myosin regulatory light chain and motility

Myosin regulatory light chain and motilityIntroduction In a translocating Dictyosteliumamoeba, myosin II forms thick filaments that localize to the cortex of the cell body (Yumura and Fukui, 1985; Fukui, 1990). Myosin II does not localize to an expanding pseudopod, although it does appear transiently in a retracting pseudopod (Moores et al., 1996). Through the motion analysis of myosin heavy-chain-deficient mutants (DeLozanne and Spudich, 1987; Knecht and Loomis, 1987), it has been demonstrated that mysoin II is necessary for normal cell polarity, cell motility and chemotaxis, and plays a role in the spatial distribution and dynamics of pseudopod extension and retraction (Wessels et al., 1988; Wessels and Soll, 1990; Spudich, 1989; Sheldon and Knecht, 1996). It has been proposed that myosin II plays a role in the suppression of lateral pseudopod formation (Wessels et al., 1988, Wessels et al., 2000b; Stites et al., 1998; Chung and Firtel, 1999), presumably through the generation of cortical tension (Clarke and Spudich, 1974; Fukui and Yumura, 1986; Pasternak et al., 1989; Egelhoff et al., 1996).
During chemotaxis in natural aggregation territories of Dictyostelium, the regulation of lateral pseudopod formation and polarity play key roles in the behavioral responses of cells to the different phases of each natural cAMP wave (Wessels et
al., 1992; Wessels et al., 2000a; Wessels et al., 2000b). Since myosin II is involved in both pseudopod formation and cell polarity, it must play an underlying role in chemotaxis. The rapid addition of the chemoattractant cAMP to cells in buffer results in phosphorylation of the myosin heavy chain (Berlot et al., 1985; Berlot et al., 1987), which in turn results in the depolymerization of myosin II thick filaments (Kuczmarski and Spudich, 1980; Cote and McCrea, 1987; Ravid and Spudich, 1989). Conversion of the three mapped threonine phosphorylation sites in the MHC tail to nonphosphorylatable alanines in the mutant 3XALA results in an increase in myosin II localization to the cell cortex and increased cortical tension (Egelhoff et al., 1996). It also results in behavioral defects in buffer and in spatial gradients of cAMP consistent with an increase in cortical tension, and a significant decrease in chemotactic efficiency (Stites et al., 1998). Together, these results suggest that the phosphorylation/dephosphorylation of MHC plays a critical role in the maintenance of cell shape and motility in buffer, and in chemotaxis in a spatial gradient of cAMP.
Cyclic AMP also stimulates myosin regulatory light chain (RLC) phosphorylation (Kuczmarski and Spudich, 1980; Berlot et al., 1985; Berlot et al., 1987), increasing myosin’s actin-activated Mg2+ ATPase activity (Griffith et al., 1987;
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The myosin regulatory light chain (RLC) of Dictyostelium discoideum is phosphorylated at a single serine site in response to chemoattractant. To investigate the role of the phosphorylation of RLC in both motility and chemotaxis, mutants were generated in which the single phosphorylatable serine was replaced with a nonphosphorylatable alanine. Several independent clones expressing the mutant RLC in the RLC null mutant, mlcR–, were obtained. These S13A mutants were subjected to high resolution computer-assisted motion analysis to assess the basic motile behavior of cells in the absence of a chemotatic signal, and the chemotactic responsiveness of cells to the spatial, temporal and concentration components of natural cAMP waves. In the absence of a cAMP signal, mutant cells formed lateral pseudopods less frequently and crawled faster than wild-type cells. In a spatial gradient of cAMP, mutant cells chemotaxed more efficiently than wild-type cells. In the front of simulated temporal and natural waves
of cAMP, mutant cells responded normally by suppressing lateral pseudopod formation. However, unlike wild-type cells, mutant cells did not lose cellular polarity at the peak and in the back of either wave. Since depolarization at the peak and in the descending phase of the natural wave is necessary for efficient chemotaxis, this deficiency resulted in a decrease in the capacity of S13A mutant cells to track natural cAMP waves relayed by wild-type cells, and in the fragmentation of streams late in mutant cell aggregation. These results reveal a regulatory pathway induced by the peak and back of the chemotactic wave that alters RLC phosphorylation and leads to cellular depolarization. We suggest that depolarization requires myosin II rearrangement in the cortex facilitated by RLC phosphorylation, which increases myosin motor function.
Key words: Myosin light chain, Myosin phosphorylation, Cell motility, Chemotaxis, Dictyostelium discoideum
Summary
Phosphorylation of the myosin regulatory light chain plays a role in motility and polarity during Dictyostelium chemotaxis Hui Zhang 1, Deborah Wessels 1, Petra Fey 2, Karla Daniels 1, Rex L. Chisholm 2 and David R. Soll 1,* 1Department of Biological Sciences, University of Iowa, Iowa City, Iowa 52242, USA 2Department of Cell and Molecular Biology, Northwestern University Medical School, Chicago, IL 60611, USA *Author for correspondence (e-mail: [email protected])
Accepted 29 January 2002 Journal of Cell Science 115, 1733-1747 (2002) © The Company of Biologists Ltd
Research Article
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Trybus, 1989). To investigate the role of RLC phosphorylation in motility and chemotaxis, we expressed either wild-type RLC or RLC in which serine 13 was substituted with alanine (S13A) in an RLC null mutant (Ostrow et al., 1994). Myosin II from mutant S13A cells exhibited only 30% of the actin-activated Mg2+ATPase activity of wild-type myosin II (Ostrow et al., 1994). Nevertheless, S13A cells underwent cell division, localized myosin II to the cortex of locomoting cells and formed fruiting bodies (Ostrow et al., 1994), suggesting that RLC phosphporylation was not essential for growth or cytoskeletal organization during locomotion and development.
Under the assumption that a modification of myosin II mediated through occupancy of the cAMP receptor must play a role in chemotaxis, we subjected S13A cells to high resolution computer-assisted motion analysis, employing a set of experimental protocols that tested, first, the basic motile behavior of mutant cells in the absence of an extracellular cAMP signal and, second, the responses of mutant cells to the different spatial, temporal and concentration components of the natural cAMP wave (Fig. 1). The results of these experiments demonstrate that RLC phosphorylation plays a role in the basic motile behavior of cells in the absence of an extracellular chemotactic signal, and in the normal response of cells to the peak and back of a natural chemotactic wave. The incapacity of mutant S13A cells to phosphorylate RLC at the peak and in the back of the wave results in less efficient chemotaxis in natural waves early in mutant cell aggregation, and to the fragmentation of streams late in aggregation. These results support a model of chemotactic regulation in which independent regulatory pathways emanating from the distinct phases of the natural chemotactic wave elicit a sequence of specific cellular behaviors that together represent the natural chemotactic response.
Materials and Methods Origin of strains The RLC null mutant mlcR– was generated from parental strain JH10 by targeting the RLC gene (mlcR) as previously described (Chen et al., 1994). The S13A mutants, which contained unphosphorylatable RLC, were generated by transforming mlcR– with the Dictyostelium integrating vector pBVN5115, which contained a mutated version of RLC [in which Ser13 (TCA) was changed to Ala (GCC), under the regulation of the actin 15 promoter] and a neomycin resistance gene for G418 selection of transformed cells (Ostrow et al., 1994). Three S13A mutants were generated in independent transformations, S13A- 1, S13A-2 and S13A-3. A control strain, WT-res, representing the RLC deletion strain rescued with wild-type RLC was generated by transforming mlcR– with the Dictyostelium integrating vector pBVN5133, which contained a wild-type version of RLC under the regulation of the actin 15 promoter, and a neomycin resistance gene for the selection of transformed cells (Ostrow et al., 1994). It was demonstrated that the WT-res RLC, but not the S13A RLC, could be phosphorylated with the myosin regulatory light chain kinase MLCK in vitro (Ostrow et al., 1994). In the computer-assisted analysis of mutant behavior, the first characterizations were performed in detail on mutant strain S13A-1, and aberrant behaviors then verified less rigorously in mutant strains S13A-2 and S13A-3.
Maintenance and development of control, mutant and rescued strains Spores of JH10, S13A and WT-res strains were frozen in 10% glycerol
and stored at –80°C. For experimental purposes, cultures were generated from spores every three weeks (Sussman, 1987). Cells were initially grown in HL-5 medium alone for two days, then in HL-5 medium containing 10 µg per ml of G418, to a final cell concentrations of 2×106 per ml. To initiate development, cells were washed in buffered salt solution (BSS) containing 20 mM KCl, 2.5 mM MgCl2 and 20 mM KH2PO4 (pH 6.4) and dispersed on a black filter pad saturated with BSS at a density of 5×106 cells per cm2 (Soll, 1987). For all analyses of single cell behavior, except those in which the developmental regulation of motility was monitored, cells were harvested at the ripple stage, which in dense cultures represents the onset of aggregation (Soll, 1979), the time at which Dictyostelium amoebae attain their highest average velocity (Varnum et al., 1986).
Analysis of the basic motile behavior of mutant cells (protocol 1, Fig. 1B) The behavior of cells in the absence of an extracellular cAMP signal, which we will refer to henceforth as the ‘basic motile behavior’ of a cell, was analyzed according to methods previously described (Varnum et al., 1985; Varnum-Finney et al., 1987a; Wessels et al., 2000a; Wessels et al., 2000b). In brief, 1.1 ml of dilute cell suspension were inoculated into a Sykes-Moore chamber (Bellco Glass, Vineland, NJ). The chamber was then inverted and positioned on the stage of an upright microscope fitted with long-range objectives and condenser. For motion analysis, cell behavior was either video-recorded or digitized directly through a 10× objective or 25× objective. The chamber was perfused with BSS at a rate that replaced the liquid volume every 15 seconds to ensure that cells did not condition the medium. This flow rate was demonstrated not to interfere with normal cellular translocation.
Analysis of mutant cell chemotaxis in a spatial gradient of cAMP (protocol 2, Fig. 1B) The motile behavior of cells in a spatial gradient of cAMP generated in a single cell spatial gradient chamber (Zigmond, 1977) was analyzed according to methods previously described (Varnum and Soll, 1984; Varnum-Finney et al., 1987b; Wessels et al., 2000a; Wessels et al., 2000b). In brief, cells were dispersed on the bridge of a Plexiglas gradient chamber, in which one of the two troughs bordering the bridge contained BSS (sink) and the other trough contained BSS plus 10–6 M cAMP (source). Cells were video- recorded through a 25× objective with bright field optics for a 10 minute period following an initial 5 minute incubation period necessary for establishing a steep gradient (Shutt et al., 1998).
Analysis of mutant cell behavior in temporal waves of cAMP (protocol 3, Fig. 1B) The motile behavior of cells in a series of temporal waves of cAMP, which simulate the temporal dynamics of natural waves in the absence of spatial gradients, was analyzed according to methods previously described (Varnum et al., 1985; Varnum-Finney et al., 1987a; Wessels et al., 2000b). In brief, cells were inoculated into a Sykes-Moore chamber as described for the analysis of cell behavior in buffer. To generate temporal waves of cAMP, cells were perfused with increasing, then decreasing, temporal gradients of cAMP, and the process repeated three times. Cells were first perfused with 5 ml of BSS, then with 2 ml of BSS containing 7.8×10–9 M cAMP over a 30 second period. At 30 second intervals thereafter, cells were perfused with 2 ml of a new solution containing twice the cAMP concentration of the preceding solution, terminating at 10–6 M cAMP, the last step in the increasing phase. Cells were then treated with 2 ml increments of BSS containing half the previous concentration of cAMP at 30 second intervals, terminating at 10–8 M cAMP. The second, third and fourth waves were generated in a similar fashion. The periodicity of
Journal of Cell Science 115 (8)
1735Myosin regulatory light chain and motility
simulated temporal waves was, therefore, 7 minutes. Fields of cells were video-recorded or directly digitized through a 10× or 25× objective. The concentration of cAMP in the chamber through the four simulated waves was assessed by fluorescent dye experiments as previously described (Wessels et al., 2000b).
Analysis of mutant cell behavior after the rapid addition of 10–6
M cAMP (protocol 4, Fig. 1B) The motile behavior of cells before and after the rapid addition of 10–6
M cAMP was analyzed according to methods described above for analysis of the basic motile behavior in buffer with one modification. That is, following perfusion for 10 minutes with BSS, the perfusion solution was rapidly switched to BSS containing 10–6 M cAMP. The concentration of cAMP in the Sykes-Moore chamber was assessed by fluorescent dye experiments as previously described (Wessels et al., 2000b). Cell behavior prior to and after addition of cAMP was continuously video recorded or digitized directly through a 10× or 25× objective.
Analysis of mutant cell behavior in self-generated waves of cAMP (protocol 5, Fig. 1B) The motile behavior of cells in self- generated waves of cAMP was analyzed according to methods previously described (Escalante et al., 1997), with the exception that the plastic surface of the tissue culture dish was not coated with agar (Wessels et al., 2000b). In brief, 2 ml of a cell suspension (2.4×106 per ml BSS) were dispersed on a 35 mm tissue culture dish. After 30 minutes of incubation, 1.0 ml of fluid was withdrawn and the dish placed on the stage of an inverted microscope. Cell behavior was continuously video-recorded or directly digitized through a 10× objective. Individual cells positioned in the same area of the field that exhibited no cell-cell contacts were selected for analysis. For streaming experiments late in aggregation, a 2.5× objective was employed.
Analysis of mutant cell behavior in wild-type waves of cAMP (protocol 6, Fig. 1B). The motile behavior of mutant cells in natural waves of cAMP generated by wild-type cells was analyzed according to methods previously described (Wessels et al., 2000a; Wessels et al., 2000b). In
brief, S13A cells were stained with the vital dye DiI (Molecular Probes, Eugene, OR), mixed with a majority of unstained JH10 cells, at a ratio of 1:9, and 2 ml of the cell mixture (2.4×106 per ml BSS) dispersed on a 35 mm tissue culture dish. After 30 minutes, 1 ml of fluid was withdrawn and the dish positioned on the stage of an Axiovert 100STV Zeiss microscope equipped for epifluorescent analysis. Cell behavior was analyzed with brightfield and fluorescence microscopy according to methods previously described (Wessels et al., 2000b). In a control experiment, unstained mutant cells were mixed with stained wild-type cells.
Experimental ProtocolTested Behavior
PHASE A
PHASE B
PHASE C
PHASE D
Front of Wave: Increasing temporal and increasing spatial gradients of cAMP
Back of Wave: Decreasing temporal and decreasing spatial gradients of cAMP
Peak
Re-extension of pseudopods in random directions; maintenance of depolarized state; no net movement in any direction (response to decreasing temporal gradient of cAMP)
Cell depolarization, cessation of translocation (response to peak concentration of cAMP).
Rapid, directed translocation towards source of wave; suppression of lateral pseudopods (response to increasing temporal gradient of cAMP.)
Direction towards aggregation center estab l ished, ce l lu lar polarization (response to positive spatial gradient of cAMP).
A. Behavioral responses to the different phases of the natural wave.
B. Experimental protocols for determining the behavioral defects of S13A mutants in basic motile behavior and chemotaxis.
1. Perfusion of amoebae with buffer in perfusion chamber.
2. Spatial gradient of cAMP generated on the bridge of a single cell spatial gradient chamber.
3. Exposure to a sequence of interspersed increasing and decreasing temporal gradients of cAMP that mimic the temporal dynamics of natural waves in the absence of established spatial gradients.
4. Rapid addition of 10-6M cAMP to cells crawling in buffer.
5. Aggregation in a dilute monolayer on a plastic surface.
6. Aggregation in which fluorescently stained mutant cells are mixed with unstained wild type cells in a 1 to 9 ratio.
1. Basic motile behavior in the absence of an extracellular cAMP signal.
2. Capacity to assess the direction of a spatial gradient of cAMP (Phase A).
3. Behavior in the middle of the front of the wave in response to increasing temporal gradient of cAMP (Phase B), the response to the concentration of cAMP at the peak of the wave (Phase C) and the behavior in the back of the wave (Phase D).
x
4. Response to the concentration of cAMP at the peak of the wave (Phase C).
5. Behavior in all phases (A, B, C and D) of self-generated natural waves of cAMP.
6. Behavior in response to all phases (A, B, C and D) of natural waves generated by wild type cells.
Direction of relayed wave
center
Fig. 1. (A) The behavioral responses of wild-type cells to the spatial, temporal and concentration components of the different phases of the natural cAMP wave, derived from results obtained in prior studies (Varnum-Finney et al., 1987a; Varnum-Finney et al., 1987b; Wessels et al., 1992; Wessels et al., 2000b). (B) Protocols used in this study to determine the behavioral defects of S13A mutant cells.
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Computer-assisted analysis of cell motility Video images were digitized at a rate of 15 frames per minute (i.e. at 4 second intervals) onto the hard disc of a Macintosh G4 computer (Apple Computers, Cupertino, CA) equipped with a Data Translation framegrabber board (Data Translation Inc., Marlboro, MA) and 2D- DIAS software (Soll, 1995; Soll and Voss, 1998). Perimeters were automatically outlined and converted to beta-spline replacement images (Soll, 1995; Soll and Voss, 1998; Soll et al., 2000). Motility parameters were computed from centroid positions and morphology parameters from perimeter contours (Soll, 1995). Instantaneous velocity of a cell in frame n was computed by drawing a line from the centroid in frame n-1 to the centroid in frame n+1 and dividing the
length of the line by twice the interval time (15 seconds) between frames. For simplicity, instantaneous velocity will be referred to simply as ‘velocity’ in the text. Directional change was computed as the direction in the interval (n-1, n) minus the direction in the interval (n, n+1). Directional change values >180° were subtracted from 360°, providing a positive value between 0° and 180°.
Difference pictures were generated by superimposing the image in frame n on the image in frame n-1. The regions of the cell image in frame n not overlapping the cell image in frame n-1 were considered the ‘expansion zones’. The summed area in the expansion zones of a difference picture divided by the total cell area in frame n and multiplied by 100 represents positive flow. The period between overlapping images in difference pictures was 1 minute. This parameter provides a measure of cellular translocation that is independent of cell centroid movement (Soll, 1995; Soll and Voss, 1998).
Maximum length was the longest chord between any two points along the perimeter of a cell. Roundness was computed by the formula 10×4π×area/perimeter2. Chemotactic index (CI) in a spatial gradient of chemoattractant was the net distance moved towards the source of chemoattractant divided by the total distance moved in the same time period. Percent positive chemotaxis was the proportion of the cell population exhibiting a positive CI over the period of analysis. In measuring the frequency of lateral pseudopod formation, a lateral pseudopod was considered to be a projection formed from the main axis of translocation at an angle ≥30° that attained a minimum of 5% total cell area and initially contained nonparticulate cytoplasm. The main axis of translocation was determined by drawing a line between the centroid of the cell in the frame 15 seconds earlier and the centroid of the cell in the present frame (Wessels et al., 1996; Wessels et al., 2000a; Wessels et al., 2000b).
For the analysis of instantaneous velocity as a function of developmental time, all cells in the population were motion analyzed. For all other experiments, motion analysis parameters were computed at 4 second intervals only for those cells crawling at instantaneous velocities above 3 µm per minute. For all strains in all tested situations, this represented over 70% of each population.
Myosin II localization Cells were stained for myosin II according to methods previously described (Wessels et al., 2000b). In brief, cells were subjected to three simulated temporal waves of cAMP. Midway through the increasing phase, at the peak and midway through the decreasing phase of the last of these waves in independent cultures, the chambers were perfused with 4% paraformaldehyde in phosphate buffer solution supplemented with 0.01% saponin. After an antigen retrieval
0 2 4 6 8 10 12 0
2
1
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6
8
10
9
7
5
3
)
Fig. 2.Motility is developmentally regulated in the myosin regulatory light chain phosphorylation mutant S13A. Cells were removed from developing JH10, S13A-1 and WT-res cultures at noted times, dispersed on the wall of a perfusion chamber and analyzed for cell velocity over a 10 minute period while perfused with buffer. The mean instantaneous velocity (Inst. Vel.) was computed at each time point from the average instantaneous velocity of 20 to 30 amoebae selected at random without a velocity threshold. Results similar to those for S13A-1 cells were obtained for cells of the independent mutant strains S13A-2 and S13A-3.
Fig. 3.Mutant S13A-1 cells retract anterior pseudopods and extend lateral pseudopods in a manner similar to that of wild-type JH10 cells. Retraction of the original anterior pseudopod and extension of a new lateral pseudopod over a 24 second period for a representative JH10 (A) and S13A (B) cell imaged through differential interference contrast optics. a, original anterior pseudopod, at time zero, and new anterior pseudopod at 24 seconds; u, uropod. Arrows indicate direction of retraction of original anterior pseudopod and direction of expansion of new lateral pseudopod for both cell types. Time is indicated in seconds in upper left hand corners of panels. Similar results were obtained for cells of strains WT-res, S13A-2 and S13A-3.
procedure (Wessels et al., 2000b), cells were incubated with rabbit anti-myosin II antibody, a generous gift of Arturo DeLozanne (University of Texas, Austin, TX), and stained with FITC-labeled anti- rabbit antibody (Jackson ImmunoResearch, West Grove, PA). DIC and confocal images were captured at 1 µm intervals beginning at the substratum with a Zeiss 510 laser-scanning confocal microscope in the Central Microscopy Facility at the University of Iowa. To measure the distribution of myosin II across a cell, intensity plots were derived along a line that did not cross the cell nucleus, using Zeiss 510 software.
Results Strains analyzed To obtain mutant cells containing a myosin regulatory light chain (RLC) that cannot be phosphorylated, the RLC deletion mutant mlcR–, derived from the parental wild-type strain JH10, was rescued with a mutated form of RLC, S13A, in which serine 13 (TCA) was substituted with alanine (GCC), under the control of the constitutive actin 15 promoter (Ostrow et al., 1994). Three independent S13A mutants were generated, S13A-1, S13A-2 and S13A-3. In addition, the mlcRmutant was rescued with wild-type RLC to generate the control strain WT- res. The parent strain JH10, the mutant strains S13A-1, S13A- 2 and S13A-3, and the rescued strain WT-res were then analyzed for basic motile behavior and chemotaxis. In every condition in which aberrant behavior was demonstrated in strain S13A-1, it was also demonstrated in the additional mutants S13A-2 and S13A-3. For simplicity, mutant S13A-1 will be referred to as strain S13A in the Results, and as strain S13A-1 in the figure legends and tables, where corroborative results with strain S13A-2 and S13A-3 are reported.
The basic motile behavior of mutant cells is aberrant During Dictyosteliumdevelopment, the velocity of individual wild-type amoebae increases to a maximum at the onset of aggregation (Varnum et al., 1986; Wessels et al., 2000b; Escalante et al., 1997). To test whether mutant cells behaved similarly, JH10, S13A and WT-res cells were removed from developing cultures at various times and analyzed for mean instantaneous velocity in buffer. All three strains attained maximum instantaneous velocity at the onset of aggregation, between 8 and 9 hours of development (Fig. 2), demonstrating that the developmental regulation of velocity was intact in the absence of RLC phosphorylation. However, the maximum instantaneous velocity achieved by S13A cells at the onset of aggregation was at least 30% higher than that of either JH10 or WT-res cells (Fig. 2). The difference in both cases was significant (P<0.01, Student t-test). To address the possibility that the increase simply reflected the proportion of motile cells, the mean instantaneous velocity of only those cells moving faster than 3 µm per minute was computed for all three cell lines. This velocity threshold has been used previously to eliminate cells not persistently translocating (Wessels et al., 1996; Wessels et al., 2000a; Wessels et al., 2000b). When applied, the peak velocity (±s.d.) of JH10, S13A and WT-res cells was 8.3±5.6 (n=46), 10.3±5.3 (n=38) and 7.4±4.6 (n=52) µm per minute, respectively. Again, the peak of S13A cells was 24% higher than that of JH10 cells and 39% higher than that of WT-res cells. These differences were significant (P<0.02, Student t-test).
Cell velocity can be affected by the rate of pseudopod expansion (Cox et al., 1992; Cox et al., 1996) and the frequency of lateral pseudopod formation, the latter correlating with the frequency of turning (Varnum-Finney et al., 1987b). In buffer, the directional change parameter, an indicator of turning frequency (Soll, 1995; Soll and Voss, 1998), was
Table 1. Lateral pseudopod formation by cells crawling in buffer or in a spatial gradient of cAMP* Average frequency of
Number 0 Lateral pseudopods 1 Lateral pseudopod 2 Lateral pseudopods >2 lateral pseudopods lateral pseudopod Condition Cell type of cells per 10 min (%)† per 10 min (%)† per 10 min (%)† per 10 min (%)† per cell per 10 min
Buffer JH10 24 4 8 8 79 3.4 S13A 24 29 29 38 4 1.2
Spatial gradient JH10 20 40 45 15 0 0.75 S13A 28 64 36 0 0 0.36
*Cells were imaged at 25× magnification. For the definition of a lateral pseudopod, see Materials and Methods. Cells were analyzed in all cases for 10 minutes. †A Chi square test was performed between JH10 and S13A cells on the combined data of the four categories of lateral pseudopod formation. The difference
between JH10 and S13A cells both in buffer and spatial gradients of cAMP was found to be highly significant (10–12 and 4×10–3, respectively).
Table 2. Motility, dynamic morphology and chemotaxis parameters in a spatial gradient of cAMP Instantaneous Directional Percent
velocity Positive flow change Area Maximum length positive Chemotactic Cell type Cell number (µm/min) (%/min) (deg./min) (µm2) (µm) Roundness (%) chemotaxis† index
JH10 20 7.7±4.6 8.0±4.6 25±13 100±18 17±3 68±11 90 0.53±0.32 S13A 28 13.6±5.1 17.6±19.4 15±9 88±17 18±3 57±8 100 0.73±0.24 WT-res 29 9.9±6.8 9.8±6.8 22±15 95±27 18±5 63±14 83 0.49±0.47 P values* JH10 vs S13A 0.0001 0.006 0.003 0.03 0.04 0.00001 0.001 S13 vs WT-res NS(0.01) NS(0.03) 0.001 NS NS 0.04 NS(0.02)
*Significance was determined by the Student t-test for all measured parameters except ‘percent positive chemotaxis’. A P value greater than 0.05 was considered non-significant (NS), but values close to 0.05 are shown in parenthesis.
†A Chi-square test found the difference between JH10 and S13A close to significant and the difference between WT-res and S13A significant.
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consistently 10-20% lower in S13A cells than JH10 cells translocating in buffer. Increased turning can depress instantaneous velocity, while decreased turning can elevate it. We tested whether the increase in velocity of S13A cells in buffer was accompanied by a decrease in lateral pseudopod formation by counting the number of lateral pseudopods formed over a 10 minute period. JH10 and S13A cells retracted old anterior pseudopods and extended new lateral pseudopods in a qualitatively similar manner (Fig. 3A and B, respectively). However, S13A cells formed lateral pseudopods at only one- third the rate of JH10 cells (Table 1). These results suggest that the increase in velocity of S13A cells in buffer may be due, at least in part, to the decreased rate of lateral pseudopod formation.
Mutant cells chemotax efficiently in a spatial gradient of cAMP To test whether S13A cells chemotax efficiently in a spatial gradient of cAMP, the mechanism presumed to be basic to the directional decision in phase A of the natural wave (Fig. 1A), the behavior of JH10, S13A, and WT-res cells were compared in spatial gradients of cAMP generated in a chamber consisting of a bridge that supports the cells, and two bordering troughs, one filled with attractant (the source) and the other with buffer (the sink) (Zigmond, 1977; Varnum and Soll, 1984; Shutt et al., 1998). Cell behavior was analyzed in a 10 minute time window (the 5-15 minute period following filling of the chamber troughs), when the evolving gradient of cAMP across the bridge elicits the maximum chemotactic response (Shutt et al., 1998). S13A cells translocated in spatial gradients of cAMP at a velocity significantly higher than that of JH10 or WT-res cells (Table 2). S13A cells also exhibited a mean positive flow value, approximately twice that of either JH10 or WT-res cells (Table 2). Positive flow is a measure of area displacement in a 4 second period that provides a measure of translocation that is independent of the cell centroid (Soll, 1995). Furthermore, S13A cells exhibited a directional change
parameter 60% that of JH10 cells and 70% that of WT-res cells (Table 2), indicating that S13A cells turned less frequently than the other two cell types during chemotaxis. Finally, both the mean chemotactic index and the proportion of the population exhibiting a positive chemotactic index (percent positive chemotaxis) were higher in S13A cells (Table 2). The higher mean chemotactic index (Table 2) was due to the very high proportion of S13A cells with chemotactic indices >0.8, as demonstrated in the histogram in Fig. 4.
The differences in both velocity and chemotactic efficiency were reflected in perimeter tracks. The perimeter tracks of the three S13A cells with the highest chemotactic indices in a spatial gradient of cAMP were more persistent in the direction of the source of chemoattractant and included fewer sharp turns (Fig. 5B) than the perimeter tracks of the three JH10 cells (Fig. 5A) and the three WT-res cells (Fig. 5C) with the highest
Fig. 4.Mutant S13A-1 cells chemotax more efficiently than JH10 cells in a spatial gradient of cAMP. A histogram of chemotactic indices indicates that S13A cells attain high CIs (>0.8-1.0) more frequently than JH10 cells or WT-res cells. The number of JH10, S13A-1 and WT-res cells analyzed was 20, 28 and 29, respectively. Results similar to those for S13A-1 cells were obtained for S13A-2 and S13A-3 cells.
Fig. 5.Mutant S13A-1 cells migrate faster and with fewer turns in a spatial gradient of cAMP. Computer-generated tracks are presented of the three JH10 (A), S13A-1 (B) and WT-res (C) cells with the highest chemotactic indices. Cells were selected from 20, 28 and 29 analyzed cells, respectively. Cell perimeters are drawn every 4 seconds. Results similar to those for S13A-1 cells were obtained for strains S13A-2 and S13A-3 cells.
chemotactic indices. In addition, the perimeters of S13A cells (Fig. 5B) were less tightly stacked than those of JH10 cells (Fig. 5A) or WT-res cells (Fig. 5C), indicating higher average velocities. The reduction in sharp turns in the S13A perimeter tracks suggested that, as in buffer, S13A cells formed fewer lateral pseudopods per unit time in a spatial gradient than either JH10 or Wt-res cells. To test this prediction, direct counts were made of the number of lateral pseudopods formed in a 10 minute period. The results demonstrated that the frequency of lateral pseudopod formation by S13A cells was half that of JH10 cells during chemotaxis in a spatial gradient of cAMP (Table 1).
Mutant cells respond abnormally to the peak and back of temporal waves The results obtained in spatial gradient chambers suggest that mutant cells, which cannot phosphorylate the myosin regulatory light chain in response to a cAMP signal, can still orient and chemotax efficiently up a spatial gradient of cAMP, the presumed mechanism for orientation and polarization in phase A of the natural wave (Fig. 1A). The behavior of cells in phases B, C and D of the natural wave, however, are in response to the temporal and concentration characteristics of the wave (Wessels et al., 1992) (Fig. 1A). In response to the increasing temporal gradient in the front of each wave (phase B), cells suppress lateral pseudopods (Varnum-Finney et al., 1987a; Wessels et al., 1992; Wessels et al., 2000b) and move in a highly persistent and directional manner towards the
aggregation center. When cells encounter the high concentration of cAMP at the peak of the wave (phase C), they round up, lose polarity and stop translocating. Finally, in response to the decreasing temporal gradient in the back of the wave (phase D), cells again extend pseudopods, but remain relatively apolar, resulting in little net movement in any direction (Varnum-Finney et al., 1987a; Wessels et al., 1992, 2000b). These responses restrict the movement of cells towards the aggregation center during natural aggregation to phase B of the natural wave. The responses to the temporal and concentration components of phases B, C and D of the natural wave are readily assessed by subjecting cells to sequential increasing and decreasing gradients of cAMP generated in a purfusion chamber (Fig. 1B, protocol 3) (Varnum et al., 1985; Varnum-Finney et al., 1987a; Wessels et al., 1992; Wessels et al., 2000b).
Because of the round shape of the chamber and perfusion rate, the temporal waves are generated in the absence of spatial gradients. In Fig. 6A and B, the instantaneous velocity of a representative JH10 and S13A cell, respectively, and the estimated concentration of cAMP, are co-plotted as functions of time through four successive simulated temporal waves. The average velocity of the representative JH10 cell (Fig. 6A) and the S13A cell (Fig. 6B) remained depressed through the first simulated temporal wave, increased at the onset of the second wave, peaked at the midpoint of the increasing phase of the second wave, decreased at the peak of the second wave and remained depressed through the remaining decreasing phase of the second wave. Behaviors in the different phases of the third
A. JH10
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Fig. 6.Mutant S13A cells respond to a sequence of simulated temporal waves of cAMP generated in the absence of spatial gradients by increasing and decreasing velocity in a manner similar to wild-type JH10 cells. The instantaneous velocity is plotted as a function of time for a representative JH10 cell (A) and a representative S13A-1 cell (B) during four simulated waves. The estimated cAMP concentration, measured in dye experiments (Wessels et al., 2000b), is presented as a function of time through the four waves. Note that the velocity of neither JH10 nor S13A-1 cells increases in the front of the first simulated wave, a result previously reported for wild-type cells (Varnum et al., 1985). Instantaneous velocity was measured at 5 second intervals. Instantaneous velocity plots were smoothed 10 times with Tukey windows of 10, 20, 40, 20 and 10. Results similar to those for the representative JH10 and S13A-1 cells were obtained for nine additional cells of each respective cell line. Results similar to those for S13A-1 cells were obtained with S13A-2 cells.
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and fourth waves were similar to those in the second wave. Similar results were obtained with WT-res cells (data not shown). The velocity data suggest, therefore, that S13A cells respond normally to the temporal dynamics of the chemotactic wave. However, scrutiny of cell shape during the different
phases of the temporal wave revealed that the instantaneous velocity plots did not provide the full story. In simulated temporal wave two to four, JH10 and WT-res cells exhibited the sequence of shape changes previously reported (Wessels et al., 2000b). In the increasing temporal gradient in the front of
Fig. 7.Cell morphology and myosin II localization during the different phases of a simulated temporal wave of cAMP generated in the absence of a spatial gradient. A to C, D to F, and G to I represent differential interference contrast (DIC) microscopy images of representative JH10 cells fixed in the front, peak and back, respectively, of a simulated temporal wave of cAMP. A′ to C′, D′ to F′ and G′ to I′ are DIC images of representative S13A-1 cells fixed in the front, peak and back, respectively, of a simulated temporal wave of cAMP. J and K are representative JH10 cells in the front and the back, respectively, of a simulated temporal wave of cAMP stained with anti-myosin II antibody (first panel in each set) and scanned along the white line shown in the first panel for staining (pixel) intensity (second panel in each set). J′ and K′ are representative S13A-1 cells in the front and the back, respectively, of a simulated temporal wave of cAMP stained and analyzed in a fashion similar to the JH10 cells in panels J and K. Over 100 JH10 and 100 S13A-1 cells were analyzed for morphology in the front, peak and back of simulated waves, and found to exhibit the morphologies of the representative cells in this figure. Nine additional JH10 and S13A-1 cells in the front and back of simulated temporal waves were found to exhibit the distribution of myosin demonstrated for the representative cells in the figure.
waves two to four (phase B), JH10 cells were highly elongate with a dominant anterior pseudopod (Fig. 7A-C). At the peak of the wave (phase C), JH10 rounded up, exhibiting a loss of polarity (Fig. 7D-F). In the decreasing temporal gradient in the back of the wave (phase D), JH10 cells again extended pseudopods, but in random directions, reflecting a lack of polarity (Fig. 7G-I). WT-res cells progressed through shape changes identical to those of JH10 cells in phases B, C and D (data not shown). In the increasing temporal gradients in the front of waves two to four (phase B), S13A cells were elongate on average (Fig. 7A′-C′), similar to JH10 (Fig. 7A-C) and WT- res cells (data not shown). However, the majority of S13A cells were still elongate and polar at the peak of the wave (phase C), each with a dominant anterior pseudopod (Fig. 7D′-F′), and remained elongate and polar during the decreasing phase of the wave (phase D) (Fig. 7G′-I′).
Therefore, although S13A cells exhibited a decrease in instantaneous velocity at the peak and in the decreasing phase of the second to fourth simulated temporal wave in a series, they did not undergo the normally associated loss of cellular polarity. The abnormal maintenance of polarity resulted in a defect in the motile behavior of S13A cells at the peak and in the decreasing phase of temporal waves. The perimeters of JH10 cells (Fig. 8A) and WT-res cells (data not shown) at the peak and in the back of simulated temporal waves became on average relatively round and polar, and tended to stack one on top of the other in a time series, indicating little net translocation in any one direction. However, the perimeters of S13A cells (Fig. 8B) remained on average elongate and polar, and generated tracks with a directional component, indicating that S13A cells continued to crawl abnormally at the peak and in the back of simulated temporal waves, albeit at reduced velocity.
Myosin distribution in the peak and back of temporal waves Myosin II localizes to the cortex of normal elongate cells
translocating in the front of a wave and is more generally distributed in less polar cells not actively suppressing lateral pseudopod formation (Wessels et al., 2000b). In the front of simulated temporal waves of cAMP, myosin II localized to the cortex of the posterior two-thirds of rapidly translocating, elongate JH10 cells (Fig. 7J) and S13A cells (Fig. 7J′). In the back of simulated temporal waves of cAMP, although myosin II was more evenly distributed throughout the cytoplasm of apolar, nontranslocating JH10 cells (Fig. 7K), it was still localized in the cortex of the posterior two-thirds of the abnormally elongate S13A cells (Fig. 7K′). Similar results were attained when the same analysis was performed on nine additional JH10 cells and nine additional S13A cells in each phase of the wave.
Mutant cells respond abnormally to the rapid addition of 10–6 M cAMP In the increasing phase of a natural wave, a cell experiences an increase in the concentration of cAMP from less than 10–8 M at the trough to 10–6 M at the peak over a period of several minutes (Tomchik and Deverotes, 1981). At the peak of a wave, a normal cell loses polarity and stops translocating (Wessels et al., 1992). One approach that has been commonly used to assess the cellular response to the peak of the wave is to add cAMP (10–6M) rapidly to cells in buffer (e.g. Ross and Newell, 1981; Hall et al., 1988; Wessels et al., 1989). When cAMP is added rapidly to cells crawling in a perfusion chamber, so that the concentration increases from 0 to 10–6 M in less then 8 seconds, the cells stop translocating, round-up and lose cellular polarity within 20 seconds from the time cAMP first enters the chamber (Wessels et al., 1989). These behavioral changes are similar to those of cells responding to the peaks of simulated temporal waves of cAMP and to the peaks of natural waves (Wessels et al., 1992). JH10 cells responded to the rapid increase in cAMP in a manner similar to that described for other wild-type strains of Dictyostelium (Wessels et al., 1989; Wessels and Soll, 1990; Cox et al., 1992; Escalante et al.,
Fig. 8.S13A cells abnormally fail to lose polarity and abnormally continue to translocate, albeit at diminished velocity, at the peak and in the back of a simulated temporal wave of cAMP and after the rapid addition of 10–6 M cAMP. (A,B) Perimeter tracks of representative JH10 and S13A cells, respectively, at the peak and in the back of the second and third waves in a series of four simulated temporal waves generated in the absence of a spatial gradient. (C,D) Perimeter tracks of representative JH10 and S13A cells, respectively, after the rapid addition of 10–6
M cAMP.
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1997). Prior to the addition of 10–6 M cAMP, the centroid tracks of JH10 cells reflected relatively persistent and rapid translocation (Fig. 9A). Within 10 seconds after the addition of cAMP to the chamber, centroids clustered, reflecting the cessation of cellular translocation (Fig. 9A). After the addition of cAMP, perimeters stacked one on top of the other, again reflecting the cessation of cellular translocation (Fig. 8C). Perimeters also became rounder, reflecting the loss of cellular polarity (Fig. 8C).
Prior to the addition of 10–6 M cAMP, the centroid tracks of the S13A cells also reflected persistent translocation (Fig. 9B). However, after the addition of 10–6 M cAMP the centroids did not cluster tightly like those of JH10 cells. Rather, they reflected continued translocation, albeit at reduced velocity. This interpretation was reinforced in perimeter tracks. After the rapid addition of cAMP, S13A cells retained their elongate morphologies and translocated in a persistent manner (Fig. 8D), similar to S13A cells responding to the peak and back of simulated temporal waves (Fig. 8B). Therefore, S13A cells abnormally retained an elongate, polar morphology and continued to translocate (albeit at reduced velocity), after the rapid addition of 10–6 M cAMP, the same abnormalities exhibited at the peak of simulated temporal waves of cAMP.
Mutant cells exhibit defects at the peak and in the back of self-generated natural waves of cAMP Based on the behavioral phenotypes of S13A cells in buffer, in a spatial gradient of cAMP and in simulated temporal waves of cAMP, one would expect S13A cells to orient correctly at the onset of each natural wave (phase A, Fig. 1A) and translocate in a persistent fashion towards the aggregation center in the front of the wave (phase B, Fig. 1A), but abnormally remain elongate (i.e. not undergo cellular depolarization) and abnormally continue to translocate at the peak and in the back of the wave (phases C,D, Fig. 1A). To assess the behavior of mutant cells in natural waves, we employed a submerged culture protocol (Escalante et al., 1997) that allowed comparison of the behavior of individual S13A, JH10 and WT-res cells in self-generated natural waves of cAMP with similar average periodicity (5 minutes for S13A
cells, 6 minutes for JH10 cells and 5 minutes for WT-res cells). Time plots of velocity for JH10 cells (Fig. 10A), WT-res cells (data not shown) and S13A cells (Fig. 10B) contained peaks and troughs at relatively constant intervals reflecting increased velocity in the front of the waves (phase B) and decreased velocity at the peak (phase C) and in the back (phase D) of waves.
Centroid tracks of both JH10 and S13A cells pointed in the general direction of their respective aggregation centers during each rapid translocation segment (phase B) (Fig. 10C and 10D, respectively), demonstrating that S13A cells assessed the correct direction of the spatial gradient of cAMP at the onset of each self-generated natural wave (phase A). However, neighboring S13A centroid tracks did not appear to exhibit on average the overall accuracy of JH10 cells (i.e. maintain the same level of directionality towards the deduced aggregation center; data not shown). In addition, S13A cells abnormally retained polarity and continued to translocate, albeit at reduced velocity, in the deduced peak and back of each self-generated wave, just as they did in the back of simulated temporal waves. The tracks of JH10 cells (Fig. 10C) included segments in which centroids were separated and aligned in the direction of the aggregation center (arrow), representing behavior in the front of each wave (phase B), interspersed with segments in which the centroids were highly clustered, reflecting little net translocation in any one direction at the peak (phase C) and in the back (phase D) of each natural wave (Fig. 10B). Outlined images of a representative JH10 cell through a wave revealed an elongate morphology during the translocation segment in the deduced front of the wave (phase B), and the loss of polarity during centroid clustering at the deduced peak and in the deduced back of the wave (phase C and D; Fig. 10E). The centroid tracks of S13A cells (Fig. 10E) also included segments in which the centroids were separated and aligned in the general direction of the aggregation center, representing behavior in front of each wave (phase B), interspersed with contracted segments in which the distances between centroids were reduced. The contracted segments still exhibited alignment, reflecting slower but still persistent translocation at the peak and in the back of the wave, the same abnormal behavior observed at the peak and in the back of simulated temporal waves of cAMP. Outlined images of a representative
A. JH10 B. S13A
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Fig. 9.Centroid tracks of representative JH10 cells (A) and S13A cells (B) prior to (–10 to 0 minutes) and after (0 to +10 minutes) the rapid addition of 10–6 M cAMP. Time interval between centroids is 10 seconds. Similar results were obtained for 17 additional JH10 and S13A cells analyzed in the same fashion. Results similar to those of JH10 were obtained for WT-res cells analyzed in the same fashion and results similar to those for S13A-1 cells were obtained for S13A-2 and S13A-3 cells analyzed in the same fashion.
S13A cell through a wave revealed the abnormal maintenance of an elongate, polar morphology at the peak and in the back of the wave (Fig. 10F), the same abnormality exhibited at the peak and in the back of a simulated temporal wave (Fig. 8B).
S13A cells respond abnormally to wild-type waves If S13A cells respond abnormally to the peaks and backs of self generated cAMP waves in aggregation territories, they should also respond abnormally to the peaks and backs of cAMP waves generated by wild-type cells. To test this prediction, S13A cells were stained with the vital dye DiI, mixed at a 1:9 ratio with unlabeled JH10 cells, and analyzed by transmitted light and fluorescence microscopy. The results (Fig. 11) were similar to those collected for the two cell types in self generated waves. In the centroid tracks of the dominant cell type JH10, expanded, persistent segments (phase B) were interspersed with highly clustered segments (phase C and D). The net direction of the representative JH10 track in Fig. 11, was towards the aggregation center. The tracks of nine additional JH10 cells analyzed in the same manner exhibited the same general characteristics. In the centroid track of a neighboring S13A cell, expanded persistent segments (phase B) were interspersed with less extensive, but still persistent segments (phases C and D). As in simulated temporal waves and self-generated natural waves, S13A cells continued to translocate at the peak and in the back of natural waves generated by JH10 cells. In addition, although the track of the representative S13A cell was in the general direction of the aggregation center, its accuracy was not as great as that of the
neighboring JH10 cells (Fig. 11). The tracks of nine additional S13A cells analyzed in the same manner exhibited the same general characteristics as the representative S13A cell in Fig. 11, and were, on average, also less on track in phase B than neighboring JH10 cells (data not shown).
Streaming is defective during S13A aggregation In the previous sections, we demonstrated behavioral defects associated with single cell chemotaxis, which occurs early in the aggregation process. However, late in the aggregation process, cells coalesce into multicellular streams, in which they move, still in a pulsatile fashion, into the final aggregate (Reitdorf et al., 1997). To test whether streaming was normal in late aggregating S13A cell populations, fields of cells were video-recorded at low magnification. Whereas JH10 cells formed normal contiguous streams late in aggregation that grew thicker as aggregation progressed, S13A cells formed streams that fragmented along their lengths (Fig. 12).
Discussion Disruption of the gene mlcR, which encodes the myosin regulatory light chain (RLC) in Dictyostelium discoideum, resulted in defects in cytokinesis, morphogenesis and motility (Chen et al., 1994). These defects were similar to those obtained with the original disruption of the myosin heavy chain (DeLozanne and Spudich, 1987; Knecht and Loomis, 1987; Wessels et al., 1988). Although it is not clear whether the defects exhibited by the RLC deletion mutant were due to the
Fig. 10.S13A cells remain abnormally elongate and continue to translocate, albeit at reduced velocity, at the deduced peak and in the deduced back of a self-generated natural wave of cAMP. (A,B) Velocity plots of a representative JH10 cell and a representative S13A-1 cell in respective homogeneous aggregation territories responding to three natural sequential waves of cAMP. The phases of the wave (A+B, C+D) are deduced from the velocity plots described previously (Wessels et al., 1992). (C,D) Centroid tracks of the representative JH10 cell and representative S13A cell through the three successive natural waves (1,2,3) in which the deduced peak plus back portions (phases C plus D) are boxed. Arrows point in the direction of the interpreted aggregation centers. (E,F) Amplified centroid tracks through one wave and associated cell morphologies. Similar results were obtained for nine additional S13A-1 cells and ten S13A-2 cells analyzed in a similar fashion.
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mislocalization of myosin II or to an alteration in motor function, a recent analysis of mutants harboring RLCs with point mutations suggested that RLC played a direct role in the motor properties of myosin (Chaudoir et al., 1999). The DictyosteliumRLC is phosphorylated in response to the rapid addition of chemoattractant (Kuczmarski and Spudich, 1980; Berlot et al., 1985; Berlot et al., 1987; Griffith et al., 1987), suggesting that phosphorylation plays a role in chemotaxis. However, the gross defects of the mlcR– mutant were reversed by the reintroduction of an RLC lacking the single phosphorylation site at serine 13 (Ostrow et al., 1994), suggesting that RLC phosphorylation was not required for myosin II function. In vertebrates, bending of the smooth muscle and nonmuscle myosin tail is regulated by RLC phosphorylation through a Ca2+-calmodulin-dependent myosin light chain kinase, which stimulates assembly and actin- activated ATPase (Suzuki et al., 1978; Somlyo and Somlyo, 1981; Kamm and Stull, 1985; Ikebe et al., 1987; Trybus and Lowery, 1987). In Dictyostelium, phosphorylation of RLC regulates only enzymatic activity (Griffith et al., 1987). The observations that RLC phosphorylation is tightly coupled to cAMP receptor occupancy, and that it plays a role in enzymatic activity led us to hypothesize that it must play a role in the complex set of responses of Dictyosteliumamoebae to the natural chemotactic wave (Wessels et al., 1992; Wessels et al., 2000a; Wessels et al., 2000b).
Dissecting the complex behavior of Dictyostelium amoebae in natural chemotactic waves The chemotactic responsiveness of Dictyosteliumamoebae has
been assessed by a variety of in vitro protocols, including the rapid addition of 10–6 M cAMP, slow release of cAMP from a micropipette and the genesis of a spatial gradient of cAMP in a gradient chamber. However, the actual chemotactic signal a cell experiences in nature is quite different. In a natural aggregation territory, individual cells respond to nondissipating, symmetrical waves of cAMP relayed from the aggregation center outwardly through the cell population (Tomchik and Devreotes, 1981). A cell responds to each phase of a natural wave in a relatively different fashion (Fig. 1A) (Varnum et al., 1985; Varnum-Finney et al., 1987a; Wessels et al., 1992; Wessels et al., 2000b). In the front of each natural wave, cells experience an increasing spatial gradient of cAMP (increasing in the direction of the aggregation center) and an increasing temporal gradient of cAMP (concentration increasing with time). It has been proposed (Wessels et al., 1992) that cells use the direction of the spatial gradient at the onset of the front of the wave to polarize in the direction of the aggregation center. Once that direction is set, cells respond to the associated increasing temporal gradient of cAMP in the front of the wave by suppressing lateral pseudopod formation, which facilitates rapid and directional movement in a blind fashion in the direction of the aggregation center (Wessels et al., 1992). At the peak of each natural wave, cells experience a cAMP concentration that has been demonstrated to cause a loss of cellular polarity and a dramatic decrease in instantaneous velocity (Varnum and Soll, 1984; Wessels et al., 1989; Wessels et al., 1992). In the back of the wave, cells experience a decreasing spatial gradient of cAMP (decreasing in the direction of the aggregation center) and a decreasing temporal gradient of cAMP (concentration decreasing with time). The decreasing temporal gradient suppresses cellular repolarization, resulting in the formation of pseudopods in random directions and no net translocation in any direction. This complex sequence of behavioral responses to the different spatial, temporal and concentration components of the four phases of the natural wave (Fig. 1A) confines directed cellular translocation towards the aggregation center to the front of the wave. The variety of experimental protocols employed in the present study (Fig. 1B) have allowed us to test which, if any, of the phase specific responses involve RLC phosphorylation.
S13A cells are faster, even in the absence of a chemotactic signal We have found that, despite their inability to phosphorylate RLC, S13A cells translocate faster than wild-type cells and form fewer lateral pseudopods in the absence of a chemotactic signal. Although the decrease in pseudopod formation must contribute to the observed increase in velocity, it does not represent the entire explanation. The increased separation of centroids and perimeters in plotted tracks of S13A cells in buffer, in spatial gradients of cAMP and in the front of simulated temporal and natural waves of cAMP suggests that the basic speed of individual mutant cells is greater than that of wild-type cells, independent of lateral pseudopod formation and turning. These results demonstrate that the serine phosphorylation site is necessary for both the normal frequency of turning and the normal velocity of a translocating cell in the absence of a cAMP signal, and that phosphorylation/
Dire ct
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Fig. 11.S13A cells respond abnormally to the deduced peak and back of natural waves generated by JH10 cells in mixed cultures. Tracks are presented of a representative labeled S13A-1 cell and a representative unlabeled neighboring JH10 cell in an aggregation territory that includes S13A-1 and JH10 cells in a ratio of 1:9. JH10 cells exhibited tracks similar to those in homogeneous JH10 cell populations, suggesting that the waves relayed in the mixed population conformed to that of the majority JH10 cell type. Note that the labeled S13A-1 cell continued to translocate in a persistent fashion, albeit at reduced velocity, at the peak and in the back of deduced waves. Note also how the S13A cell veers off track. The decrease in tracking efficiency was observed in a majority of labeled S13A-1 cells in JH10 aggregation territories. Reverse labeling experiments were performed that demonstrated that labeling did not contribute to the observed effects.
dephosphorylation of RLC plays a role in the basic motile behavior of a cell in the absence of extracellular cAMP.
S13A cells migrate faster and with higher chemotactic efficiency in spatial gradients of cAMP S13A cells also exhibited a higher average chemotactic index than either of the two control cell types. This observation was at first counter intuitive, since one would have expected most mutations in cytoskeletal events downstream of cAMP receptor occupancy to decrease the efficiency of chemotaxis. However, it may not have been completely surprising given the inverse relationship demonstrated between the efficiency of chemotaxis and the frequency of lateral pseudopod formation. Varnum-Finney et al. demonstrated that as the chemotactic index increases, the rate of lateral pseudopod formation decreases (Varnum-Finney et al., 1987b). S13A cells already exhibit depressed rates of lateral pseudopod formation in their basic motile behavior, which appear to enhance chemotactic efficiency in a spatial gradient. Therefore, if S13A cells can still assess the direction of a spatial gradient and adjust direction by relying more heavily on biased anterior pseudopod expansion than new lateral pseudopod formation, they may chemotax more efficiently. Why, then, does Dictyosteliumgo to the trouble of phosphorylating the RLC? One possible answer is that cells in vivo must assess not only the spatial characteristics, but also the temporal dynamics of a natural cAMP wave in order to chemotax properly, and that the phosphorylation/dephosphorylation of RLC is intricately involved in this complex process.
S13A cells respond abnormally to the peak and back of simulated temporal and natural waves of cAMP To test whether S13A cells were defective in responding to the temporal characteristics of natural waves, they were subjected to a series of increasing and decreasing temporal gradients that simulated the temporal dynamics of sequential waves in the absence of spatial gradients (Varnum-Finney et al., 1987a; Wessels et al., 1992). The velocity responses of mutant cells were generally normal. Cells moved at peak velocities in the front of waves, and at trough velocities at the peak and in the back of waves. However, mutant cells failed to round up at the peak of the wave and remained abnormally polarized
(elongate) in the back of waves. Mutant cells continued to move in a directed fashion at the peak and in the back of simulated temporal waves, although at greatly reduced average velocity. These results demonstrate that phosphorylation of RLC is necessary for the morphological response to the high concentration of cAMP at the peak of the natural wave, but is not essential for the general reduction in velocity. Mutant cells also abnormally retained polarity in the back of the wave even though velocity remained generally suppressed. To confirm that the absence of depolarization in response to the peak of the wave represented a failure of mutant cells to respond to the peak concentration of cAMP (Varnum and Soll, 1984), we tested the response of mutant cells to the rapid addition of 10–6
M cAMP (Wessels et al., 1989). Although the rapid increase in cAMP caused a dramatic reduction in velocity, it did not elicit a loss of cellular polarity, as it did in wild-type cells. This defect had an impact on the motile behavior of mutant cells at the peak and in the back of both simulated temporal waves and natural waves of cAMP. While there is very little net translocation by wild-type cells at the peak and in the back of simulated temporal cAMP waves (Varnum et al., 1985; Varnum-Finney et al., 1987a) and deduced natural waves (Wessels et al., 1992), abnormally elongate mutant cells continued to translocate in a directed fashion, albeit at reduced velocity.
S13A cells respond less efficiently to natural cAMP waves If normal depolarization at the peak of a natural wave and the normal maintenance of the depolarized state in the back of the natural wave are necessary components of chemotaxis, then S13A cells must be less efficient in natural chemotaxis. Our results demonstrate that this is indeed the case. S13A cells exhibited the same defects at the peak and in the back of self- generated natural waves as those exhibited at the peak and in the back of simulated temporal waves. In addition, the tracks of S13A cells, although generally directed at a common aggregation center, were on average less on course than tracks of JH10 cells responding to self-generated natural waves. When a minority of labeled S13A cells were mixed with unlabeled JH10 cells, their tracks were also generally directed towards the aggregation center, but again were on average less on course than the tracks of neighboring JH10 cells. The
Fig. 12.Streams of S13A cells late in aggregation fragment. Homogeneous populations of JH10 and S13A-1 cells were video-recorded at low magnification late in aggregation during stream formation. Stream formation and fragmentation are obvious in the S13A-1 cultures. In repeat experiments, S13A-1 and S13A-2 streams formed and fragmented, as in the representative panels in B. Zero minutes represents the time at which video-recording was initiated.
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decrease in efficiency appeared to be in the decision on direction in deduced phase A of each natural wave. This result suggests that cells may be more efficient in making the correct directional decision at the onset of the front of a natural wave if they have undergone depolarization at the peak and in the back of the preceding wave, hence the importance of RLC phosphorylation
Late in aggregation the streams of S13A cells abnormally fragmented, suggesting that depolarization in response to the peak and back of waves also plays a role in maintaining the integrity of streams. This is not a surprising result given the fact that changes in light defraction, which reflect cell shape changes, move outwardly through streams in association with naturally moving cAMP waves late in aggregation (Reitdorf et al., 1997) (H.Z., D.W. and D.R.S., unpublished).
Mechanism In buffer, S13A cells migrate with increased persistence as a result of a decrease in the frequency of lateral pseudopod formation. Together, localization of myosin II in crawling cells and the behavioral phenotype of myosin heavy chain deletion mutants (Wessels et al., 1988) suggest that myosin is involved in the suppression of lateral pseudopod formation in the posterior two thirds of a polarized cell. The results presented here suggest that RLC phosphorylation is involved in overcoming this suppression. In Dictyostelium, RLC phosphorylation increases myosin motor activity by increasing the rate of actin-activated ATP hydrolysis. This increase in motor function is manifested in in vitro motility assays as increased myosin movement of actin. In vivo, localized RLC phosphorylation in a cortical region may increase the relative mobility of myosin, producing a site where the cortex is more conducive to the nucleation of actin assembly, resulting in pseudopod extension. The S13A mutant, unable to increase myosin motor function, would be deficient in the production of these sites, resulting in an overall decrease in lateral pseudopod formation, as has been observed.
There is growing evidence that myosin activity facilitates pseudopod extension. The movement of actin and myosin in a localized region in the lateral cortex would produce a local decrease in rigidity, effectively generating an opening for actin polymerization. Consistent with this model, myosin heavy chain null mutants extend pseudopods in all directions (i.e. do not suppress pseudopod formation in the posterior two thirds of the cell body) (Wessels et al., 1988); myosin heavy chain kinase, which promotes the disassembly of myosin filaments, localizes in pseudopodial regions (Steimle et al., 2001); and PAKa, which promotes myosin assembly, localizes to the posterior of the cell (Chung and Firtel, 1999).
As described here, wild-type cells undergo a loss in polarity at the peak of simulated temporal and natural waves. The rapid addition of 10–6 M cAMP to wild-type cells also causes a rapid loss in polarity (Wessels et al., 1988) and may correlate temporally with RLC phosphorylation (Berlot et al., 1985). S13A cells fail to exhibit this rapid loss of polarity at the peak of simulated temporal and natural waves, and after the rapid addition of 10–6 M cAMP, demonstrating that RLC phosphorylation is necessary for depolarization. We suggest that, in response to the increasing temporal gradient of cAMP in the front of a natural wave, there is an increased association
of myosin with the cortex. However, as the concentration of cAMP approaches 10–6 M, there is an increase in RLC phosphorylation that facilitates the general relocalization of myosin and the loss of polarity. In the S13A mutant, relocalization does not occur (i.e. myosin II remains localized in the cortex of the posterior two-thirds of the elongate cell), providing an explanation for the failure of S13A mutants to depolarize at the peak of a cAMP wave.
The identification of independent pathways emanating from different phases of the natural wave We recently demonstrated through the behavioral characterization of a mutant of the internal phosphodiesterase RegA that a pathway emanating specifically from the front of the wave is responsible for the suppression of lateral pseudopods. The regA– mutant cannot suppress lateral pseudopods in response to the increasing temporal wave in the front of the wave and, therefore, cannot chemotax (Wessels et al., 2000b). By contrast, regA– cells respond normally to the peak and back of the wave (Wessels et al., 2000b). Here, we have demonstrated that, whereas the RLC phosphorylation mutant S13A responds normally to the front of the wave, it does not respond normally to the peak and back of the wave. These results lead to a model in which independent regulatory pathways emanating from different phases of the natural wave effect very different behavioral responses in the complex sequence of behavioral changes accompanying natural chemotaxis.
The authors are indebted to J. Swails for help in assembling the manuscript. The research was supported by National Institutes of Health grants HD-18577 (D.R.S.) and GM39264 (R.L.C.). The authors acknowledge use of the W. M. Keck Dynamic Image Analysis Facility at the University of Iowa, funded by the W.M. Keck Foundation.
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