 1995). According to this model, the retractive force generated by the myosin-independent component is strong enough to retract cell extensions under less challenging conditions, as occur for example on a glass substrate, and so myosin-null cells can move on glass. In more challenging environments, as occur on highly adhesive substrates, more retractive force is needed and so myosin-null cells become stuck (Jay et al., 1995; Doolittle et al., 1995). Consistent with a role for myosin in rear retraction is the observation that myosin-null cells fail to form ultrathin lamellae (Jay et al., 1995). As determined by interference reflection microscopy, ultrathin lamellae reflect a very close apposition of the cell’s ventral and dorsal membrane surfaces, analogous to a squeezed tube of toothpaste. In polarized cells crawling on a glass substrate, these lamellae form at the cell posterior during rear retraction (Gingell and Vince, 1982; Gingell et al., 1982). The absence of these lamellae in myosin-null cells suggests firstly that these cells cannot execute the contraction required for membrane squeezing. Secondly since these lamellae are ordinarily associated with retraction, their absence in myosin-null cells suggests abnormal retraction in this mutant. Several myosin localization studies are likewise consistent with a role for myosin in rear retraction. In fixed Dictyostelium cells, myosin concentrates along an arc at the posterior cell cortex (Yumura and Fukui, 1985). A comparable posterior, cortical stain has been observed in living Dictyostelium cells tagged with a GFP-myosin (Clow and McNally, 1999). This arc-like, posterior distribution of GFPmyosin, termed a ‘C’, forms prior to rear retraction and then condenses to a much smaller spot, coincident with retraction of the cell rear. The myosin redistribution correlates with a cell shape change as visualized by combined fluorescence and bright-field microscopy. This so-called ‘C’-to-spot redistribution has been hypothesized to reflect a myosin contraction that contributes to rear retraction (Clow and McNally, 1999). To test the role of the ‘C’-to-spot in rear retraction, we examined both features in two different myosin mutants that affect myosin filament assembly. In Dictyostelium, filament assembly is regulated by the phosphorylation state of three threonines in the tail of the myosin heavy chain (Egelhoff et al., 1993). Monomers are favored when these residues are phosphorylated and filaments are favored when they are dephosphorylated. These two different phosphorylation states have been mimicked in two different site-directed mutants. In one mutant (3Asp), the three threonines have
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been converted to aspartic acid to mimic phosphorylation, and in the other mutant (3Ala) the three threonines have been converted to alanine to mimic dephosphorylation. Each of these sitedirected mutant myosins behaves as expected: 3Asp-myosin is predominantly monomeric and 3Ala-myosin is predominantly filamentous (Egelhoff et al., 1993). By examining cells expressing exclusively one or other of these mutant myosins tagged with GFP (Sabry et al., 1997), we have shown that a defective ‘C’-to-spot can result in defective retraction, and that the formation of a ‘C’ depends on filament assembly while the dissolution of a spot requires filament disassembly.
MATERIALS AND METHODS Cell culture Cells were grown in HL5 medium in 100 mm Petri dishes. The following strains were used: HS1, a myosin II—null (Ruppel et al., 1994); JH10 (Hadwiger and Firtel, 1992), a derivative of KAx 3 as a parent strain for HS1; HS1GFPmyo (Clow and McNally, 1999), a derivative of HS1 containing wild-type GFP-tagged myosin II, and pBIG 3Asp and pBIG 3Ala (Egelhoff et al., 1993), both derivatives of HS1. To generate cells with GFP-3Asp or GFP-3Ala fusion proteins, we used constructs generously provided by Drs James Spudich and Ji-Hong Zang, Stanford University (Sabry et al., 1997). These constructs were used to transform HS1 and several different clones were selected for study. Both the mutant and wild-type GFP-myosins have been previously characterized (Moores et al., 1996; Sabry et al., 1997). Cell preparation for microscopy Cells were collected from Petri dishes by centrifugation and washing in phosphate buffer (Clow and McNally, 1999). To initiate development to the mound stage, the cells were plated on dialysis membrane laid on 2% agar. As mounds formed, the membranes were transferred to a cover slip kept in a humid chamber. For most experiments, labeled cells were mixed with a much higher percentage of unlabeled cells to permit clear identification of the labeled cells within the multicellular mass. In such mixtures, we typically used from 1–5% labeled cells. The GFP-3Asp-myosin was used directly as a cell label, since the GFP stain was largely static and always filled the cytoplasmic compartment. This was not the case for GFP-3Ala-myosin which
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stained only the cell posterior. For tracking of this cell line, we labeled cell cytoplasm with 5-chloromethylfluorescein diacetate (CMF, molecular Probes, Eugene, OR, U.S.A.) to permit a more accurate determination of the cell’s center of mass (Clow and McNally, 1999). Microscopy Images were collected with either a Leica TCS NT SP confocal microscope (Heidelberg, Germany) or a scientific grade cooled CCD camera on a conventional wide-field Olympus IX70 microscope running Deltavision software (Applied Precision, Issaquah, WA, U.S.A.). Depending on the experiment, we used either a 10, 20 or 25 objective lens. For the confocal microscope, the pinhole was set to 1.0 Airy disk unit. Images were collected at intervals from 10–60 s, depending on the experiment. In some cases, multiple focal plane images were collected and then a maximum intensity projected image was created. For additional details of the microscopy, including controls to insure cell viability during imaging, and cell tracking see Clow and McNally (1999). RESULTS 3Asp-myosin was engineered to form few if any myosin filaments, and instead to remain almost exclusively monomeric (Egelhoff et al., 1993). The inability to form thick filaments yields a phenotype in 3Asp-myosin that is virtually identical to a myosin-null. In particular, both mutants arrest development at the first multicellular stage, known as the mound (DeLozanne and Spudich, 1987; Knecht and Loomis, 1987; Egelhoff et al., 1993). At this stage, cell motion defects in both 3Asp and myosin-nulls become pronounced. In each mutant, cells within the mound jiggle in place instead of the vigorous, directed cell motion seen in the wild-type (Doolittle et al., 1995; Clow and McNally, 1999). GFP-3Asp-myosin mutant cells exhibited a comparable developmental phenotype to the conventional 3Asp mutant: development arrested at the mound stage, at which time directional cell motion in the mound ceased, and individual cells instead jiggled largely in place (Fig. 1A). When these GFP-3Asp cells were mixed with predominantly wild-type cells, we found that the mutant cells entered the mound and moved directionally (Fig. 1B), unlike myosin-null cells which drift aimlessly to the mound core or periphery when seeded into wild-type mounds (Clow and McNally, 1999).
Fig. 1. 3Asp cell trajectories within a mound. Individual cells tagged with GFP-3Asp-myosin were mixed with unlabeled cells and then the fluorescent cells tracked from time-lapse images of the mound. A cell position at each time-point is marked by a square, and successive time points are connected by thin lines. the interval between time points if 30 s. Different colors represent different cells. The mound border as determined from a bright field image is shown as a solid black line. (A) Both labeled and unlabeled cells carry 3Asp mutant myosin. The mutant cells translocate only small distances and so lines connecting separate time-points are obscured. The trajectories appear instead as small blobs. This is characteristic of a myosin-null phenotype (Doolittle et al., 1995; Clow and McNally, 1999) and radically different from the rotational cell trajectories observed in the parent strain (Kellerman and McNally, 1999). (B) Labeled (and tracked) cells are 3Asp mutants and unlabeled cells are wild-type. The mound is composed of predominantly wild-type cells (95%). In this largely wild-type mound, the 3Asp mutant cells are capable of executing essentially normal rotational motion. This is in marked contrast to myosin-null cells which drift to the outer edge or inner core of the mound (Clow and McNally, 1999). Scale bars 50 m.
Directional motion in the 3Asp mutant makes it feasible to define a rear retraction. In wild-type cells, rear retraction is consistently associated with a stereotyped ‘C’-to-spot redistribution of GFPmyosin at the cell posterior (Fig. 2A) (see also Clow and McNally, 1999). GFP-3Asp-myosin exhibited one characteristic of the ‘C’-to-spot redistribution of wild-type myosin: it formed a posterior spot (arrows in Fig. 2B,C) that was followed by a rear retraction (Fig. 2B, time-points 320–460; Fig. 2C time-points 180, 200). But compared to wild-type GFP-myosin (Fig. 2A), GFP3Asp-myosin was diffusely and dimly distributed throughout the cytoplasm and not enhanced at the cell cortex (Fig. 2B,C) except for the spot patterns noted above. In particular, cortical ‘C’ patterns were never observed at the cell posterior of GFP3Asp-myosin cells. Thus, both wild-type and GFP-3Asp-myosins exhibited spot patterns associated with retraction, but in wild-type cells, this was preceded by a cortical ‘C’ pattern, whereas in the mutant it was preceded by a more diffuse cytoplasmic stain.
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Fig. 2. Representative timelapse sequences of individual wild-type GFP-myosin (a) and GFP-3Asp-myosin cells (B,C) moving within a wild-type mound. The cells undergo rotational motion in the mound (Fig. 1B), but have been oriented here such that motion in each panel is upward. Time points in seconds are shown below each image. As previously described (Clow and McNally, 1999), the wild-type myosin (A) is initially rather uniformly distributed, but then exhibits a ‘C’-to-spot at the posterior (timepoints 288, 320, 352). The 3Asp GFP mutant myosin is also distributed rather uniformly throughout the cytoplasm. However, at certain time points in the movement sequence, a myosin spot can be discerned (arrows in B,C), although a precursor ‘C’ is never visible. The formation of the myosin spot precedes retraction, as judged by the shortening and fattening of the cell body at subsequent time points (320–460 in B and 180–200 in C). Arrows indicate location of myosin spots in wild-type and 3Asp cells. Dotted lines indicate the cell periphery as judged by contrast enhancement of the cytoplasmic fluorescence. Scale bars 5 m.
Based on its correlation with retraction, we previously hypothesized that the ‘C’-to-spot pattern was mechanistically involved in rear retraction (Clow and McNally, 1999). To test if the absence of a ‘C’ in 3Asp cells had functional consequences, we compared the time between the first appearance of a posterior myosin spot and the time when a rear retraction occurred, for 3Asp cells vs wild-type cells. Included in this analysis were only those cells for which both a GFP-myosin spot and a subsequent retraction could be clearly identified. We found that nearly 60% of 3Asp cells had retraction times that were significantly slower (i.e. more than 3 standard deviations longer) than the average for wild-type cells (Fig. 3). These measurements suggest that both ‘C’ patterns and filamentous myosin (largely absent in the 3Asp mutant) are required for efficient rear retraction. Even though cortical ‘C’ patterns did not form at the cell posterior, other aspects of the wild-type ‘C’-to-spot myosin sequence were not altered in the 3Asp mutant, namely the formation of a myosin
spot and the subsequent dissolution of this spot. This suggests that filamentous myosin was required for formation of the ‘C’, but not dissolution of the spot. To further investigate these possibilities, we examined a second myosin mutant, 3Ala, which forms constitutive thick filaments (Egelhoff et al., 1993). The 3Ala mutant proceeds through development normally (Egelhoff et al., 1993). Consistent with this, we found that cell movement in the 3Ala mound was indistinguishable from wildtype (Fig. 4A). That is, cells exhibited rotational trajectories characteristic of the KAx 3 parent strain (Kellerman and McNally, 1999). Normal, directional movement also continued when the mutant cells were mixed into wild-type mounds (Fig. 4B), suggesting that the mutant cells could compete effectively among wild-type neighbors. Cells containing GFP-3Ala-myosin also behaved like cells containing 3Ala-myosin in these assays, but their patterns of myosin distribution were different from wild-type GFP-myosin. Virtually
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Fig. 3. Rear retraction is significantly less efficient in the 3Asp mutant. To assess the role in rear retraction of myosin and its ‘C’-to-spot redistribution, we compared the time between the spot’s appearance and the subsequent retraction of the cell body in wild-type (white bars) vs 3Asp-myosin mutant cells (black bars). 51 wild-type and 34 3Asp rear retractions were identified from cells moving within the mounds. The measured time intervals for each retraction event are shown on the x axis, and the percent of retractions at that time interval are shown on the y axis. Negative numbers for the wild-type indicate that in some cells retraction occurred before the myosin spot could be detected. The mean of the wild-type distribution is shown (gray arrow) as is the distance encompassing three standard deviations from the mean (gray bar +3SD). Nearly 60% of the 3Asp data fall outside this three standard deviation range, demonstrating a significant difference in rear retraction efficiency in the mutant vs wild-type.
no diffuse cytoplasmic fluorescence was observed. Instead a single large and bright myosin spot was typically found at the posterior cortex of locomoting cells (Fig. 5A,B). ‘C’ patterns appeared to form directly from the myosin in the spot, growing bidirectionally to yield a ‘C’ still superimposed on a spot (a ‘spotted’ ‘C’, Fig. 5B time-points 96 and 120), or in some cells eventually a thickened ‘C’ by itself with only a hint of a superimposed spot (Fig. 5A time point 196). As the ‘C’ condensed to a spot once more, the spot typically persisted until a new ‘C’ pattern eventually formed. In contrast, in wild-type cells completing a rear retraction, the myosin spot ordinarily disappeared entirely leaving a diffuse cytoplasmic distribution of myosin, before a subsequent de novo appearance of a cortical ‘C’ pattern (see Fig. 2A and also Clow and McNally, 1999).
Fig. 4. 3Ala cell trajectories within a mound. Individual 3Ala cells tagged with a fluorescent dye (CMF) were mixed with unlabeled cells and then the fluorescent cells tracked from time lapse images of the mound. A cell position at each timepoint is marked by a square, and successive time-points are connected by thin lines. Different colors represent different cells. The mound border as determined from a bright field image is shown as a solid black line. The interval between timepoints is 120 s in (A) and 104 s in (B). Both labeled and unlabeled cells carry 3Ala mutant myosin. The mutant cells exhibit normal rotational motion characteristic of the parent strain (Kellerman and McNally, 1999). (B) Labeled (and tracked) cells are 3Ala mutants and unlabeled cells are wild-type. The mound is composed of predominantly wild-type cells (95%). In this largely wild-type mound, the 3Ala mutant cells compete effectively and still exhibit normal rotational trajectories. Scale bars 50 m.
To quantify these observations of a persistent myosin spot in GFP 3Ala-myosin cells, we categorized and counted myosin patterns observed at the posterior of both 3Ala and wild-type cells (Fig. 6). The fraction of cells (40% in wild-type vs 33% in 3Ala) exhibiting some form of ‘C’ pattern was similar between the two mutants. This suggests that both the stimulus and the ability to form a ‘C’ pattern were comparable in both cell lines. However, the fraction of cells exhibiting only a posterior myosin spot was radically different: 13% in wildtype vs 67% in the 3Ala mutant (Fig. 6). The dramatic increase of mutant cells with myosin spots was at the expense of two wild-type characteristics that were never observed in the mutant, namely diffuse cytoplasmic fluorescence (15%), or uniform cortical staining (32%). Such wild-type patterns (Fig. 6A–D) appear either before or after a ‘C’-tospot condensation, and persist for variable periods until a new ‘C’-to-spot condensation is initiated (Clow and McNally, 1999). Our quantitative measurements demonstrate that these intermediate patterns are absent in the 3Ala mutant and suggest that they are replaced by myosin spots, presumably because the myosin spot fails to disintegrate normally.
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Fig. 5. Representative timelapse sequences (A) and (B) of GFP-3Ala-myosin cells moving within the mound. The cells undergo rotational motion in the mound (Fig. 4), but have been oriented here such that motion in each panel is upward. Time points in seconds are shown below each image. Virtually all of the GFP mutant myosin collects at the cell posterior where it typically forms a large spot. This spot redistributes into ‘C’-like patterns at the posterior (time points 172–196 in (A) and time points 72–120 in (B) which then condense to form large spots once more (time points 220, 244 in (A) and time points 168, 288 in (B)). At no time is 3Ala-myosin found diffusely distributed throughout the cytoplasm, in marked contrast to wild-type or the 3Asp mutant (Fig. 2). Scale bar 5 m.
Despite the failure of the myosin spot to dissolve, we did not observe significant impairment in cell motility, or more specifically rear retraction. The only obvious defect in the 3Ala cells was that they were less elongated, and in particular less curved at their posterior. This was manifested in ‘C’ patterns that were also less curved or in some cases even straight, forming in the extreme an ‘I’ perpendicular to the cell’s direction of motion (Fig. 6F). DISCUSSION Previous studies have shown that mysosin II is required both for cell division in suspension (DeLozanne and Spudich, 1987; Knecht and Loomis, 1987) and for providing cortical stiffness during interphase (Pasternak et al., 1989; Knecht and Shelden, 1995; Shelden and Knecht, 1995; Shelden and Knecht, 1996). Other studies have provided some evidence of myosin involvement in rear retraction (Yumura and Fukui, 1985; Jay et al., 1995; Clow and McNally, 1999). We now extend these studies with a mutant analysis demonstrating that rear retraction was markedly delayed in cells containing largely monomeric myosin. Retraction times in this 3Asp mutant ranged from twice as long to ten times as long as the average for wild-type. These data suggest that
myosin thick filaments are normally required for efficient rear retraction, as first hypothesized by Jay et al. (1995). In the absence of filaments, we find that retraction is still possible, but inefficient. These observations therefore strongly support the view that myosin II plays an important role in rear retraction. Based on the early myosin knockouts in Dictyostelium, it has repeatedly been stated that myosin is essential for cell division, but not cell motion. The data reported here, plus recent studies of cell division, have changed this picture and demonstrate that both processes, cell division and cell motion, have myosin-dependent and myosinindependent components. Dictyostelium myosinnull cells fail to divide in suspension culture, but in the wild, these cells never grow in suspension and so the physiological significance of this phenotype is unclear. In contrast, the same myosin-null cells undergo fairly normal cytokinesis on an adhesive substrate (Zang et al., 1997; Neujahr et al., 1997), exhibiting mitotic rounding, cell elongation, polar ruffling, furrow ingression and separation of daughter cells. This demonstrates that cytokinesis has a myosin-independent component. Dictyostelium myosin-null cells also exhibit a myosinindependent component to motility. They move sufficiently well to form a cell mass known as the mound. However, this mound arrests development and a fruiting body never forms. Cell movement in
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Fig. 6. 3Ala myosin exists predominantly as posterior spots at the expense of diffuse cytoplasmic or cortical distributions. To quantify differences between 3Ala myosin and wild type myosin, we categorized the different myosin distributions observed in live cell movies. Wild-type GFP myosin exhibited four different patterns: A: ‘cytoplasmic’—a rather uniform distribution throughout the cytoplasm; B: ‘cortical’—a dim enhancement of staining throughout much of the cell membrane (typically much dimmer than the enhancement seen at the posterior cortex and defined as a ‘C’); C: ‘C’—a brightly stained arc-like pattern found at the posterior cortex; D: ‘spot’—a bright puncta located at the posterior cortex. GFP3Ala-myosin exhibited a different assortment of staining patterns; E: ‘C’—a posterior cortical arc very similar to that observed for wild-type myosin; F: ‘I’—a bright, posterior cortical stain that was straight instead of arc-like; G: ‘spotted C’—a ‘C’-like posterior pattern with a spot superimposed; H: ‘spot’—a bright puncta located at the posterior cortex, typically somewhat larger in diameter and brighter than those seen in wild type. Using these categories as a guide, we randomly selected single time point images from five different time-lapse movies of wild-type or 3Ala cells and then classified the myosin distribution in each cell at those selected timepoints. For purposes of this quantification, we classified all of the 3Ala patterns shown in E–G as a ‘C’. We examined 317 wild type cells, and 324 3Ala cells. As shown in the histogram, cytoplasmic and cortical myosin are never observed in 3Ala cells, and appear to be replaced by an increase in spots in this mutant. ‘C’-type patterns account for roughly similar percentages in the two strains. Scale bars 5 m.
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the myosin-null mound is radically defective (Elliot et al., 1993; Doolittle et al., 1995; Clow and McNally, 1999), demonstrating in this case that under certain normal physiological conditions, myosin is vital for cell motion. Our present data show that one key role for myosin in locomotion is rear retraction, and argue more generally that myosin is equally important for cell locomotion as it is for cell division. Our observations suggest that myosin’s role in rear retraction is mediated by its ‘C’-to-spot condensation at the cell posterior. We measured a delay in retraction times in the 3Asp mutant that was calculated as the time interval between the first appearance of the posterior myosin spot and the completion of retraction. That this time was on average much longer in the mutant implies that the condensation from a ‘C’ to a spot has a functional consequence in wild-type, namely subsequent retraction of the rear. In principle, the ‘C’ could provide structural support for the cell posterior during retraction. We think this unlikely given that 3Asp cells which lack a ‘C’ showed no unusual deformation localized to a region at the cell posterior where a ‘C’ should have formed. It seems more likely that the condensation of a ‘C’ to a spot reflects myosin motor function that is important for normal rear retraction. It is of interest that the predominantly monomeric myosin in the 3Asp mutant was capable of forming a spot, but that the mutant cells were nonetheless incapable of efficient retraction. This implies that monomeric myosin responds to the signal to form a spot, but yet cannot produce a retraction. One plausible interpretation is that spot formation occurs via movement over actin, a process that both monomers and thick filaments could accomplish. Such cortical flow of myosin II occurs in cell division (Yumura, 2001) and so might also occur in the transition from a ‘C’ to a spot. Our observation here that 3Asp cells participate in rotational motion in a wild type mound is the first phenotypic difference detected between 3Asp and myosin-null cells. Unlike 3Asp cells, myosin-null cells do not exhibit rotational motion in a wild type mound, but rather drift to the mound periphery or interior (Clow and McNally, 1999). This difference between the two myosin mutants most likely arises due to small myosin multimers that are thought to be present in 3Asp cells (Egelhoff et al., 1993). These multimers may supply sufficient force or structural support to enable the 3Asp cells to participate in rotational motion within the mound. For example, small multimers might enable inefficient retraction as
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observed in the 3Asp cells. These multimers may also supply sufficient cortical integrity to 3Asp cells such that they are stiff enough to compete with wild-type cells and therefore participate in rotational motion within the mound. In this latter scenario, inefficient rear retraction could occur via the myosin-independent retractive force described for myosin-null cells crawling on a substrate (Jay et al., 1995). Regardless of the precise mechanism of inefficient retraction, our data show that 3Asp cells have enough residual myosin function to enable movement in a wild-type mound, but not enough myosin function to permit normal rear retraction. Our current data suggest that the myosin ‘C’-tospot may be regulated in part by the phosphorylation state of myosin. We found that normal dissolution of the myosin spot fails to occur in a mutant, 3Ala, that forms thick filaments constitutively. In this mutant, ‘Cs’ form from the spot, instead of de novo from a diffuse cytoplasmic distribution of myosin, as occurs with wild-type myosin. These observations suggest that ordinarily the myosin spot dissolves due to a signal for thick filament disassembly. In the 3Ala mutant, this signal cannot overcome the tendency of this myosin to form filaments, and so the myosin spot fails to disintegrate upon completion of retraction. These observations parallel earlier studies in which 3Ala-myosin was shown to persist at sites where conA was capped (Egelhoff et al., 1993), also suggesting a failure to dissociate in response to a signal for disassembly. A logical candidate for a disassembly signal is myosin phosphorylation at three threonines in the heavy chain tail. 3Ala cells cannot be phosphorylated at these residues because they have been mutated to alanine, thereby mimicking a continuously dephosphorylated state. As a consequence, 3Ala-myosin should be recalcitrant to disassembly by a kinase specific for these residues, and so should fail to disassemble if this kinase normally triggers disintegration of the myosin spot. Our data, however, do not strictly rule out other, as yet unidentified, signals for thick filament disassembly that are simply overridden by the tendency of 3Ala-myosin to remain filamentous. Despite the inability of the 3Ala-myosin spot to disintegrate, this same myosin can form a cortical ‘C’. In contrast, ‘Cs’ were absent in the complementary mutant, 3Asp, in which myosin is largely monomeric. This suggests that formation of a cortical ‘C’ requires thick filaments. These observations and conclusions are paralleled by studies of myosin localization during cell division.
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Myosin normally accumulates in the cleavage furrow, but GFP-3Asp-myosin fails to localize to the furrow and remains diffusely cytoplasmic during cell division, suggesting also in this case that thick filament formation is required for normal myosin localization (Sabry et al., 1997). In both cases (cell division and rear retraction), a logical candidate for an assembly signal is myosin dephosphorylation at the same three threonine residues in the heavy chain tail. 3Asp cells cannot be dephosphorylated at these sites because these residues have been mutated to aspartic acid, thereby mimicking a continuously phosphorylated state. As a consequence, 3Asp-myosin should be recalcitrant to an assembly signal by a phosphatase that may normally help induce thick filament assembly to form a ‘C’ at the posterior cortex during rear retraction or an accumulation at the cleavage furrow during cytokinesis. Our results with 3Ala however suggest that dephosphorylation of these three threonines is not absolutely required to induce formation of a ‘C’ pattern. 3Ala-myosin cannot be dephosphorylated at these three threonines, but ‘Cs’ still formed in this mutant. Thus some additional signal beyond dephosphorylation at these three residues must induce 3Ala-myosin to leave the spot and spread out to form a ‘C’ along the posterior cortex. The ability of 3Ala cells to form a ‘C’-to-spot pattern could account for their relatively normal rear retractions. In this mutant, the only obvious defect in the ‘C’-to-spot pattern was the failure of the spot to disintegrate normally. One potential consequence of this defect that we could discern was the formation of straighter, less extended ‘C’ patterns in comparison to wild-type ‘Cs’ which presumably form from a more dispersed distribution of myosin monomers at the cell posterior. These constrained ‘C’s’ in 3Ala-myosin might make retraction slightly less efficient in comparison to wild-type, but such a difference would be difficult to detect. In summary, our observations define a role for myosin in rear retraction and suggest a model for this process (Fig. 7). The initial assembly of myosin as a ‘C’ at the posterior cortex requires filaments, and may be regulated in part by myosin heavy chain dephosphorylation (Fig. 7A,B). The condensation to a spot could reflect a myosin contraction over a cortical actin meshwork that helps break adhesive bonds between cells (Fig. 7C,D) or on an adhesive substrate. Once freed from these bonds, the cell rear retracts via a myosin-independent retractive force, such as elasticity of the cytoskeleton (Fig. 7E). The final dissolution of the
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Fig. 7. A working model for myosin’s role and regulation in rear retraction. The panels are shown in clockwise order to emphasize the cyclic nature of the process. The schematic shows a group of cells within a multicellular mass. The center cell undergoes a rear retraction. Within this cell, myosin monomers are indicated by red dots, myosin thick filaments by thick red lines, and cell–cell adhesion bonds by blue lollipops. At the outset, a signal for retraction localized to the posterior yields thick filament assembly at the cortex A,B. This signal may involve myosin dephosphorylation. The subsequent condensation of the myosin from a cortical ‘C’ to a spot C,D may reflect an actomyosin contraction that breaks adhesion bonds by virtue of their attachment to the cortical actin cytoskeleton. Once freed from neighboring cells, the cell rear is retracted forward E by a myosin-independent mechanism that under less adhesive conditions is sufficient by itself to produce rear retraction. Finally, phosphorylation of the posterior myosin spot leads to its dissolution F in preparation for the start of a new cycle.
myosin spot and subsequent proper relocalization requires disassembly of myosin thick filaments regulated by myosin heavy chain phosphorylation (Fig. 7F). ACKNOWLEDGEMENTS This work was supported by NIH grant GM-47330 to JGM. We thank Dr Patricia Clow for collecting some of the wild type movies that were used for quantitative analysis. We are also grateful to Dr Kathy Miller and Dr Gordon Hager for providing lab space for some of the work carried out here. Finally we acknowledge Dr Tatiana Karpova for assistance with the imaging done in the National Cancer Institute Fluorescence Imaging Facility. REFERENCES C PA, MN JG, 1999. In vivo observations of myosin II dynamics support a role in rear retraction. Mol Biol Cell 10: 1309–1323.
DL A, S JA, 1987. Disruption of the Dictyostelium myosin heavy chain gene by homologous recombination. Science 236: 1086–1091. D KW, R I, MN JG, 1995. 3D analysis of cell movement during normal and myosin-II-null cell morphogenesis in Dictyostelium. Dev Biol 167: 118– 129. E TT, L RJ, S JA, 1993. Dictyostelium myosin heavy chain phosphorylation sites regulate myosin filament assembly and localization in vivo. Cell 75: 363–371. E S, J GH, S A, W KL, 1993. Patterns in Dictyostelium discoideum: the role of myosin II in the transition from the unicellular to the multicellular phase. J Cell Sci 104: 457–466. G D, T I, O N, 1982. Interaction between intracellular vacuoles and the cell surface analysed by finite aperture theory interference reflection microscopy. J Cell Sci 54: 287–298. G D, V S, 1982. Substratum wettability and charge influence the spreading of Dictyostelium amoebae and the formation of ultrathin cytoplasmic lamellae. J Cell Sci 54: 255–285. H JA, F RA, 1992. Analysis of G4, a G-protein subunit required for multicellular development in Dictyostelium. Genes Dev 6: 38–49.
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J PY, P PA, W SA, E EL, 1995. A mechanical function of myosin II in cell motility. J Cell Sci 108: 387–393. K KA, MN JG, 1999. Mound cell movement and morphogenesis in Dictyostelium. Dev Biol 208: 416–429. K DA, L WF, 1987. Antisense RNA inactivation of myosin heavy chain gene expression in Dictyostelium discoideum. Science 236: 1081–1086. K DA, S E, 1995. Three-dimensional localization of wild-type and myosin II mutant cells during morphogenesis of Dictyostelium. Dev Biol 170: 434–444. L DA, H AF, 1996. Cell migration: a physically integrated molecular process. Cell 84: 359–369. M TJ, C LP, 1996. Actin-based cell motility and cell locomotion. Cell 84: 371–379. M SL, S JH, S JA 1996. Myosin dynamics in live Dictyostelium cells. Proc. Natl Acad Sci USA 93: 443–446. N R, H C, G G, 1997. Myosin II independent processes in mitotic cells of Dictyostelium discoideum: redistribution of the nuclei, re-arrangement of the actin system and formation of the cleavage furrow. J Cell Sci 110: 123–137. P C, S JA, E EL, 1989. Capping of surface receptors and concomitant cortical tension are generated by conventional myosin. Nature 341: 549–551. P SP, H AF, L DA, 1999. Kinetic model for integrin-mediated adhesion release during cell migration. Ann Biomed Eng 27: 219–235.
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R KM, U TQP, S JA, 1994. Role of highly conserved lysine 130 of myosin motor domain. J Biol Chem 269: 18773–18780. S JH, M SL, R S, Z JH, S JA, 1997. Myosin heavy chain phosphorylation sites regulate myosin localization during cytokinesis in live cells. Mol Biol Cell 8: 2605–2615. S E, K DA, 1995. Mutants lacking myosin II cannot resist forces generated during multicellular morphogenesis. J Cell Sci 108: 1105–1115. S E, K DA, 1996. Dictyostelium cell shape generation requires myosin II. Cell Motil Cytoskeleton 35: 59–67. W D, S DR, K D, L WF, D A, S J, 1988. Cell motility and chemotaxis in Dictyostelium amoebae lacking myosin heavy chain. Dev Biol 128: 164–177. Y S, 2000. Myosin II dynamics and cortical flow during contractile ring formation in Dictyostelium cells. J Cell Biol 154: 137–145. Y S, F Y, 1985. Reversible cyclic AMP-induced change in distribution of myosin thick filaments in Dictyostelium. Nature 314: 194–196. Z JH, C G, S JH, W P, M SL, S JA, 1997. On the role of myosin-II in cytokinesis: division of Dictyostelium cells under adhesive and nonadhesive conditions. Mol Biol Cell 12: 2617–2629.
MYOSIN II REDISTRIBUTION DURING REAR RETRACTION AND THE ROLE OF FILAMENT ASSEMBLY AND DISASSEMBLY GUDRUN KOEHL* and JAMES G. MCNALLY Laboratory of Receptor Biology and Gene Expression, National Cancer Institute, Bethesda, MD 20892, U.S.A. Received 17 July 2001; accepted 14 November 2001
The literature to date suggests a role for myosin II in rear retraction, including evidence that myosin undergoes a characteristic ‘C’-to-spot redistribution at the cell posterior which is associated with retraction. Here we investigate the mechanism of both retraction and the ‘C’-to-spot using Dictyostelium cells containing mutant forms of myosin that affect its polymerization. 3Asp-myosin forms few if any filaments. When 3Asp cells are added to a wild-type mound, the mutant cells move directionally, but rear retraction is markedly delayed, demonstrating that myosin II filaments are essential for efficient retraction. In addition, using a GFP-tagged 3Asp-myosin, we observed a posterior spot pattern associated with retraction, but no cortical ‘C’ pattern preceding it. This suggests that filamentous myosin is required to produce the ‘C’, and that its failure to form results in defective rear retraction. In contrast, an alternate mutant myosin that forms filaments constitutively, 3Ala-myosin, forms ‘Cs’ and then spot patterns at the posterior, but in the interim the spots do not disintegrate. This suggests that spot dissolution occurs by filament depolymerization. In summary our data demonstrate a role for myosin II and the ‘C’-to-spot in efficient rear retraction, and define filament assembly as critical for formation of the ‘C’ and filament disassembly as critical for dissolution of the spot. Published by Elsevier Science Ltd.
K: myosin II; retraction; motility; GFP; time-lapse. A: GFP, green fluorescent protein; MHC, myosin heavy chain
INTRODUCTION The simplest conceptual model for amoeboid cell locomotion involves extension of a cell’s leading edge followed by retraction of its rear (Lauffenburger and Horwitz, 1996; Mitchison and Cramer, 1996). Much data suggest that expansion of the leading edge involves actin polymerization. Less is known about the molecular mechanism of rear retraction, although several lines of evidence have implicated a role for myosin II (hereafter referred to as myosin). *Present address: Klinik und Poliklinik fu¨r Chirurgie, Universita¨tsklinikum Regensburg, Forschungsbau H4, Franz-Josef-Strauss Allee 11, 93053 Regensburg, Germany. To whom correspondence should be addressed: James G. McNally, Laboratory of Receptor Biology and Gene Expression, National Cancer Institute, Bethesda, MD 20892, U.S.A. E-mail: [email protected] 1065–6995/02/$-see front matter
In Dictyostelium, myosin knockout cells cannot translocate when placed on an adhesive substrate (Jay et al., 1995), or within a cell mass where adhesive forces are also high (Eliott et al., 1993; Doolittle et al., 1995). However, the same cells can translocate (slowly) on a glass or agar substrate (Wessels et al., 1988). This substrate-dependent disparity in myosin-null cell motility could reflect a specific requirement for myosin in overcoming high adhesive forces (Jay et al., 1995; Doolittle et al., 1995), which is likely to be a principal task accomplished during retraction of the cell rear (Palecek et al., 1999). But if myosin plays a key role in such retraction, how can myosin-null cells still translocate on less adhesive substrates? This question has led to the suggestion that retraction of the cell rear involves two components, one myosindependent and the other myosin-independent (Jay Published by Elsevier Science Ltd.
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et al., 1995). According to this model, the retractive force generated by the myosin-independent component is strong enough to retract cell extensions under less challenging conditions, as occur for example on a glass substrate, and so myosin-null cells can move on glass. In more challenging environments, as occur on highly adhesive substrates, more retractive force is needed and so myosin-null cells become stuck (Jay et al., 1995; Doolittle et al., 1995). Consistent with a role for myosin in rear retraction is the observation that myosin-null cells fail to form ultrathin lamellae (Jay et al., 1995). As determined by interference reflection microscopy, ultrathin lamellae reflect a very close apposition of the cell’s ventral and dorsal membrane surfaces, analogous to a squeezed tube of toothpaste. In polarized cells crawling on a glass substrate, these lamellae form at the cell posterior during rear retraction (Gingell and Vince, 1982; Gingell et al., 1982). The absence of these lamellae in myosin-null cells suggests firstly that these cells cannot execute the contraction required for membrane squeezing. Secondly since these lamellae are ordinarily associated with retraction, their absence in myosin-null cells suggests abnormal retraction in this mutant. Several myosin localization studies are likewise consistent with a role for myosin in rear retraction. In fixed Dictyostelium cells, myosin concentrates along an arc at the posterior cell cortex (Yumura and Fukui, 1985). A comparable posterior, cortical stain has been observed in living Dictyostelium cells tagged with a GFP-myosin (Clow and McNally, 1999). This arc-like, posterior distribution of GFPmyosin, termed a ‘C’, forms prior to rear retraction and then condenses to a much smaller spot, coincident with retraction of the cell rear. The myosin redistribution correlates with a cell shape change as visualized by combined fluorescence and bright-field microscopy. This so-called ‘C’-to-spot redistribution has been hypothesized to reflect a myosin contraction that contributes to rear retraction (Clow and McNally, 1999). To test the role of the ‘C’-to-spot in rear retraction, we examined both features in two different myosin mutants that affect myosin filament assembly. In Dictyostelium, filament assembly is regulated by the phosphorylation state of three threonines in the tail of the myosin heavy chain (Egelhoff et al., 1993). Monomers are favored when these residues are phosphorylated and filaments are favored when they are dephosphorylated. These two different phosphorylation states have been mimicked in two different site-directed mutants. In one mutant (3Asp), the three threonines have
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been converted to aspartic acid to mimic phosphorylation, and in the other mutant (3Ala) the three threonines have been converted to alanine to mimic dephosphorylation. Each of these sitedirected mutant myosins behaves as expected: 3Asp-myosin is predominantly monomeric and 3Ala-myosin is predominantly filamentous (Egelhoff et al., 1993). By examining cells expressing exclusively one or other of these mutant myosins tagged with GFP (Sabry et al., 1997), we have shown that a defective ‘C’-to-spot can result in defective retraction, and that the formation of a ‘C’ depends on filament assembly while the dissolution of a spot requires filament disassembly.
MATERIALS AND METHODS Cell culture Cells were grown in HL5 medium in 100 mm Petri dishes. The following strains were used: HS1, a myosin II—null (Ruppel et al., 1994); JH10 (Hadwiger and Firtel, 1992), a derivative of KAx 3 as a parent strain for HS1; HS1GFPmyo (Clow and McNally, 1999), a derivative of HS1 containing wild-type GFP-tagged myosin II, and pBIG 3Asp and pBIG 3Ala (Egelhoff et al., 1993), both derivatives of HS1. To generate cells with GFP-3Asp or GFP-3Ala fusion proteins, we used constructs generously provided by Drs James Spudich and Ji-Hong Zang, Stanford University (Sabry et al., 1997). These constructs were used to transform HS1 and several different clones were selected for study. Both the mutant and wild-type GFP-myosins have been previously characterized (Moores et al., 1996; Sabry et al., 1997). Cell preparation for microscopy Cells were collected from Petri dishes by centrifugation and washing in phosphate buffer (Clow and McNally, 1999). To initiate development to the mound stage, the cells were plated on dialysis membrane laid on 2% agar. As mounds formed, the membranes were transferred to a cover slip kept in a humid chamber. For most experiments, labeled cells were mixed with a much higher percentage of unlabeled cells to permit clear identification of the labeled cells within the multicellular mass. In such mixtures, we typically used from 1–5% labeled cells. The GFP-3Asp-myosin was used directly as a cell label, since the GFP stain was largely static and always filled the cytoplasmic compartment. This was not the case for GFP-3Ala-myosin which
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stained only the cell posterior. For tracking of this cell line, we labeled cell cytoplasm with 5-chloromethylfluorescein diacetate (CMF, molecular Probes, Eugene, OR, U.S.A.) to permit a more accurate determination of the cell’s center of mass (Clow and McNally, 1999). Microscopy Images were collected with either a Leica TCS NT SP confocal microscope (Heidelberg, Germany) or a scientific grade cooled CCD camera on a conventional wide-field Olympus IX70 microscope running Deltavision software (Applied Precision, Issaquah, WA, U.S.A.). Depending on the experiment, we used either a 10, 20 or 25 objective lens. For the confocal microscope, the pinhole was set to 1.0 Airy disk unit. Images were collected at intervals from 10–60 s, depending on the experiment. In some cases, multiple focal plane images were collected and then a maximum intensity projected image was created. For additional details of the microscopy, including controls to insure cell viability during imaging, and cell tracking see Clow and McNally (1999). RESULTS 3Asp-myosin was engineered to form few if any myosin filaments, and instead to remain almost exclusively monomeric (Egelhoff et al., 1993). The inability to form thick filaments yields a phenotype in 3Asp-myosin that is virtually identical to a myosin-null. In particular, both mutants arrest development at the first multicellular stage, known as the mound (DeLozanne and Spudich, 1987; Knecht and Loomis, 1987; Egelhoff et al., 1993). At this stage, cell motion defects in both 3Asp and myosin-nulls become pronounced. In each mutant, cells within the mound jiggle in place instead of the vigorous, directed cell motion seen in the wild-type (Doolittle et al., 1995; Clow and McNally, 1999). GFP-3Asp-myosin mutant cells exhibited a comparable developmental phenotype to the conventional 3Asp mutant: development arrested at the mound stage, at which time directional cell motion in the mound ceased, and individual cells instead jiggled largely in place (Fig. 1A). When these GFP-3Asp cells were mixed with predominantly wild-type cells, we found that the mutant cells entered the mound and moved directionally (Fig. 1B), unlike myosin-null cells which drift aimlessly to the mound core or periphery when seeded into wild-type mounds (Clow and McNally, 1999).
Fig. 1. 3Asp cell trajectories within a mound. Individual cells tagged with GFP-3Asp-myosin were mixed with unlabeled cells and then the fluorescent cells tracked from time-lapse images of the mound. A cell position at each time-point is marked by a square, and successive time points are connected by thin lines. the interval between time points if 30 s. Different colors represent different cells. The mound border as determined from a bright field image is shown as a solid black line. (A) Both labeled and unlabeled cells carry 3Asp mutant myosin. The mutant cells translocate only small distances and so lines connecting separate time-points are obscured. The trajectories appear instead as small blobs. This is characteristic of a myosin-null phenotype (Doolittle et al., 1995; Clow and McNally, 1999) and radically different from the rotational cell trajectories observed in the parent strain (Kellerman and McNally, 1999). (B) Labeled (and tracked) cells are 3Asp mutants and unlabeled cells are wild-type. The mound is composed of predominantly wild-type cells (95%). In this largely wild-type mound, the 3Asp mutant cells are capable of executing essentially normal rotational motion. This is in marked contrast to myosin-null cells which drift to the outer edge or inner core of the mound (Clow and McNally, 1999). Scale bars 50 m.
Directional motion in the 3Asp mutant makes it feasible to define a rear retraction. In wild-type cells, rear retraction is consistently associated with a stereotyped ‘C’-to-spot redistribution of GFPmyosin at the cell posterior (Fig. 2A) (see also Clow and McNally, 1999). GFP-3Asp-myosin exhibited one characteristic of the ‘C’-to-spot redistribution of wild-type myosin: it formed a posterior spot (arrows in Fig. 2B,C) that was followed by a rear retraction (Fig. 2B, time-points 320–460; Fig. 2C time-points 180, 200). But compared to wild-type GFP-myosin (Fig. 2A), GFP3Asp-myosin was diffusely and dimly distributed throughout the cytoplasm and not enhanced at the cell cortex (Fig. 2B,C) except for the spot patterns noted above. In particular, cortical ‘C’ patterns were never observed at the cell posterior of GFP3Asp-myosin cells. Thus, both wild-type and GFP-3Asp-myosins exhibited spot patterns associated with retraction, but in wild-type cells, this was preceded by a cortical ‘C’ pattern, whereas in the mutant it was preceded by a more diffuse cytoplasmic stain.
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Fig. 2. Representative timelapse sequences of individual wild-type GFP-myosin (a) and GFP-3Asp-myosin cells (B,C) moving within a wild-type mound. The cells undergo rotational motion in the mound (Fig. 1B), but have been oriented here such that motion in each panel is upward. Time points in seconds are shown below each image. As previously described (Clow and McNally, 1999), the wild-type myosin (A) is initially rather uniformly distributed, but then exhibits a ‘C’-to-spot at the posterior (timepoints 288, 320, 352). The 3Asp GFP mutant myosin is also distributed rather uniformly throughout the cytoplasm. However, at certain time points in the movement sequence, a myosin spot can be discerned (arrows in B,C), although a precursor ‘C’ is never visible. The formation of the myosin spot precedes retraction, as judged by the shortening and fattening of the cell body at subsequent time points (320–460 in B and 180–200 in C). Arrows indicate location of myosin spots in wild-type and 3Asp cells. Dotted lines indicate the cell periphery as judged by contrast enhancement of the cytoplasmic fluorescence. Scale bars 5 m.
Based on its correlation with retraction, we previously hypothesized that the ‘C’-to-spot pattern was mechanistically involved in rear retraction (Clow and McNally, 1999). To test if the absence of a ‘C’ in 3Asp cells had functional consequences, we compared the time between the first appearance of a posterior myosin spot and the time when a rear retraction occurred, for 3Asp cells vs wild-type cells. Included in this analysis were only those cells for which both a GFP-myosin spot and a subsequent retraction could be clearly identified. We found that nearly 60% of 3Asp cells had retraction times that were significantly slower (i.e. more than 3 standard deviations longer) than the average for wild-type cells (Fig. 3). These measurements suggest that both ‘C’ patterns and filamentous myosin (largely absent in the 3Asp mutant) are required for efficient rear retraction. Even though cortical ‘C’ patterns did not form at the cell posterior, other aspects of the wild-type ‘C’-to-spot myosin sequence were not altered in the 3Asp mutant, namely the formation of a myosin
spot and the subsequent dissolution of this spot. This suggests that filamentous myosin was required for formation of the ‘C’, but not dissolution of the spot. To further investigate these possibilities, we examined a second myosin mutant, 3Ala, which forms constitutive thick filaments (Egelhoff et al., 1993). The 3Ala mutant proceeds through development normally (Egelhoff et al., 1993). Consistent with this, we found that cell movement in the 3Ala mound was indistinguishable from wildtype (Fig. 4A). That is, cells exhibited rotational trajectories characteristic of the KAx 3 parent strain (Kellerman and McNally, 1999). Normal, directional movement also continued when the mutant cells were mixed into wild-type mounds (Fig. 4B), suggesting that the mutant cells could compete effectively among wild-type neighbors. Cells containing GFP-3Ala-myosin also behaved like cells containing 3Ala-myosin in these assays, but their patterns of myosin distribution were different from wild-type GFP-myosin. Virtually
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Fig. 3. Rear retraction is significantly less efficient in the 3Asp mutant. To assess the role in rear retraction of myosin and its ‘C’-to-spot redistribution, we compared the time between the spot’s appearance and the subsequent retraction of the cell body in wild-type (white bars) vs 3Asp-myosin mutant cells (black bars). 51 wild-type and 34 3Asp rear retractions were identified from cells moving within the mounds. The measured time intervals for each retraction event are shown on the x axis, and the percent of retractions at that time interval are shown on the y axis. Negative numbers for the wild-type indicate that in some cells retraction occurred before the myosin spot could be detected. The mean of the wild-type distribution is shown (gray arrow) as is the distance encompassing three standard deviations from the mean (gray bar +3SD). Nearly 60% of the 3Asp data fall outside this three standard deviation range, demonstrating a significant difference in rear retraction efficiency in the mutant vs wild-type.
no diffuse cytoplasmic fluorescence was observed. Instead a single large and bright myosin spot was typically found at the posterior cortex of locomoting cells (Fig. 5A,B). ‘C’ patterns appeared to form directly from the myosin in the spot, growing bidirectionally to yield a ‘C’ still superimposed on a spot (a ‘spotted’ ‘C’, Fig. 5B time-points 96 and 120), or in some cells eventually a thickened ‘C’ by itself with only a hint of a superimposed spot (Fig. 5A time point 196). As the ‘C’ condensed to a spot once more, the spot typically persisted until a new ‘C’ pattern eventually formed. In contrast, in wild-type cells completing a rear retraction, the myosin spot ordinarily disappeared entirely leaving a diffuse cytoplasmic distribution of myosin, before a subsequent de novo appearance of a cortical ‘C’ pattern (see Fig. 2A and also Clow and McNally, 1999).
Fig. 4. 3Ala cell trajectories within a mound. Individual 3Ala cells tagged with a fluorescent dye (CMF) were mixed with unlabeled cells and then the fluorescent cells tracked from time lapse images of the mound. A cell position at each timepoint is marked by a square, and successive time-points are connected by thin lines. Different colors represent different cells. The mound border as determined from a bright field image is shown as a solid black line. The interval between timepoints is 120 s in (A) and 104 s in (B). Both labeled and unlabeled cells carry 3Ala mutant myosin. The mutant cells exhibit normal rotational motion characteristic of the parent strain (Kellerman and McNally, 1999). (B) Labeled (and tracked) cells are 3Ala mutants and unlabeled cells are wild-type. The mound is composed of predominantly wild-type cells (95%). In this largely wild-type mound, the 3Ala mutant cells compete effectively and still exhibit normal rotational trajectories. Scale bars 50 m.
To quantify these observations of a persistent myosin spot in GFP 3Ala-myosin cells, we categorized and counted myosin patterns observed at the posterior of both 3Ala and wild-type cells (Fig. 6). The fraction of cells (40% in wild-type vs 33% in 3Ala) exhibiting some form of ‘C’ pattern was similar between the two mutants. This suggests that both the stimulus and the ability to form a ‘C’ pattern were comparable in both cell lines. However, the fraction of cells exhibiting only a posterior myosin spot was radically different: 13% in wildtype vs 67% in the 3Ala mutant (Fig. 6). The dramatic increase of mutant cells with myosin spots was at the expense of two wild-type characteristics that were never observed in the mutant, namely diffuse cytoplasmic fluorescence (15%), or uniform cortical staining (32%). Such wild-type patterns (Fig. 6A–D) appear either before or after a ‘C’-tospot condensation, and persist for variable periods until a new ‘C’-to-spot condensation is initiated (Clow and McNally, 1999). Our quantitative measurements demonstrate that these intermediate patterns are absent in the 3Ala mutant and suggest that they are replaced by myosin spots, presumably because the myosin spot fails to disintegrate normally.
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Fig. 5. Representative timelapse sequences (A) and (B) of GFP-3Ala-myosin cells moving within the mound. The cells undergo rotational motion in the mound (Fig. 4), but have been oriented here such that motion in each panel is upward. Time points in seconds are shown below each image. Virtually all of the GFP mutant myosin collects at the cell posterior where it typically forms a large spot. This spot redistributes into ‘C’-like patterns at the posterior (time points 172–196 in (A) and time points 72–120 in (B) which then condense to form large spots once more (time points 220, 244 in (A) and time points 168, 288 in (B)). At no time is 3Ala-myosin found diffusely distributed throughout the cytoplasm, in marked contrast to wild-type or the 3Asp mutant (Fig. 2). Scale bar 5 m.
Despite the failure of the myosin spot to dissolve, we did not observe significant impairment in cell motility, or more specifically rear retraction. The only obvious defect in the 3Ala cells was that they were less elongated, and in particular less curved at their posterior. This was manifested in ‘C’ patterns that were also less curved or in some cases even straight, forming in the extreme an ‘I’ perpendicular to the cell’s direction of motion (Fig. 6F). DISCUSSION Previous studies have shown that mysosin II is required both for cell division in suspension (DeLozanne and Spudich, 1987; Knecht and Loomis, 1987) and for providing cortical stiffness during interphase (Pasternak et al., 1989; Knecht and Shelden, 1995; Shelden and Knecht, 1995; Shelden and Knecht, 1996). Other studies have provided some evidence of myosin involvement in rear retraction (Yumura and Fukui, 1985; Jay et al., 1995; Clow and McNally, 1999). We now extend these studies with a mutant analysis demonstrating that rear retraction was markedly delayed in cells containing largely monomeric myosin. Retraction times in this 3Asp mutant ranged from twice as long to ten times as long as the average for wild-type. These data suggest that
myosin thick filaments are normally required for efficient rear retraction, as first hypothesized by Jay et al. (1995). In the absence of filaments, we find that retraction is still possible, but inefficient. These observations therefore strongly support the view that myosin II plays an important role in rear retraction. Based on the early myosin knockouts in Dictyostelium, it has repeatedly been stated that myosin is essential for cell division, but not cell motion. The data reported here, plus recent studies of cell division, have changed this picture and demonstrate that both processes, cell division and cell motion, have myosin-dependent and myosinindependent components. Dictyostelium myosinnull cells fail to divide in suspension culture, but in the wild, these cells never grow in suspension and so the physiological significance of this phenotype is unclear. In contrast, the same myosin-null cells undergo fairly normal cytokinesis on an adhesive substrate (Zang et al., 1997; Neujahr et al., 1997), exhibiting mitotic rounding, cell elongation, polar ruffling, furrow ingression and separation of daughter cells. This demonstrates that cytokinesis has a myosin-independent component. Dictyostelium myosin-null cells also exhibit a myosinindependent component to motility. They move sufficiently well to form a cell mass known as the mound. However, this mound arrests development and a fruiting body never forms. Cell movement in
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Fig. 6. 3Ala myosin exists predominantly as posterior spots at the expense of diffuse cytoplasmic or cortical distributions. To quantify differences between 3Ala myosin and wild type myosin, we categorized the different myosin distributions observed in live cell movies. Wild-type GFP myosin exhibited four different patterns: A: ‘cytoplasmic’—a rather uniform distribution throughout the cytoplasm; B: ‘cortical’—a dim enhancement of staining throughout much of the cell membrane (typically much dimmer than the enhancement seen at the posterior cortex and defined as a ‘C’); C: ‘C’—a brightly stained arc-like pattern found at the posterior cortex; D: ‘spot’—a bright puncta located at the posterior cortex. GFP3Ala-myosin exhibited a different assortment of staining patterns; E: ‘C’—a posterior cortical arc very similar to that observed for wild-type myosin; F: ‘I’—a bright, posterior cortical stain that was straight instead of arc-like; G: ‘spotted C’—a ‘C’-like posterior pattern with a spot superimposed; H: ‘spot’—a bright puncta located at the posterior cortex, typically somewhat larger in diameter and brighter than those seen in wild type. Using these categories as a guide, we randomly selected single time point images from five different time-lapse movies of wild-type or 3Ala cells and then classified the myosin distribution in each cell at those selected timepoints. For purposes of this quantification, we classified all of the 3Ala patterns shown in E–G as a ‘C’. We examined 317 wild type cells, and 324 3Ala cells. As shown in the histogram, cytoplasmic and cortical myosin are never observed in 3Ala cells, and appear to be replaced by an increase in spots in this mutant. ‘C’-type patterns account for roughly similar percentages in the two strains. Scale bars 5 m.
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the myosin-null mound is radically defective (Elliot et al., 1993; Doolittle et al., 1995; Clow and McNally, 1999), demonstrating in this case that under certain normal physiological conditions, myosin is vital for cell motion. Our present data show that one key role for myosin in locomotion is rear retraction, and argue more generally that myosin is equally important for cell locomotion as it is for cell division. Our observations suggest that myosin’s role in rear retraction is mediated by its ‘C’-to-spot condensation at the cell posterior. We measured a delay in retraction times in the 3Asp mutant that was calculated as the time interval between the first appearance of the posterior myosin spot and the completion of retraction. That this time was on average much longer in the mutant implies that the condensation from a ‘C’ to a spot has a functional consequence in wild-type, namely subsequent retraction of the rear. In principle, the ‘C’ could provide structural support for the cell posterior during retraction. We think this unlikely given that 3Asp cells which lack a ‘C’ showed no unusual deformation localized to a region at the cell posterior where a ‘C’ should have formed. It seems more likely that the condensation of a ‘C’ to a spot reflects myosin motor function that is important for normal rear retraction. It is of interest that the predominantly monomeric myosin in the 3Asp mutant was capable of forming a spot, but that the mutant cells were nonetheless incapable of efficient retraction. This implies that monomeric myosin responds to the signal to form a spot, but yet cannot produce a retraction. One plausible interpretation is that spot formation occurs via movement over actin, a process that both monomers and thick filaments could accomplish. Such cortical flow of myosin II occurs in cell division (Yumura, 2001) and so might also occur in the transition from a ‘C’ to a spot. Our observation here that 3Asp cells participate in rotational motion in a wild type mound is the first phenotypic difference detected between 3Asp and myosin-null cells. Unlike 3Asp cells, myosin-null cells do not exhibit rotational motion in a wild type mound, but rather drift to the mound periphery or interior (Clow and McNally, 1999). This difference between the two myosin mutants most likely arises due to small myosin multimers that are thought to be present in 3Asp cells (Egelhoff et al., 1993). These multimers may supply sufficient force or structural support to enable the 3Asp cells to participate in rotational motion within the mound. For example, small multimers might enable inefficient retraction as
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observed in the 3Asp cells. These multimers may also supply sufficient cortical integrity to 3Asp cells such that they are stiff enough to compete with wild-type cells and therefore participate in rotational motion within the mound. In this latter scenario, inefficient rear retraction could occur via the myosin-independent retractive force described for myosin-null cells crawling on a substrate (Jay et al., 1995). Regardless of the precise mechanism of inefficient retraction, our data show that 3Asp cells have enough residual myosin function to enable movement in a wild-type mound, but not enough myosin function to permit normal rear retraction. Our current data suggest that the myosin ‘C’-tospot may be regulated in part by the phosphorylation state of myosin. We found that normal dissolution of the myosin spot fails to occur in a mutant, 3Ala, that forms thick filaments constitutively. In this mutant, ‘Cs’ form from the spot, instead of de novo from a diffuse cytoplasmic distribution of myosin, as occurs with wild-type myosin. These observations suggest that ordinarily the myosin spot dissolves due to a signal for thick filament disassembly. In the 3Ala mutant, this signal cannot overcome the tendency of this myosin to form filaments, and so the myosin spot fails to disintegrate upon completion of retraction. These observations parallel earlier studies in which 3Ala-myosin was shown to persist at sites where conA was capped (Egelhoff et al., 1993), also suggesting a failure to dissociate in response to a signal for disassembly. A logical candidate for a disassembly signal is myosin phosphorylation at three threonines in the heavy chain tail. 3Ala cells cannot be phosphorylated at these residues because they have been mutated to alanine, thereby mimicking a continuously dephosphorylated state. As a consequence, 3Ala-myosin should be recalcitrant to disassembly by a kinase specific for these residues, and so should fail to disassemble if this kinase normally triggers disintegration of the myosin spot. Our data, however, do not strictly rule out other, as yet unidentified, signals for thick filament disassembly that are simply overridden by the tendency of 3Ala-myosin to remain filamentous. Despite the inability of the 3Ala-myosin spot to disintegrate, this same myosin can form a cortical ‘C’. In contrast, ‘Cs’ were absent in the complementary mutant, 3Asp, in which myosin is largely monomeric. This suggests that formation of a cortical ‘C’ requires thick filaments. These observations and conclusions are paralleled by studies of myosin localization during cell division.
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Myosin normally accumulates in the cleavage furrow, but GFP-3Asp-myosin fails to localize to the furrow and remains diffusely cytoplasmic during cell division, suggesting also in this case that thick filament formation is required for normal myosin localization (Sabry et al., 1997). In both cases (cell division and rear retraction), a logical candidate for an assembly signal is myosin dephosphorylation at the same three threonine residues in the heavy chain tail. 3Asp cells cannot be dephosphorylated at these sites because these residues have been mutated to aspartic acid, thereby mimicking a continuously phosphorylated state. As a consequence, 3Asp-myosin should be recalcitrant to an assembly signal by a phosphatase that may normally help induce thick filament assembly to form a ‘C’ at the posterior cortex during rear retraction or an accumulation at the cleavage furrow during cytokinesis. Our results with 3Ala however suggest that dephosphorylation of these three threonines is not absolutely required to induce formation of a ‘C’ pattern. 3Ala-myosin cannot be dephosphorylated at these three threonines, but ‘Cs’ still formed in this mutant. Thus some additional signal beyond dephosphorylation at these three residues must induce 3Ala-myosin to leave the spot and spread out to form a ‘C’ along the posterior cortex. The ability of 3Ala cells to form a ‘C’-to-spot pattern could account for their relatively normal rear retractions. In this mutant, the only obvious defect in the ‘C’-to-spot pattern was the failure of the spot to disintegrate normally. One potential consequence of this defect that we could discern was the formation of straighter, less extended ‘C’ patterns in comparison to wild-type ‘Cs’ which presumably form from a more dispersed distribution of myosin monomers at the cell posterior. These constrained ‘C’s’ in 3Ala-myosin might make retraction slightly less efficient in comparison to wild-type, but such a difference would be difficult to detect. In summary, our observations define a role for myosin in rear retraction and suggest a model for this process (Fig. 7). The initial assembly of myosin as a ‘C’ at the posterior cortex requires filaments, and may be regulated in part by myosin heavy chain dephosphorylation (Fig. 7A,B). The condensation to a spot could reflect a myosin contraction over a cortical actin meshwork that helps break adhesive bonds between cells (Fig. 7C,D) or on an adhesive substrate. Once freed from these bonds, the cell rear retracts via a myosin-independent retractive force, such as elasticity of the cytoskeleton (Fig. 7E). The final dissolution of the
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Fig. 7. A working model for myosin’s role and regulation in rear retraction. The panels are shown in clockwise order to emphasize the cyclic nature of the process. The schematic shows a group of cells within a multicellular mass. The center cell undergoes a rear retraction. Within this cell, myosin monomers are indicated by red dots, myosin thick filaments by thick red lines, and cell–cell adhesion bonds by blue lollipops. At the outset, a signal for retraction localized to the posterior yields thick filament assembly at the cortex A,B. This signal may involve myosin dephosphorylation. The subsequent condensation of the myosin from a cortical ‘C’ to a spot C,D may reflect an actomyosin contraction that breaks adhesion bonds by virtue of their attachment to the cortical actin cytoskeleton. Once freed from neighboring cells, the cell rear is retracted forward E by a myosin-independent mechanism that under less adhesive conditions is sufficient by itself to produce rear retraction. Finally, phosphorylation of the posterior myosin spot leads to its dissolution F in preparation for the start of a new cycle.
myosin spot and subsequent proper relocalization requires disassembly of myosin thick filaments regulated by myosin heavy chain phosphorylation (Fig. 7F). ACKNOWLEDGEMENTS This work was supported by NIH grant GM-47330 to JGM. We thank Dr Patricia Clow for collecting some of the wild type movies that were used for quantitative analysis. We are also grateful to Dr Kathy Miller and Dr Gordon Hager for providing lab space for some of the work carried out here. Finally we acknowledge Dr Tatiana Karpova for assistance with the imaging done in the National Cancer Institute Fluorescence Imaging Facility. REFERENCES C PA, MN JG, 1999. In vivo observations of myosin II dynamics support a role in rear retraction. Mol Biol Cell 10: 1309–1323.
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