Journal of Cell Science
Myosin heavy chain kinases play essential roles in
Ca2+, but not cAMP, chemotaxis and the natural
aggregation of Dictyostelium discoideum
Deborah Wessels1, Daniel F. Lusche1, Paul A. Steimle2, Amanda Scherer1, Spencer Kuhl1, Kristen Wood1,
Brett Hanson1, Thomas T. Egelhoff3and David R. Soll1,*
1Developmental Studies Hybridoma Bank, Department of Biology, University of Iowa, Iowa City, IA 52242, USA
2Department of Biology, University of North Carolina at Greensboro, NC 27412, USA
3Department of Biology, The Lerner Research Institute, Cleveland Clinic Foundation, Cleveland, OH 44195, USA
*Author for correspondence (email@example.com)
Accepted 1 July 2012
Journal of Cell Science 125, 4934–4944
? 2012. Published by The Company of Biologists Ltd
Behavioral analyses of the deletion mutants of the four known myosin II heavy chain (Mhc) kinases of Dictyostelium discoideum
revealed that all play a minor role in the efficiency of basic cell motility, but none play a role in chemotaxis in a spatial gradient of
cAMP generated in vitro. However, the two kinases MhckA and MhckC were essential for chemotaxis in a spatial gradient of Ca2+,
shear-induced directed movement, and reorientation in the front of waves of cAMP during natural aggregation. The phenotypes of the
mutants mhckA2and mhckC2were highly similar to that of the Ca2+channel/receptor mutant iplA2and the myosin II phosphorylation
mutant 3XALA, which produces constitutively unphosphorylated myosin II. These results demonstrate that IplA, MhckA and MhckC
play a selective role in chemotaxis in a spatial gradient of Ca2+, but not cAMP, and suggest that Ca2+chemotaxis plays a role in the
orientation of cells in the front of cAMP waves during natural aggregation.
Key words: Calcium chemotaxis, Microfluidic chamber, Calcium receptor, Myosin II
Phosphorylation and dephosphorylation of the heavy chain plays a
major role in the polymerization, localization and function of
nonmuscle myosin II in both lower and higher eukaryotes
(Middelbeek et al., 2010; Bosgraaf and van Haastert, 2006; Clark
et al., 2008). In the lower eukaryote Dictyostelium discoideum,
phosphorylation of the myosin heavy chain plays a role in a variety
of basic cell behaviors, including cell polarity, the extension of the
anterior pseudopod, repression of lateral pseudopods, uropod
formation and the dynamics of cell migration (Stites et al., 1998;
De La Roche et al., 2002; Heid et al., 2004). Phosphorylation,
therefore, impacts the directed translocation of a D. discoideum
amoeba up a spatial gradient of the chemoattractant cAMP during
chemotaxis (Yumura and Uyeda, 1997; Heid et al., 2004). In human
cells there are several myosin II isoforms (Vicente-Manzanares
et al., 2009; Conti et al., 2008; Berg et al., 2001). Phosphorylation
and dephosphorylation of the myosin IIA, which regulates the
polymerized state of this isoform, has been shown to play a role in
migration, lamellipod formation and the stabilization of stress fibers
(Dulyaninova et al., 2007; Redowicz, 2001). Therefore, the myosin
IIheavychainkinasesand phosphatasesmustplayrolesin basiccell
motility and the efficiency of chemotaxis. Here, we have explored
the roles of the four identified myosin II heavy chain (Mhc) kinases
of D. discoideum by analyzing the behavior of null mutants during
translocation in the absence of chemoattractant, chemotaxis in a
spatial gradient of either of the two chemoattractants, cAMP or
Ca2+, chemokinesis intemporal gradientsof cAMP, and chemotaxis
in naturally aggregating cell populations.
During natural aggregation, D. discoideum amoebae respond
chemotactically and chemokinetically to the spatial and temporal
dynamics, respectively, of waves of the chemoattractant cAMP.
These waves are relayed outwardly through an aggregating cell
population (Tomchik and Devreotes 1981; Soll et al., 2002).
During the developmental program leading to aggregation, cells
not only acquire the receptor and signal transduction pathways
for cAMP chemotaxis, but they also acquire the capacity to
undergo chemotaxis in a spatial gradient of Ca2+(Scherer et al.,
2010). The discovery of Ca2+chemotaxis (Scherer et al., 2010;
Soll et al., 2011) and its selective loss in the Ca2+channel/
receptor mutant iplA2(Lusche et al., 2012), has led to the
hypothesis that transient Ca2+gradients formed between cells at
the onset of each natural cAMP wave may augment orientation in
the direction of the aggregation center.
When D. discoideum amoebae polarize and translocate on a
substratum, myosin II polymerizes in the cortex of the posterior
cell body and uropod (Yumura et al., 1984; Yumura and Fukui,
1985; Soll et al., 2009). The role of the phosphorylation–
dephosphorylation cycle of the myosin heavy chain, MhcA, was
revealed in the behavioral analyses of two mutants. In the mutant
3XASP, the three threonine phosphorylation sites were substituted
withaspartic acid to mimic the constitutivelyphosphorylatedstate,
and in the mutant 3XALA, the three sites were substituted with
alanine to mimic the constitutively unphosphorylated state
(Egelhoff et al., 1993; Lu ¨ck-Vielmetter et al., 1990). In 3XASP
cells, myosin II was primarily disassembled, and in 3XALA cells,
it was overassembled (Egelhoff et al., 1993). 3XASP cells
Journal of Cell Science
exhibited decreases in velocity, polarity, the repression of lateral
pseudopod formation and directional persistence (Heid et al.,
2004). They did not possess a tapered uropod and formed lateral
pseudopods on average at twice the rate of wild-type cells (Heid
et al., 2005). 3XASP cells, however, oriented normally in a spatial
motility (Heid et al., 2004). 3XALA cells, however, translocated
relatively normally, except for frequent bifurcations of the anterior
pseudopod (Yumura and Uyeda, 1997; Stites et al., 1998),
suggesting an increase in cortical tension (Egelhoff et al., 1993;
Stites et al., 1998; Laevsky and Knecht, 2003). In contrast to
3XASP cells, 3XALA cells exhibited a 50% reduction in the
efficiency of chemotactic orientation. The defects in the 3XASP
dephosphorylation cycle plays a role both in basic cell motility
and chemotaxis. This cycle is regulated by myosin heavy chain
kinases and phosphatases (Murphy and Egelhoff, 1999; Rai and
Egelhoff, 2011), which one would assume are the targets of signal
transduction pathways that coordinate remodeling of the cortical
cytoskeleton during basic cell motility and chemotaxis.
To investigate the role of the myosin heavy chain kinases, the
behavior of the individual Mhc null mutants mhckA2(Co ˆte and
Bukiejko, 1987), mhckB2(Clancy et al., 1997; Rico and Egelhoff,
2003), mhckC2(Nagasaki et al., 2002) and mhckD2(Yumura
et al., 2005), and the triple mutant, mhckA2mhckB2mhckC2
(Yumura et al., 2005) (hereafter referred to as mhckA2B2C2), has
been analyzed by computer-assisted live-cell reconstruction and
motion analysis systems. We fully expected to find that one or
more of the mutants would exhibit behavioral defects similar to
those of the mutant 3XALA (Heid et al., 2004). The behavior of
each mutant and its parent strain were analyzed during rapid
perfusion with buffer (in the absence of attractants), in a spatial
gradient of cAMP, in temporal gradients of cAMP, in spatial
gradients of Ca2+and in wild-type aggregation territories. We
found that three of the mutants, mhckA2, mhckC2, mhckD2and
the triple mutant, mhckA2B2C2, exhibited defects in basic motile
behavior in the absence of attractant as well as in a spatial gradient
of cAMP. We found, however, that all five mutants oriented
normally in spatial gradients of cAMP generated in vitro and
exhibited normal chemokinetic responses to temporal increases
and decreases in cAMP in temporal waves generated in vitro to
mimic the dynamics of natural waves. However, while mhckB2
and mhckD2cells oriented normally in a spatial gradient of Ca2+,
mhckA2, mhckC2and mhckA2B2C2had completely lost the
capacity to orient in a spatial gradient of Ca2+. They also lost flow-
directed orientation. More importantly, the three mutants lost the
capacity to reorient accurately in the front of each natural cAMP
wave relayed through a wild-type aggregation territory. These
results indicate that none of the Mhcks are each essential for
orientation in a cAMP gradient, but the two Mhcks, MhckA and
MhckC, are essential for Ca2+chemotaxis and flow-directed
orientation, the same phenotype exhibitedby the null mutant of the
putative Ca2+channel/receptor, IplA (Lusche et al., 2012).
Basic motile behavior in the absence of attractrant
Cells of the mutants mhckA2, mhckB2, mhckC2, mhckD2,
mhckA2B2C2and their parent strains JH10 and Ax2, were
allowed to attachat lowdensity to the glass wall ofa Sykes–Moore
chamber and were then perfused with Tricine buffer (TB)
containing 10 mM Ca2+(10 mM Ca2+solution), which has been
persistence and velocity in the absence of attractant (Lusche
et al., 2009; Lusche et al., 2011). Buffer solution was perfused at a
of the cell perimeters were reconstructed by the computer-assisted
program 2D-DIAS (Soll, 1995; Soll and Voss, 1998). The tracks of
the mutants revealed no obvious qualitative differences with those
of the parental strains (Fig. 1A–D). However, quantification
revealed that the parameters reflecting the speed of cellular
translocation (i.e. instantaneous velocity, percentage of cells
§9 mm per second, positive flow) for the individual mutants
mhckA2, mhckB2and mhckD2, were consistently below those of
their parental strains (Fig. 1E). The decreases in instantaneous
velocity were significant (P,0.05). The persistence measurements
ofmhckA2,mhckB2cells and mhckD2cellswerealso belowthose
of their respective parental strains (Fig. 1E). The results for the
triple mutant mhckA2B2C2were consistent with those for
individual mutants mhckA2and mhckB2and, if anything,
revealed some additivity (Fig. 1E). In marked contrast, the same
motility parameters for the mutant mhckC2were indistinguishable
from those of parental cells (Fig. 1E; P.0.1). These results
demonstrate that none of the four Mhcks is essential for cellular
translocation, but three of them (MhckA, MhckB, MhckD) play
roles in fine tuning basic motile behavior and in attaining
to facilitatemaximum polarity,directional
All mutants undergo cAMP chemotaxis
Cells of the mutant and parental strains were analyzed in spatial
gradients of cAMP generated in a chamber consisting of a bridge,
which supports test cells, and two bordering troughs, one filled
with 10 mM Ca2+solution and the other filled with 10 mM Ca2+
solution containing 1026M cAMP. In this chamber, a gradient
forms across the bridge after five minutes and remains steep
enough to induce chemotaxis for 15 additional minutes (Varnum
and Soll, 1981; Varnum and Soll, 1984). Cell behavior was video
Cells of all five mutants moved in a directed fashion up a cAMP
the five mutants varied between +0.40 and +0.55, while those of
parental strains JH10 and Ax2 were +0.57 and +0.60, respectively
(Fig. 2E). The percentage of cells of the five mutant strains with
positive chemotactic indices varied between 79 and 91%, while
those of the parental strains JH10 and Ax2 were 94 and 90%,
respectively (Fig. 2E). These results demonstrate that none of the
Mhcks are individually essential for cAMP chemotaxis. The triple
mutant underwent cAMP chemotaxis, demonstrating that at least
in the case of MhckA, MhckB and MhckC, the results do not
reflect a case of redundancy.
As was the case in the absence of attractant (Fig. 1), the three
motility parameters instantaneous velocity, percentage of cells
with velocities §9 mm per minute and positive flow of the
mutants mhckA2, mhckB2, mhckD2and mhckA2B2C2cells were
lower than that of parental cells (Fig. 2E). The differences were
attractant (Fig. 1), themotility parameters of mutant mhckC2were
similar to those of the parental strain Ax2 (P.0.05; Fig. 2E).
Therefore, the velocity defects of mhckA2, mhckB2, mhckD2and
mhckA2B2C2observed in the absence of chemoattractant
(Fig. 1E) were also observed when cells were undergoing
chemotaxis in a spatial gradient of cAMP (Fig. 2E).
MhckA and MhckC are required for Ca2+chemotaxis in Dictyostelium4935
Journal of Cell Science
All mutants respond normally to temporal waves of cAMP
During natural aggregation, cells undergo changes in motility in
response to the temporal dynamics of each relayed wave of
cAMP. Instantaneous velocity increases in response to the
increasing temporal gradient in the front of each wave, and
decreases in response to the peak and decreasing temporal
gradient in the back of each wave (Varnum et al., 1985; Soll
et al., 2002). These chemokinetic responses can be assessed in
vitro by measuring instantaneous velocity in a series of temporal
waves of cAMP generated with pumps attached to a round
perfusion chamber (Geiger et al., 2003; Wessels et al., 2009).
Under these conditions, spatial gradients of cAMP were not
established. The wave periodicity was seven minutes, the
average periodicity of natural waves in an aggregation
territory (Tomchik and Devreotes, 1981). The response of a
representative parental wild-type cell is shown in Fig. 3A. In
each temporal wave, instantaneous velocity increased in the
increasing phase, and decreased at the peak and in the decreasing
phase (Fig. 3A). All five MhckA mutants (mhckA2, mhckB2,
mhckC2, mhckD2, mhckA2B2C2) responded normally to a
series of four temporal waves (Fig. 3B–F, respectively). In the
front of each of the last three in a series of four temporal waves
there was an increase in instantaneous velocity. Although the
triple mutant exhibited an increase in the front of each wave, it
consistently exhibited aberrant surges in the backs of waves
(Fig. 3F). Ten cells of each parental and mutant strain were
analyzed, with similar results.
mhckA2, mhckC2and mhckA2B2C2cells do not undergo
To test whether the five mutants underwent Ca2+chemotaxis, TB
plus 10 mM Ca2+was pumped along one side of a microfluidic
chamber, and TB lacking Ca2+along the other (Scherer et al.,
2010). A series of chevron micromixers between the two flow
channels of the chamber generated a stable Ca2+gradient that was
perpendicular to the direction of flow (Scherer et al., 2010). We
also demonstrated that wild-type cells were mechanoresponsive
to the shear force caused by the high level of flow (Scherer et al.,
2010), as others have shown (De ´cave ´ et al., 2003; Fache et al.,
2005; Lombardi et al., 2008). Because wild-type cells respond to
both a chemotactic signal and mechanosignal, wild-type cells
move up a Ca2+gradient generated in the microfluidic chamber
with a bias in the direction of flow (i.e. at an angle between the
direction of the Ca2+gradient and the angle of flow) (Scherer
et al., 2010). Parental JH10 and Ax2 cells, in the experiments
reported here, moved up Ca2+gradients with chemotactic indices
percentage positive chemotaxis measures of 75 and 80%,
respectively (Fig. 4A). These values were very close to those
previously reported for wild-type cells (Scherer et al., 2010).
Cells of both parental strains exhibited rightward bias, which was
evident in the rightward direction parameter, calculated as the
degrees directed away from the rightward vector (i.e. the
direction of flow) (Scherer et al., 2010) (Fig. 4A), and in
representative cell tracks (Fig. 4B).
Fig. 1. Deleting the individual MHCK genes can have minor
effects on the basic motile behavior of cells in the absence of
chemoattractant. Cell behavior was assessed in a chamber
perfused with buffer lacking attractant. (A–D) Representative
perimeter tracks of cells of parental strain JH10 and mutants
mhckA2, mhckB2and mhckC2, respectively. (E) Quantification
of the motility paremeters instantaneous velocity (Inst. vel.),
percentage of cells with velocities §9 mm per min (Percent cells
§9 mm/min), positive flow (Posit. Flow) and persistence
(Persist.). Arrows in A–D indicate net direction of a cell.
Journal of Cell Science 125 (20) 4936
Journal of Cell Science
Cells of the mutants mhckB2
measures similar to those of cells of the parental strains
(Fig. 4A). mhckB2and mhckD2cells also exhibited positive
rightward directionality values of +0.14 and +0.13 (Fig. 4A),
which were half that of parental strains, but still far above 0.00, the
measure of unresponsiveness (Scherer et al., 2010). The capacity
of mhckB2cells (Fig. 4D) and mhckD2cells (data not shown) to
undergo chemotaxis and mechanoreception albeit reduced, in a
Ca2+gradient was evident in representative perimeter tracks.
In marked contrast, the chemotactic indices of mhckA2, mhckC2
and mhckA2B2C2cells were 20.0360.28, +0.0060.18 and
20.0260.24, respectively, and the percentage positive chemotaxis
measure in all cases was close to 50% (Fig. 4A). Rightward
directionality measurements were 20.0460.27, +0.0460.26 and
20.0560.29, respectively (Fig. 4A). The differences between the
mutants (mhckA2, mhckC2and mhckA2B2C2) and their parental
strains for the two chemotactic parameters and rightward
directionality were significant (P,0.05). Representative tracks
of mhckA2(Fig. 4C), mhckC2(Fig. 4E) and mhckA2B2C2(data
not shown) cells reflected directional randomness in a Ca2+
gradient. A comparison of fields of cells in Ca2+gradients at low
magnification revealed that although mhckA2cells were mobile,
they remained randomly dispersed throughout the field of analysis
(Fig. 4G) whereas parental cells moved towards the upper right
hand corner of the field (Fig. 4F). These results demonstrate that
each of the two Mhcks, MhckA and MhckC, but not MhckB or
MhckD, is essential for Ca2+chemotaxis and flow-induced
Defects in chemotactic orientation in the front of natural
Since all four individual Mhck mutants underwent normal
chemotaxis in a spatial gradient of cAMP and normal
chemokinetic responses to increasing and decreasing temporal
gradients of cAMP in vitro, one might expect all four to respond
normally to the spatial and temporal dynamics of cAMP waves
relayed in a natural aggregation territory of majority wild-type
cells. This expectation was based on the premise that every
aspect of the complex behavior of cells in a natural wave of
cAMP could be explained by their responses to the increasing
and decreasing spatial gradients of cAMP, and the increasing and
decreasing temporal gradients of cAMP that accompany the front
and back, respectively, of each relayed wave (Soll et al., 2002).
To test this, vitally stained mutant cells and unstained JH10 cells
were mixed at a ratio of 1:9, respectively, and allowed to
aggregate on the surface of a 35 mm plastic Petri dish (Wessels
et al., 2000; Wessels et al., 2007; Lusche et al., 2012). The
outwardly moving, nondissipating waves of cAMP in these
mixed populations are dictated by the majority wild-type cells.
Fig. 2. Deleting the individual MHCK genes does not affect
the efficiency of chemotaxis in a spatial gradient of cAMP.
Chemotaxis was assessed in a gradient chamber designed after
that of Zigmond (Zigmond, 1977; Varnum and Soll, 1984).
(A–D) Representative perimeter tracks of cells of parental strain
JH10 and mutants mhckA2, mhckB2and mhckC2, respectively.
(E) Quantification of motility and chemotaxis parameters. The
motility parameters are defined in the legend in Fig. 1. The
chemotaxis parameters were chemotactic index (Chem. Index,
C.I.) and percentage of cells with a positive chemotactic index
(Percent + C.I.). Large arrows in A–D represent direction of
MhckA and MhckC are required for Ca2+chemotaxis in Dictyostelium4937
Journal of Cell Science
The form and frequency of each natural wave passing over an
individual minority mutant cell was deduced by the transient
increase in instantaneous velocity of neighboring majority wild-
type cells. Majority wild-type cells responded chemotactically
and chemokinetically to the front of each wave (Fig. 5A,B). In
the front of each deduced wave, wild-type cells moved with
increased velocity in the direction of the aggregation center, and
at the peak and in the back of each deduced wave, the velocity of
cells decreased dramatically, with little net movement towards
the aggregation center. These behaviors were evident in velocity
plots (Fig. 5A) and tracks of the cell centroid (Fig. 5B). Minority
mhckA2, mhckB2and mhckC2, mhckD2and mhckA2B2C2
cells all exhibited normal chemokinetic increases and decreases
in instantaneous velocity in the fronts and backs of consecutive
relayed wild-type waves (red dot plots), behaviors similar to
neighboring wild-type cells (black dot plots; Fig. 5C, E and G,
respectively; data for mhckD2and mhckA2B2C2not shown).
The majority of cell centroid tracks (90%) of the mutants
mhckB2and mhckD2(n58 and 10, respectively) also appeared
to be similar to those of neighboring wild-type cells, although
they did display a slightly higher tendency to make sharp turns
(Fig. 5F; data for mhckD2not shown). However, only a minority
(,20%) of the cell centroid tracks of the mutants mhckA2
(Fig. 5D), mhckC2(Fig. 5H) and mhckA2B2C2(data not
shown) exhibited normal trajectories (n59 and 11, respectively).
In the majority of cases, the cells did not appear to reorient at the
onset of each wave, resulting in a high frequency of sharp turns
(Fig. 5D,H). To quantify this abnormality in reorientation, the
angle was measured between the direction of movement in the
front of each of four successive waves and the direction of
the aggregation center, the source of the wave, as diagrammed in
Fig. 5I. In a previous study using this technique (Lusche et al.,
2012), we found the average angle of orientation for wild-type
cells to be 30˚. Here, parental JH10 and Ax2 cells oriented with
average angles of 31.269.2˚and 34.769.5˚, respectively (Fig. 5J).
Mutant mhckB2and mhckD2cells oriented at angles (6 standard
deviation) of 41.2612.6˚and 36.6610.8˚, respectively (Fig. 5J).
These values were found to be statistically indistinguishable from
those of the respective controls (Fig. 5J). Cells of the mutants
mhckA2, mhckC2and mhckA2B2C2, however, exhibited average
angles of 71.0622.1˚, 67.4618.3˚and 68.0617.9˚, respectively
(Fig. 5J). The difference in angle between the three latter mutants
and their respective parental strain washighlysignificant (Fig. 5J).
Together, these results indicate that MhckA and MhckC play
major roles in reorientation in the front of each natural, relayed
wave, but MhckB and MhckD do not.
Myosin II localization in mutants
In parental Ax2 (Fig. 6A,B) and JH10 (data not shown) cells
translocating in buffer, anti-myosin II antibody stained the
cytoplasm diffusely and the posterior cell cortex and uropod
intensely (n525 for both strains), as previously reported (Yumura
et al., 1984; Yumura and Fukui, 1985). A similar pattern was
observed for JH10 cells (data not shown). In a minority (,10%)
of control cells, there was distinct myosin II staining in
pseudopods (data not shown), presumably reflecting pseudopod
retraction, as suggested by Spudich and co-workers, who
monitored GFP-tagged myosin II in live translocating cells
(Moores et al., 1996). Cells of all five mutants exhibited diffuse
myosin II staining in the cytoplasm, and more intense staining in
the uropod and posterior cell cortex (Fig. 6C–J). However, the
pseudopods of a majority (.60%, n520) of mhckA2cells
(Fig. 6C,D) and a majority (.60%, n520) of mhckA2B2C2
cells (Fig. 6I,J) also stained intensely for myosin II. These latter
results suggest that in the absence of MhckA, which localizes in
pseudopods (Steimle et al., 2001a; Steimle et al., 2001b; Steimle
et al., 2002; Liang et al., 2002), myosin II abnormally
unphosphorylated and thus polymerized state.
Ca2+and in vitro MhckA activity
Assessing the direction of a spatial gradient of Ca2+(Scherer
et al., 2010) most likely functions through surface receptors that
activate signal transduction pathways, as is the case for cAMP
chemotaxis in D. discoideum (Swaney et al., 2010; Wang et al.,
2011; Cai and Devreotes, 2011) and chemotaxis to a number of
attractants in animal cells (von Philipsborn and Bastmeyer,
2007), including Ca2+chemotaxis (Aguirre et al., 2010). In
addition, assessing a Ca2+gradient may also be mediated by the
gradient-dependent release of Ca2+if the receptor is a Ca2+
Fig. 3. Deleting the individual MHCK genes does not affect the
chemokinetic response to temporal waves of cAMP generated in vitro that
mimic the temporal dynamics of naturally relayed cAMP waves.
Instantaneous velocity is plotted as a function of time. The cAMP waves are
plotted above the velocity plots. ‘‘f’’ and blue shading denotes the front
(increasing temporal gradient) of each wave.
Journal of Cell Science 125 (20) 4938
Journal of Cell Science
channel (Treves et al., 2010; Prevarskaya et al., 2010). In the
latter case, orientation may be effected by generating a cytosolic
Ca2+gradient that activates MhckA and MhckC in a gradient-
dependent fashion (Yumura et al., 1996). To explore this latter
possibility, we tested whether changes in the concentration of
Ca2+affected MhckA and MhckC activity in vitro. The substrate
in this assay was the MH-1 peptide described elsewhere (Steimle
et al., 2001a). The cytosolic concentration of Ca2+has been
estimated to be 50 nM in the absence of cAMP and 200 nM upon
global stimulation with very high concentrations of cAMP
(Schlatterer et al., 1994; Yumura et al., 1996; Nebl and Fisher,
1997). Increasing the in vitro Ca2+concentration from 0 to
100 mM in the reaction mixture had only a minor effect on
MhckAactivity (Fig. 7A). Increasing
nonphysiological concentration, caused a 30–40% decrease in
activity. Addition of calmodulin, which binds Ca2+and facilitates
its interaction with target proteins, had little effect on MhckA
activity (Fig. 7B,C).Similar
measurements of MhckC activity (data not shown). These
results suggest that the activity of MhckA and MhckC are not
it to500 mM,a
under the regulation of receptor- or channel-dependent changes
in the concentration of cytosolic Ca2+. They do not, however,
contradict the suggestion that Ca2+chemotaxis is receptor
mediated (Scherer et al., 2010).
If MhckA and MhckC regulate Ca2+, but not cAMP, chemotaxis,
through the phosphorylation of Mhc, then the mutant 3XALA,
which produces a constitutively unphosphorylated Mhc, should
share the characteristics of the mutants mhckA2, mhckC2and
mhckA2B2C2. A previous computer-assisted study (Heid et al.,
2004) revealed that cells of the 3XALA mutant underwent
chemotaxis in a spatial gradient of cAMP, but with reduced
efficiency. 3XALA also underwent chemokinesis in the front of
temporal waves of cAMP generated in vitro and in the front of
deduced natural waves of cAMP generated in mixtures of wild-
type and mutant cells (9:1), as described here. We therefore
retested whether 3XALA cells underwent chemotaxis in a spatial
gradient of cAMP in 10 mM Ca2+solution and tested for the first
time whether they underwent chemotaxis in a spatial gradient of
Fig. 4. The mutants mhckA2, mhckB2and mhckA2B2C2have
lost the capacity to undergo chemotaxis in a spatial gradient of
Ca2+. They also lose flow-induced directed motility. Chemotaxis and
flow-induced directed motility were assessed in a microfluidic
chamber (Scherer et al., 2010). (A) Quantification of motility
parameters, flow-induced movement and chemotaxis. Motility and
chemotaxis parameters are defined in the legends to Figs 1 and 2,
respectively. Right. direct., rightward directionality.
(B–E) Representative perimeter tracks of cells of parental strain Ax2
and mutants mhckA2, mhckB2and mhckC2, respectively.
(F,G) Consecutive video frames at 2 minute intervals of Ax2 and
mhckA2cells, respectively. In B–E, thick and thin arrows represent
direction of Ca2+gradient and cell movement, respectively. In F and
G, the white dashed line denotes the middle of the chamber, and time
is presented in minutes (M).
MhckA and MhckC are required for Ca2+chemotaxis in Dictyostelium 4939
Journal of Cell Science
Ca2+. Cells of the mutant strain 3XALA, which was derived from
strain JH10 as previously reported, underwent chemotaxis in a
cAMP gradient with an average chemotactic index of +0.2860.36,
and a percentage positive chemotaxis valueof75%(n548), results
very similar to those previously reported (Heid et al., 2004).
However, 3XALA cells did not undergo chemotaxis in a spatial
gradient of Ca2+. The chemotactic index in the latter case was
+0.0360.32 (n544). As was the case for cells of the mutants
mhckA2, mhckC2and mhckA2B2C2, 3XALA cells did not
exhibit rightward directionality in response to flow (rightward
directionality50.0460.30). And as previously reported for
3XALA cells in a spatial gradient (Stites et al., 1998), myosin II
stained strongly in the cortex and uropod of 3XALA cells in buffer
(data not shown), reflecting overpolymerization (Egelhoff et al.,
The role of myosin II phosphorylation
Behavioral analyses of the phosphorylation mutants of the
myosin II heavy chain (Mhc), 3XASP and 3XALA (Egelhoff
et al., 1993; Egelhoff et al., 1996), indicated that the
phosphorylation–dephosphorylation cycle played a role not
only in the basic motile behavior of a cell, but also in
chemotaxis in a spatial gradient of cAMP and in response to
temporal gradients of cAMP (Heid et al., 2004; Soll et al., 2009;
Stites et al., 1998). The aberrant phenotypes of these mutants
suggested that the dynamic phosphorylation–dephosphorylation
cycle of myosin II was not absolutely essential for these
behaviors, but rather played a role in fine tuning chemotaxis in
a spatial gradient of cAMP. Because the Mhcks had been shown
to phosphorylate myosin II in vitro, we fully expected that one or
more of the Mhck null mutants would partially or fully exhibit
the behavioral defects of the mutant 3XALA.
The role of the Mhck proteins in basic motile behavior
All of the Mhck null mutants except mhckC2exhibited
reductions in velocity. MhckA had been shown to localize to
the pseudopod (Steimle et al., 2001a; Steimle et al., 2001b;
Steimle et al., 2002; Liang et al., 2002), which, when extending,
is relatively devoid of myosin II (Moores et al., 1996). The
Fig. 5. The mutants mhckA2, mhckC2, and
mhckA2B2C2have lost the capacity to reorient in the
front of each natural wave, in the direction of the
aggregation center, the source of the waves. Parental
and mutant cells were mixed at a ratio of 9:1 and allowed
to aggregate. (A,C,E,G) Representative velocity plots of
cells responding to the temporal dynamics of successive
natural waves (drawn above each velocity plot) for parent
strain JH10 and mutants mhckA2, mhckB2and mhckC2,
respectively (mhckD2and mhckA2B2C2not shown). In
A, the data are for two representative JH10 cells in a
homogeneous JH10 population (both black plots), and in
C, E and G, the data are for a JH10 (black) and mutant
(red) cell. (B,D,F,H) Representative centroid tracks at
20 sec. intervals in successive natural waves. In B, the
data are for two representative JH10 cells (black). In D, F
and H, data are for a representative control cell (black)
and representative mutant (red) cells in 9:1 mixtures.
(I) Method for assessing the angle in the front of a wave.
(J) Measured average angle in the front of
Journal of Cell Science 125 (20)4940
Journal of Cell Science
accumulation of myosin II in the pseudopods of mhckA2cells, as
we have shown here, supports the suggestion that MhckA
phosphorylates Mhc in the pseudopod, thus blocking myosin II
polymerization in the dynamic actin gel that drives pseudopodial
expansion (Steimle et al., 2001a; Steimle et al., 2001b; Steimle
et al., 2002). In marked contrast, MhckB localizes to the
cytoplasm (Liang et al., 2002; Underwood et al., 2010), where it
may play a role in maintaining the monomeric myosin II pool
necessary for remodeling the cortical cytoskeleton during cellular
translocation and lateral pseudopod formation. MhckC localizes
to the myosin II-rich posterior cortex and uropod (Nagasaki et al.,
2002; Yumura et al., 2005), where it may play a role in
dismantling the actin–myosin cytoskeleton during remodeling.
Remodeling has been indicated in morphometric studies to occur
at the interface between the cell body and uropod (Soll et al.,
2009). However, this suggested role is not reflected in the
mhckC2mutant, which does not exhibit any measurable defect in
velocity, polarity or uropod maintenance. MhckD localization
has not been reported. These observations suggest that in a wild-
type cell, at least three of the four Mhcks exhibit differences in
localization and, therefore, are unlikely to be functionally
Mhcks and cAMP chemotaxis
All four of the individual Mhck mutants and the triple mutant
mhckA2B2C2, underwent robust chemotaxisinspatial gradients of
cAMP generated in vitro. All five mutants also exhibited relatively
normal chemokinetic responses to increasing and decreasing
temporal gradients of cAMP generated in vitro in temporal waves
that mimicked the temporal dynamics of natural waves. These
Fig. 6. Myosin II distribution in wild-type and mutant
strains during cellular translocation. Cells were mixed
and stained with anti-myosin II antibody. ps, pseudopod;
Fig. 7. Varying Ca2+concentration or adding calmodulin has little effect on MhckA activity in vitro. (A–C) The phosphorylation reaction was performed
with the MH-1 peptide described elsewhere (Steimle et al., 2001a). Similar results were obtained with MhckC.
MhckA and MhckC are required for Ca2+chemotaxis in Dictyostelium4941
Journal of Cell Science
results indicate that no single Mhck, nor a combination of the three
Mhcks, MhckA, MhckB and MhckC, is essential for chemotaxis in
a spatial gradient of cAMP or for the chemokinetic responses to
temporal gradients of cAMP generated in vitro. This leads to the
tentative conclusion that none of these kinases may represent an
exclusive, or essential, target for the signal transduction pathways
regulating cAMP chemotaxis (Yumura et al., 2005). One might
suggest either that other kinases may be responsible for Mhc
phosphorylation and hence serve as targets for these pathways, or
phosphorylation may regulate myosin assembly (Levi et al.,
2002). However, even this explanation seems unlikely given that
3XALA, which contains myosin II heavy chain in a constitutively
unphosphorylated state, undergoes chemotaxis in a spatial gradient
of cAMP and chemokinesis in temporal gradients of cAMP
generated in vitro (Heid et al., 2004).
Mhcks and Ca2+chemotaxis
Although none of the Mhcks were essential for chemotaxis in a
spatial gradient of cAMP, two of them were essential for
chemotaxis in a spatial gradient of Ca2+. The two mutants
mhckA2and mhckC2, and the triple mutant mhckA2B2C2, were
incapable of undergoing chemotaxis in a spatial gradient of Ca2+
(Scherer et al., 2010). These mutants also lost shear-induced
motility in the direction of fluid flow (De ´cave ´ et al., 2002; De ´cave ´
et al., 2003; Fache et al., 2005; Lombardi et al., 2008). The two
and mhckD2, however, underwent Ca2+
chemotaxis and shear-induced directional movement, although
the latter at a reduced level. These results indicate that MhckA and
MhckC are eachessential ina nonredundant fashionfor chemotaxis
in a spatial gradient of Ca2+, but not cAMP. This resultis surprising
for several reasons. First, because the behavioral response appears
so similar during cAMP and Ca2+chemotaxis (Scherer et al., 2010;
Lusche et al., 2012), we hypothesized that while the receptors and
upstream components of the signal transduction machinery may
differ, the downstream pathways and targets regulating the
response, such as the Mhcks, would be common. This
expectation was reinforced by the observation that the capacity to
undergo chemotaxis in both a cAMP and a Ca2+gradient were
attained at approximately the same time in the developmental
program initiatedby starvation
Demonstration here that MhckA and MhckC are each essential
for Ca2+, but not cAMP, chemotaxis suggests that either the
downstream targets ofthesignaltransductionpathwaysdiffer inthe
two chemotactic processes, or that Mhc phosphorylation plays a
role upstream in the regulation of Ca2+, but not cAMP, chemotaxis.
Mhck proteins and natural aggregation
Given that all four of the individual Mhck null mutants underwent
chemotaxis in a spatial gradient of cAMP and chemokinesis in an
increasing temporal gradient of cAMP, we expected all of them to
behave normally in natural wild-type aggregation territories. This
expectation was predicated on the assumption that the behavior of
cellsduring naturalaggregation isdictatedsolelybythe spatial and
temporal dynamics of cAMP in thenatural wave (Soll et al., 2002).
We found instead in mixing experiments (Lusche et al., 2012) that
neither minority mhckA2cells nor minority mhckC2cells undergo
normal chemotaxis in natural waves generated by majority wild-
type cells. They were unable to reorient in the front of each
successive wave. In contrast, the mutants mhckB2and mhckD2
reoriented in the front of natural wild-type waves. Loss of the
capacity to reorient by mhckA2and mhckC2cells correlated with
loss of the capacity to undergo chemotaxis in a spatial gradient of
the hypothesis that Ca2+chemotaxis plays a fundamental role in
natural chemotaxisinanaggregation territory(Scherer et al.,2010;
Lusche et al., 2012).
mhckA2and mhckC2are phenocopies of the mutant iplA2
Like cells of the Mhck null mutants mhckA2and mhckC2, cells of
iplA2, the null mutant of IplA, a putative Ca2+channel/receptor
(Traynor et al., 2000), also undergo chemotaxis in a spatial
gradient of cAMP, but not Ca2+(Lusche et al., 2012). Like
mhckA2and mhckC2cells, iplA2cells have also lost shear-
induced movement in the direction of fluid flow (Lusche et al.,
2012). Finally, like mhckA2and mhckC2cells, iplA2cells have
lost the capacity to reorient in the front of cAMP waves, during
natural aggregation (Lusche et al., 2012). These similarities
suggest that IplA, MhckA and MhckC are components of a signal
transduction pathway for Ca2+, but not cAMP, chemotaxis.
mhckA2, mhckC2and iplA2are phenocopies of 3XALA
mutant 3XALA. If the phenotypic consequences of deleting either
MHCKA or MHCKC is an overabundance of unphosphorylated
Mhc, and, therefore, higher levels of more stable polymerized
myosin II, then the mutant 3XALA should be phenotypically
similar to mhckA2and mhckC2, and that was indeed the case.
Therefore, it is reasonable to conclude that the consequence of
deleting either MHCKA or MHCKC is the overpolymerization of
myosin II, in the cortex and uropod in the former case and in
the pseudopod in the latter case. And based on the assumption
that cAMP chemotaxis employs the same downstream motility
machinery as Ca2+chemotaxis and shear-induced directed
motility, but is intact in the mutants mhckA2, mhckC2, iplA2
and 3XALA, we suggest that myosin phosphorylation plays a role
upstream in the pathway regulating Ca2+chemotaxis and shear-
induced directed motility. If IplA proves to be the common
receptor for these two responses, as has been suggested (Lombardi
et al., 2008; Lusche et al., 2012), then the possibility can be
entertained that the phosphorylation of Mhc by MhckA and
MhckC plays a role in receptor (IplA) function, rather than
downstream in the motile response. The in vitro experiments
reported here reveal that since neither MhckA nor MhckC appears
to be affected in vitro by the changes in Ca2+concentration in the
physiological range, the possibility must be entertained that IplA
may functionas a classical chemotactic receptor at the cell surface,
where it interacts directly or indirectly with MhckA and MhckC,
rather than as a Ca2+channel that regulates these kinases by
affecting the intracellular concentration of Ca2+. However, the
disparate locations of the two Mhcks and the disparate effects
deleting either has on myosin localization are difficult to reconcile
with the common phenotype exhibited by either deletion mutant.
Nonetheless we have identified components in a novel and
selective pathway for Ca2+chemotaxis and shear-induced motility.
Because the components are just emerging, it seems premature to
attempt to design a model for this pathway at the present time.
Materials and Methods
Strain maintenance, growth and development
The mhckA2, mhckB2, mhckC2, and mhckA2B2C2strains were provided by
Thomas Egelhoff (Cleveland Clinic Foundation, Cleveland, OH) and are also
Journal of Cell Science 125 (20)4942
Journal of Cell Science
available in the Dictyostelium stock center (http://dictybase.org/StockCenter/
StockCenter.html. The 3XALA and mhckD2strains were deposited by Thomas
Egelhoff in the Dictyostelium stock center and obtained from there.
To generate the mhckA2strain (Kolman et al., 1996), the pD15 gene
replacement vector containing 59 flanking and 39 regions of the mhckA gene
was electroporated into JH10, a thymidine auxotrophic cell line. The JH10 control
strain was grown in HL-5 medium (http://dictybase.org/techniques/index.html)
supplemented with 100 mg/ml thymidine (Sigma Aldrich, St. Louis, MO) while the
mhckA2cells were maintained in HL-5 alone.
The mhckB2strain was also obtained by insertion of a gene targeting construct
into JH10 (Rico and Egelhoff, 2003). In this case, the construct contained 59 and 39
sequences of the mhckB gene while the central portion of the gene had been
replaced with the blasticidin resistance cassette. In studies presented here, mhckB2
cells were grown in HL-5 medium supplemented with 10 mg/ml Blasticidin S
(Sigma Aldrich, St. Louis, MO).
The mhckC2and mhckD2strains were generated by insertion of the appropriate
blasticidin resistance disruption cassette into the mhckC or mhckD genes of the
Ax2 control strain (Nagasaki et al., 2002). These cells were grown in HL-5
supplemented with 10 mg/ml Blasticidin S.
The mhckA2B2C2strain was obtained using the nonselected co-transformation
protocol described in detail by Betapudi et al. (Betapudi et al., 2004). This
approach was designed to generate a multiple gene knockout strain while
circumventing problems inherent to G418 and hygromycin selection (Betapudi
et al., 2004). Briefly, targeting vectors for the mhckB and mhckC genes containing
blasticidin and hygromycin resistance cassettes, respectively, within the coding
regions were mixed in a 1:6 molar ratio, electroporated into the mhckA2cell line,
and selected for blasticidin resistance. Transformants were screened by PCR to
confirm disruption of all genes (Yumura et al., 2005).
Finally, the 3XALA strain (Egelhoff et al., 1993) in which the three threonines in
the tail region of the myosin II heavy chain were replaced with nonphosphorylatable
alanines, was obtained from Dr Egelhoff (Egelhoff et al., 1993). These cells were
grown in HL-5 supplemented with 10 mg/ml G418 (Sigma Aldrich, St. Louis, MO).
Frozen stocks of the strains were reconstituted every two weeks for experimental
use (Wessels et al., 2007; Lusche et al., 2009). Methods for obtaining aggregation-
competent amoebae for motility assays are described elsewhere in detail (Scherer
et al., 2010). For analyses of basic motility and chemotaxis, cells were harvested
from developmental filters at the onset of aggregation, when velocity and
chemotactic responsiveness were maximal (Varnum et al., 1986).
Analysis of basic motile behavior and behavior in temporal cAMP waves
The method and conditions for analyzing basic motile behavior in the Sykes–
Moore perfusion chamber have been described previously in detail (Soll et al.,
2009; Varnum et al., 1986; Lusche et al., 2009; Wessels et al., 2009). Here, basic
motile behavior was analyzed during perfusion with Tricine buffer (TB) containing
10 mM Ca2+. Sequential temporal waves of cAMP were generated in 40 mM K+
buffer (Varnum et al., 1985; Lusche et al., 2011) in the Sykes–Moore chamber by
pressure-driven pumps as previously described (Geiger et al., 2003; Wessels et al.,
2009). The wave periodicity was 7 min. Digital images were acquired at an
interval of 4 secs for 10 mins as described elsewhere (Lusche et al., 2011).
Analysis of chemotaxis
Cells were distributed across the bridge of the plexiglass gradient chamber
(Zigmond, 1977) according to methods previously described (Varnum and Soll,
1984). Gradients were generated by filling one trough bordering the bridge with
10 mM Ca2+solution in TB lacking cAMP and the other trough with 10 mM Ca2+
solution in TB plus 1026M cAMP. Cells were video recorded for a 10 minute
period following 5–7 minutes of incubation after adding solutions to the troughs.
Cells were inoculated into the custom built microfluidic chamber and a Ca2+
gradient generated using a method identical to the one described elsewhere in
considerable detail (Scherer et al., 2010). The concentration of Ca2+used in the
microfluidic chamber in this study was 10 mM.
Myosin II immunofluorescent staining
Myosin II was stained with Dictyostelium anti-myosin II polyclonal antibody
(Burns et al., 1995), a generous gift from Arturo de Lozanne (University of Texas,
Austin). The methods were described elsewhere in detail (Wessels et al., 2000).
DIAS analysis of behavior
Cell behavior under all conditions was analyzed by 2-D DIAS software and a
description of the parameters is described elsewhere in detail (Soll, 1995; Soll and
with a velocity of §9 mm per minute (% cells §9 mm/min), directional persistence,
chemotacticindexandpercentagepositive chemotaxis werecomputedfromcentroid
positions (Soll, 1995; Soll and Voss 1998; Wessels et al., 2009; Lusche et al., 2012).
‘Instantaneous velocity’ was computed at 4 second intervals between each
consecutive pair of centroids over a ten minute period (Soll, 1995; Soll and Voss,
1998). The parameter ‘% cells §9 mm/min’ was computed as the proportion of cells
in a population with an average instantaneous velocity greater than or equal to 9 mm
per minute. ‘Positive flow’ was determined from overlapping perimeter outlines of
two consecutive cell images. The area in the second of the two images that did not
overlap the first was calculated and expressed as a percentage of the area of the first
image. This was performed at 4 second intervals over a 10 min period for each cell.
‘Directional persistence’ was computed as the net distance between the first and last
centroid of a centroid track divided by the summed distances between consecutive
centroid positions of the track. The ‘chemotactic index’ (C.I.) was computed as the
net distance traveled in the direction of the source of chemoattractant divided by the
total distance traveled. Similarly, rightward directionality (RD) was computed as the
net distance traveled towards the right (direction of flow in the microfluidic
chamber) divided by the total distance traveled. The ‘percentage positive
chemotaxis’ parameter was measured as the proportion of cells in a population
with a chemotactic index greater than 0.00. Values given are the mean (6 standard
deviation) computed for the population.
Mixing experiments for analysis of natural aggregation
cells were labeled with the vital dye DiI (Invitrogen, Carlsbad, CA), mixed with a
majority of unlabeled control cells, and motion analyzed during aggregation
according to methods described in detail elsewhere (Lusche et al., 2012). Briefly,
mhck mutants were labeled by incubation in HL-5 containing 0.05 mM DiI
(Invitrogen) for 24 hrs in the dark. HL-5 was then removed from labeled mutant
cells and excess dye removed by washing in 40 mM K+solution (Lusche et al.,
2011). The appropriate control strain (JH10 or Ax2) and labeled mhck cell
populations were then mixed at a 9:1 ratio, respectively, to a final density of 56106
cells per 2 ml. This suspension was inoculated into a 35 mm Petri dish and the dish
placed on the stage of a Nikon Eclipse TE-2000 microscope connected to a Bio-Rad
Radiance 2100MP laser scanning confocal microscope (Bio-Rad Microscience Ltd,
Hemel Hempstead, UK). After 6 hours, image acquisition was begun using a green
HeNe laser at 543 nm and a 106 objective. Transmitted light images were
continuously collected through a transmitted light detector at 543 nm. Transmitted
and fluorescent images were collected with LaserSharp 2000 software at 20 sec
intervals and converted to QuickTimeTMformat. Labeled mhck2cells and unlabeled
2D-DIAS as described above (Soll, 1995; Soll and Voss 1998; Wessels et al., 2009).
This research was supported by the Developmental Studies
Hybridoma bank at the University of Iowa, a National Resource
initiated by the National Institutes of Health; and by National
Institutesof Health[grant numbers
2R15GM066789 to P.A.S.]. Deposited in PMC for release after 12
Aguirre, A., Gonza ´lez, A., Planell, J. A. and Engel, E. (2010). Extracellular calcium
modulates in vitro bone marrow-derived Flk-1+CD34+progenitor cell chemotaxis
and differentiation through a calcium-sensing receptor. Biochem. Biophys. Res.
Commun. 393, 156-161.
Berg, J. S., Powell, B. C. and Cheney, R. E. (2001). A millennial myosin census. Mol.
Biol. Cell 12, 780-794.
Betapudi, V., Shoebotham, K. and Egelhoff, T. T. (2004). Genertion of double gene
disruptons in Dictyostelium discoideum using a single antibiotic marker selection.
Biotechniques 36, 106-112.
Bosgraaf, L. and van Haastert, P. J. (2006). The regulation of myosin II in
Dictyostelium. Eur. J. Cell Biol. 85, 969-979.
Burns, C. G., Larochelle, D. A., Erickson, H., Reedy, M. and De Lozanne, A. (1995).
Single-headed myosin II acts as a dominant negative mutation in Dictyostelium. Proc.
Natl. Acad. Sci. USA 92, 8244-8248.
Cai, H. and Devreotes, P. N. (2011). Moving in the right direction: how eukaryotic cells
migrate along chemical gradients. Semin. Cell Dev. Biol. 22, 834-841.
Clancy, C. E., Mendoza, M. G., Naismith, T. V., Kolman, M. F. and Egelhoff, T. T.
(1997). Identification of a protein kinase from Dictyostelium with homology to the
novel catalytic domain of myosin heavy chain kinase A. J. Biol. Chem. 272, 11812-
Clark, K., Middelbeek, J., Lasonder, E., Dulyaninova, N. G., Morrice, N. A.,
Ryazanov, A. G., Bresnick, A. R., Figdor, C. G. and van Leeuwen, F. N. (2008).
TRPM7 regulates myosin IIA filament stability and protein localization by heavy
chain phosphorylation. J. Mol. Biol. 378, 790-803.
Conti, M., Kawamoto, S. and Adelsatein, R. (2008). Nonmuscle myosin II. In
Myosins: A Superfamily of Molecular Motors, Vol. 7, Proteins and Cell Regulation
(ed. L.M. Coluccio), pp. 223-264, Dordrecht, The Netherlands: Springer.
Co ˆte, G. P. and Bukiejko, U. (1987). Purification and characterization of a myosin
heavy chain kinase from Dictyostelium discoideum. J. Biol. Chem. 262, 1065-1072.
De La Roche, M. A., Smith, J. L., Betapudi, V., Egelhoff, T. T. and Co ˆte ´, G. P.
(2002). Signaling pathways regulating Dictyostelium myosin II. J. Muscle Res. Cell
Motil. 23, 703-718.
MhckA and MhckC are required for Ca2+chemotaxis in Dictyostelium4943
Journal of Cell Science Download full-text
De ´cave ´, E., Garrivier, D., Bre ´chet, Y., Fourcade, B. and Bruckert, F. (2002). Shear
flow-induced detachment kinetics of Dictyostelium discoideum cells from solid
substrate. Biophys. J. 82, 2383-2395.
De ´cave, E., Rieu, D., Dalous, J., Fache, S., Brechet, Y., Fourcade, B., Satre, M. and
Bruckert, F. (2003). Shear flow-induced motility of Dictyostelium discoideum cells
on solid substrate. J. Cell Sci. 116, 4331-4343.
Dulyaninova, N. G., House, R. P., Betapudi, V. and Bresnick, A. R. (2007). Myosin-
IIA heavy-chain phosphorylation regulates the motility of MDA-MB-231 carcinoma
cells. Mol. Biol. Cell 18, 3144-3155.
Egelhoff, T. T., Lee, R. J. and Spudich, J. A. (1993). Dictyostelium myosin heavy
chain phosphorylation sites regulate myosin filament assembly and localization in
vivo. Cell 75, 363-371.
Egelhoff, T. T., Naismith, T. V. and Brozovich, F. V. (1996). Myosin-based cortical
tension in Dictyostelium resolved into heavy and light chain-regulated components.
J. Muscle Res. Cell Motil. 17, 269-274.
Fache, S., Dalous, J., Engelund, M., Hansen, C., Chamaraux, F., Fourcade, B., Satre,
M., Devreotes, P. and Bruckert, F. (2005). Calcium mobilization stimulates
Dictyostelium discoideum shear-flow-induced cellmotility.J. CellSci.118, 3445-3458.
to waves of chemoattractant, like Dictyostelium. Cell Motil. Cytoskeleton 56, 27-44.
Heid, P. J., Wessels, D., Daniels, K. J., Gibson, D. P., Zhang, H., Voss, E. and Soll,
D. R. (2004). The role of myosin heavy chain phosphorylation in Dictyostelium
motility, chemotaxis and F-actin localization. J. Cell Sci. 117, 4819-4835.
Heid, P. J., Geiger, J., Wessels, D., Voss, E. and Soll, D. R. (2005). Computer-assisted
analysis of filopod formation and the role of myosin II heavy chain phosphorylation in
Dictyostelium. J. Cell Sci. 118, 2225-2237.
Kolman, M. F., Futey, L. M. and Egelhoff, T. T. (1996). Dictyostelium myosin heavy
chain kinase A regulates myosin localization during growth and development. J. Cell
Biol. 132, 101-109.
Laevsky, G. and Knecht, D. A. (2003). Cross-linking of actin filaments by myosin II is
a major contributor to cortical integrity and cell motility in restrictive environments.
J. Cell Sci. 116, 3761-3770.
Levi, S., Polyakov, M. V. and Egelhoff, T. T. (2002). Myosin II dynamics in
Dictyostelium: determinants for filament assembly and translocation to the cell cortex
during chemoattractant responses. Cell Motil. Cytoskeleton 53, 177-188.
Liang, W., Licate, L., Warrick, H., Spudich, J. and Egelhoff, T. (2002). Differential
localization in cells of myosin II heavy chain kinases during cytokinesis and polarized
migration. BMC Cell Biol. 3, 19.
Lombardi, M. L., Knecht, D. A. and Lee, J. (2008). Mechano-chemical signaling
maintains the rapid movement of Dictyostelium cells. Exp. Cell Res. 314, 1850-1859.
Replacement of threonine residues by serine and alanine in a phosphorylatable heavy
chain fragment of Dictyostelium myosin II. FEBS Lett. 269, 239-243.
Lusche, D. F., Wessels, D. and Soll, D. R. (2009). The effects of extracellular calcium
on motility, pseudopod and uropod formation, chemotaxis, and the cortical
localization of myosin II in Dictyostelium discoideum. Cell Motil. Cytoskeleton 66,
Lusche, D. F., Wessels, D., Ryerson, D. E. and Soll, D. R. (2011). Nhe1 is essential for
potassium but not calcium facilitation of cell motility and the monovalent cation
requirement for chemotactic orientation in Dictyostelium discoideum. Eukaryot. Cell
Lusche, D., Wessels, D., Scherer, A., Daniels, K., Kuhl, S. and Soll, D. R. (2012). The
IplA Ca++channel of Dictyostelium discoideum is necessary for chemotaxis mediated
through Ca++, but not through cAMP chemotaxis, and plays a fundamental role in
natural aggregation. J. Cell Sci. 125, 1770-1783.
Middelbeek, J., Clark, K., Venselaar, H., Huynen, M. A. and van Leeuwen, F. N.
(2010). The alpha-kinase family: an exceptional branch on the protein kinase tree.
Cell. Mol. Life Sci. 67, 875-890.
Moores, S. L., Sabry, J. H. and Spudich, J. A. (1996). Myosin dynamics in live
Dictyostelium cells. Proc. Natl. Acad. Sci. USA 93, 443-446.
Murphy, M. B. and Egelhoff, T. T. (1999). Biochemical characterization of a
Dictyostelium myosin II heavy-chain phosphatase that promotes filament assembly.
Eur. J. Biochem. 264, 582-590.
Nagasaki, A., Itoh, G., Yumura, S. and Uyeda, T. Q. (2002). Novel myosin heavy
chain kinase involved in disassembly of myosin II filaments and efficient cleavage in
mitotic Dictyostelium cells. Mol. Biol. Cell 13, 4333-4342.
Nebl, T. and Fisher, P. R. (1997). Intracellular Ca2+signals in Dictyostelium
chemotaxis are mediated exclusively by Ca2+influx. J. Cell Sci. 110, 2845-2853.
Prevarskaya, N., Skryma, R. and Shuba, Y. (2010). Ion channels and the hallmarks of
cancer. Trends Mol. Med. 16, 107-121.
Rai, V. and Egelhoff, T. T. (2011). Role of B regulatory subunits of protein phosphatase
type 2A in myosin II assembly control in Dictyostelium discoideum. Eukaryot. Cell
Redowicz, M. J. (2001). Regulation of nonmuscle myosins by heavy chain
phosphorylation. J. Muscle Res. Cell Motil. 22, 163-173.
Rico, M. and Egelhoff, T. T. (2003). Myosin heavy chain kinase B participates in the
regulation of myosin assembly into the cytoskeleton. J. Cell. Biochem. 88, 521-532.
Scherer, A., Kuhl, S., Wessels, D., Lusche, D. F., Raisley, B. and Soll, D. R. (2010).
Ca2+chemotaxis in Dictyostelium discoideum. J. Cell Sci. 123, 3756-3767.
Schlatterer, C., Gollnick, F., Schmidt, E., Meyer, R. and Knoll, G. (1994). Challenge
with high concentrations of cyclic AMP induces transient changes in the cytosolic
free calcium concentration in Dictyostelium discoideum. J. Cell Sci. 107, 2107-2115.
Soll, D. R. (1995). The use of computers in understanding how animal cells crawl. Int.
Rev. Cytol. 163, 43-104.
Soll, D. and Voss, E. (1998). Two and three dimensional computer systems for
analyzing how cells crawl. In Motion Analysis of Living Cells (ed. D. Soll and
D. Wessels), pp. 25-52. New York, NY: John Wiley Inc.
Soll, D. R., Wessels, D., Heid, P. J. and Zhang, H. (2002). A contextual framework for
characterizing motility and chemotaxis mutants in Dictyostelium discoideum.
J. Muscle Res. Cell Motil. 23, 659-672.
Soll, D. R., Wessels, D., Kuhl, S. and Lusche, D. F. (2009). How a cell crawls and the
role of cortical myosin II. Eukaryot. Cell 8, 1381-1396.
Soll, D., Wessels, D., Lusche, D. F., Kuhl, S., Scherer, A. and Grimm, S. (2011). Role of
extracellular cations incellmotility,polarity,and chemotaxis. Res. Rep. Biol.2011, 89-99.
Steimle, P. A., Naismith, T., Licate, L. and Egelhoff, T. T. (2001a). WD repeat
domains target Dictyostelium myosin heavy chain kinases by binding directly to
myosin filaments. J. Biol. Chem. 276, 6853-6860.
Steimle, P. A., Yumura, S., Co ˆte ´, G. P., Medley, Q. G., Polyakov, M. V., Leppert, B.
and Egelhoff, T. T. (2001b). Recruitment of a myosin heavy chain kinase to actin-
rich protrusions in Dictyostelium. Curr. Biol. 11, 708-713.
Steimle, P. A., Licate, L., Co ˆte ´, G. P. and Egelhoff, T. T. (2002). Lamellipodial
localization of Dictyostelium myosin heavy chain kinase A is mediated via F-actin
binding by the coiled-coil domain. FEBS Lett. 516, 58-62.
Stites, J., Wessels, D., Uhl, A., Egelhoff, T., Shutt, D. and Soll, D. R. (1998).
Phosphorylation of the Dictyostelium myosin II heavy chain is necessary for
maintaining cellular polarity and suppressing turning during chemotaxis. Cell Motil.
Cytoskeleton 39, 31-51.
Swaney, K. F., Huang, C. H. and Devreotes, P. N. (2010). Eukaryotic chemotaxis: a
network of signaling pathways controls motility, directional sensing, and polarity.
Annu. Rev. Biophys. 39, 265-289.
Tomchik, K. J. and Devreotes, P. N. (1981). Adenosine 39,59-monophosphate waves in
Dictyostelium discoideum: a demonstration by isotope dilution-fluorography. Science
Traynor, D., Milne, J. L., Insall, R. H. and Kay, R. R. (2000). Ca(2+)signalling is not
required for chemotaxis in Dictyostelium. EMBO J. 19, 4846-4854.
Treves, S., Vukcevic, M., Griesser, J., Armstrong, C.-F., Zhu, M. X. and Zorzato,
F. (2010). Agonist-activated Ca2+influx occurs at stable plasma membrane and
endoplasmic reticulum junctions. J. Cell Sci. 123, 4170-4181.
Underwood, J., Greene, J. and Steimle, P. A. (2010). Identification of a new
mechanism for targeting myosin II heavy chain phosphorylation by Dictyostelium
myosin heavy chain kinase B. BMC Res. Notes 3, 56.
Varnum, B. and Soll, D. R. (1981). Chemoresponsiveness to cAMP and folic acid
during growth, development, and dedifferentiation in Dictyostelium discoideum.
Differentiation 18, 151-160.
Varnum, B. and Soll, D. R. (1984). Effects of cAMP on single cell motility in
Dictyostelium. J. Cell Biol. 99, 1151-1155.
Varnum, B., Edwards, K. B. and Soll, D. R. (1985). Dictyostelium amebae alter
motility differently in response to increasing versus decreasing temporal gradients of
cAMP. J. Cell Biol. 101, 1-5.
Varnum, B., Edwards, K. B. and Soll, D. R. (1986). The developmental regulation of
single-cell motility in Dictyostelium discoideum. Dev. Biol. 113, 218-227.
Vicente-Manzanares, M., Ma, X., Adelstein, R. S. and Horwitz, A. R. (2009). Non-
muscle myosin II takes centre stage in cell adhesion and migration. Nat. Rev. Mol.
Cell Biol. 10, 778-790.
von Philipsborn, A. and Bastmeyer, M. (2007). Mechanisms of gradient detection: a
comparisonofaxonpathfindingwith eukaryotic cellmigration.Int.Rev.Cytol. 263,1-62.
Wang, Y., Chen, C.-L. and Iijima, M. (2011). Signaling mechanisms for chemotaxis.
Dev. Growth Differ. 53, 495-502.
Wessels, D. J., Zhang, H., Reynolds, J., Daniels, K., Heid, P., Lu, S., Kuspa, A.,
Shaulsky, G., Loomis, W. F. and Soll, D. R. (2000). The internal phosphodiesterase
RegA is essential for the suppression of lateral pseudopods during Dictyostelium
chemotaxis. Mol. Biol. Cell 11, 2803-2820.
Wessels, D., Lusche, D. F., Kuhl, S., Heid, P. and Soll, D. R. (2007). PTEN plays a
role in the suppression of lateral pseudopod formation during Dictyostelium motility
and chemotaxis. J. Cell Sci. 120, 2517-2531.
Wessels, D. J., Kuhl, S. and Soll, D. R. (2009). Light microscopy to image and quantify
cell movement. Methods Mol. Biol. 571, 455-471.
Yumura, S. and Fukui, Y. (1985). Reversible cyclic AMP-dependent change in
distribution of myosin thick filaments in Dictyostelium. Nature 314, 194-196.
Yumura, S. and Uyeda, T. Q. (1997). Myosin II can be localized to the cleavage furrow
and to the posterior region of Dictyostelium amoebae without control by
phosphorylation of myosin heavy and light chains. Cell Motil. Cytoskeleton 36,
Yumura, S., Mori, H. and Fukui, Y. (1984). Localization of actin and myosin for the
study of ameboid movement in Dictyostelium using improved immunofluorescence.
J. Cell Biol. 99, 894-899.
Yumura, S., Furuya, K. and Takeuchi, I. (1996). Intracellular free calcium responses
during chemotaxis of Dictyostelium cells. J. Cell Sci. 109, 2673-2678.
Yumura, S., Yoshida, M., Betapudi, V., Licate, L. S., Iwadate, Y., Nagasaki, A., Uyeda,
assembly control and proper cytokinesis in Dictyostelium. Mol. Biol. Cell 16, 4256-4266.
Zigmond, S. H. (1977). Ability of polymorphonuclear leukocytes to orient in gradients
of chemotactic factors. J. Cell Biol. 75, 606-616.
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