JCS ePress online publication date 28 June 2011
2521
Short Report
Myosin-1C associates with microtubules and
stabilizes the mitotic spindle during cell division
Agrani Rump1,*, Tim Scholz2,*, Claudia Thiel1, Falk K. Hartmann1, Petra Uta2, Maike H. Hinrichs2,
Manuel H. Taft1 and Georgios Tsiavaliaris1,‡
1
Laboratory for Cellular Biophysics, Institute for Biophysical Chemistry, Hannover Medical School, 30625 Hannover, Germany
Institute for Molecular and Cell Physiology, Hannover Medical School, 30625 Hannover, Germany
2
*These authors contributed equally to this work
‡
Author for correspondence ([email protected])
Journal of Cell Science
Accepted 26 April 2011
Journal of Cell Science 124, 2521-2528
© 2011. Published by The Company of Biologists Ltd
doi:10.1242/jcs.084335
Summary
The mitotic spindle in eukaryotic cells is composed of a bipolar array of microtubules (MTs) and associated proteins that are required
during mitosis for the correct partitioning of the two sets of chromosomes to the daughter cells. In addition to the well-established
functions of MT-associated proteins (MAPs) and MT-based motors in cell division, there is increasing evidence that the F-actin-based
myosin motors are important mediators of F-actin–MT interactions during mitosis. Here, we report the functional characterization of
the long-tailed class-1 myosin myosin-1C from Dictyostelium discoideum during mitosis. Our data reveal that myosin-1C binds to MTs
and has a role in maintenance of spindle stability for accurate chromosome separation. Both myosin-1C motor function and taildomain-mediated MT–F-actin interactions are required for the cell-cycle-dependent relocalization of the protein from the cell periphery
to the spindle. We show that the association of myosin-1C with MTs is mediated through the tail domain. The myosin-1C tail can
inhibit kinesin motor activity, increase the stability of MTs, and form crosslinks between MTs and F-actin. These data illustrate that
myosin-1C is involved in the regulation of MT function during mitosis in D. discoideum.
Key words: Myosin, Mitosis, Microtubules, Actin
Introduction
With their ability to mediate dynamic interactions between the
cytoskeleton and membrane compartments by exerting motor
domain driven force along actin filaments, the single-headed, nonfilamentous class-1 myosins have key roles in cell motility,
organelle transport and cytoskeleton remodeling (Coluccio, 1997;
McConnell and Tyska, 2010). Class-1 myosins localize to cortical
regions of high actin turnover during cell migration, associate with
cell–cell junctions, bind endocytic vesicles, and support the
formation and maintenance of actin-rich protrusive structures during
endocytosis (Kim and Flavell, 2008). The eukaryotic model system
Dictyostelium discoideum has been used extensively to study class1 myosins in processes of the endocytic pathway and cell movement
(Falk et al., 2003; Jung et al., 1996; Neuhaus and Soldati, 1999).
Of the seven class-1 myosins produced in D. discoideum, the longtailed members myosin-1B, myosin-1C and myosin-1D have been
implicated in the management of cortical tension during endocytosis
(Novak et al., 1995; Morita et al., 1996); they are important for
efficient cell motility (Wessels et al., 1991; Jung et al., 1996), and
essential for the recycling of plasma membrane components from
endosomes back to the cell surface (Neuhaus and Soldati, 2000).
Compared with the shorter isoforms with which they share the
small basic phospholipid-binding TH1 domain that mediates
membrane interactions, long-tailed class-1 myosins are
characterized by two additional tail homology domains (TH2 and
TH3) (Hokanson and Ostap, 2006). TH2 is a Gly–Pro–Ala-rich
domain capable of F-actin binding (Doberstein and Pollard, 1992;
Vargas et al., 1997), which, together with TH3, a Src-homology 3
domain, links the motors to the machinery responsible for actin
dynamics (Soldati, 2003).
Although class-1 myosins have been widely investigated in
vegetative cells during interphase (Falk et al., 2003; Rivero et al.,
2008), their functions during mitosis remain still elusive. Here, we
characterized D. discoideum myosin-1C, in addition to its
endocytotic roles (Peterson et al., 1995; Dieckmann et al., 2010),
as a high-affinity MT-binding myosin that associates with the
spindle throughout mitosis. We show that the myosin-1C tail
domain affects the dynamic instability of MTs by protecting them
from depolymerization. Impairment of myosin-1C function causes
defects in spindle assembly and positioning, which results in
perturbed mitotic progression. Thus, myosin-1C appears to be
important for the structural integrity of the entire spindle to facilitate
high-fidelity chromosome separation during mitosis.
Results and Discussion
Myosin-1C associates with microtubules of the mitotic
spindle
The subcellular distribution of myosin-1C in D. discoideum was
analyzed using AX2 cells and myosin-1C-knockout cells, producing
recombinant full-length myosin-1C tagged either at the N- or Cterminus with YFP. Western blot analysis of whole cell lysates
revealed comparable protein levels of recombinant and endogenous
myosin-1C (supplementary material Fig. S1A). Confocal analysis
of fixed and living cells showed myosin-1C at the plasma
membrane, crown-like cortical structures, and endosomes that were
formed during phagocytic and pinocytic uptake (Fig. 1A,B). This
localization pattern resembles that of endogenous protein (Peterson
et al., 1995), indicating that the presence and position of the YFP
tag did not affect the dynamic distribution of the motor. During
yeast internalization, myosin-1C gradually accumulated at the
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Fig. 1. Cellular localization of myosin-1C during endocytosis and mitosis. (A) Confocal time series of a Dictyostelium cell ingesting Atto-633-labeled yeast
(red). During uptake, myosin-1C (green) is localized at the cell membrane of the phagocytic cup that surrounds the yeast cell (arrows). (B) Localization of myosin1C during Texas-Red–Dextran uptake. Myosin-1C is associated with the membrane at macropinocytic cups (arrows). (C) Time lapse of a mitotic cell showing that
myosin-1C associates with the spindle. (D) Confocal images of YFP–myosin-1C cells stained for chromatin during interphase, mitosis and cytokinesis. At
interphase, myosin-1C is concentrated at the plasma membrane. In mitotic cells, myosin-1C localizes at the spindle poles during prophase. From metaphase to
anaphase, myosin-1C is concentrated along the entire mitotic spindle, including newly formed astral MTs. At telophase, myosin-1C is enriched at the spindle poles
and associates with astral MTs that extend towards the cell cortex. Scale bars: 5 m.
entire phagosomal membrane and disappeared from the proximal
phagosomal region after completion of phagocytosis. The
enrichment at phagocytic cups suggests a similar role of myosin1C in exerting force at membrane extensions of endocytic cup
pseudopods as reported for myosin-1D (Morita et al., 1996; Dai et
al., 1999). During mitosis, however, myosin-1C was completely
absent from the cell periphery and localized to the cell center. Live
imaging of mitotic cells revealed a well-defined enrichment of the
protein at structures indicative of the spindle apparatus. This new
and striking myosin-1C localization pattern is depicted in Fig. 1C
as a confocal time-lapse sequence from metaphase until the
complete detachment of the daughter cells.
To obtain a more detailed picture of the temporal and spatial
localization of myosin-1C with the spindle, we analyzed the
distribution of the protein in synchronized and fixed cells at
different mitotic phases (Fig. 1D). D. discoideum undergoes a
closed mitosis in which the spindle is formed inside the nucleus
and MTs penetrate the intact nuclear envelope (Moens, 1976).
Because the nuclear envelope is maintained throughout the cell
cycle, it forms a barrier for nuclear import. This excludes
unspecific enrichment of YFP–myosin to the nucleus. During
prometaphase, myosin-1C was mainly present at the spindle
poles. In metaphase cells, myosin-1C decorated the entire length
of the central spindle. From metaphase to telophase, spindle
association persisted and extended to astral MTs. During
cytokinesis, myosin-1C was mainly present at the spindle poles
and astral MTs that reached the cell cortex. The disappearance of
myosin-1C from the center of the spindle is concomitant with the
decomposition of MTs during spindle breakdown upon cleavage
furrow constriction. Closer analysis of mitotic cells showed
colocalization of myosin-1C with both - tubulin and -tubulin
(Fig. 2A, supplementary material Fig. S1B). In synchronized,
G2-phase-arrested cells (Weijer et al., 1984), myosin-1C
exclusively localized at the periphery of centrosomes
(supplementary material Fig. S1C), but did not colocalize with
the centrosomal protein Spc97 (supplementary material Fig. S1D),
suggesting that before mitosis, myosin-1C specifically binds to
MTs that emanate from the centrosomal corona (Euteneuer et al.,
1998). The observed stepwise association of myosin-1C with
centrosomal, interzonal, and astral MTs during mitotic progression
supports a role of the motor in contributing to MT dynamics of
the spindle, a process that needs to be tightly regulated to ensure
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Function of myosin-1C in mitosis
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Fig. 2. The myosin-1C tail binds MTs and mediates crosslinking of MTs to F-actin. (A) Confocal images of an anaphase cell immunostained for -tubulin and
chromatin showing YFP–myosin-1C co-localized with MTs of the entire spindle apparatus. (B) TIRFM images reveal direct binding of YFP–TH1-TH2-TH3 to
surface-immobilized Cy5-labeled MTs. (C) Domain organization and schematic drawings of myosin-1C constructs with amino acid numbering on top. (D) TRITC–
phalloidin-labeled F-actin binds to surface-immobilized myosin-1C tail molecules. (E) Fluorescence images of surface-immobilized Cy5-labeled MTs (red)
incubated with TRITC–phalloidin-labeled F-actin (green) in the presence of 58 nM myosin-1C tail reveal the crosslinking ability of the myosin-1C tail.
(F) Confocal section of a mitotic YFP–myosin-1C (green) cell double stained for F-actin (red) and chromatin (blue). Scale bars: 5 m.
proper spindle assembly or positioning, and subsequent spindle
breakdown for successful cell division.
The myosin-1C tail binds MTs, modulates their dynamics
and crosslinks them to F-actin
To test whether myosin-1C has a role in MT dynamics, we studied
the protein in more detail in vitro. Because the elements targeting
class-1 myosins to distinct cellular locations are implicated to
reside within the C-terminal domains (Ruppert et al., 1995), we
generated a set of recombinant tail-truncation constructs (Fig. 2C)
to test for direct MT binding and dissect the functional properties
of the individual TH domains. Quantitative analyses of GST pulldowns and co-sedimentation assays revealed nanomolar affinity
binding of the entire tail (TH1-TH2-TH3) to MTs (supplementary
material Fig. S2). The specificity of this interaction was further
investigated by in vitro TIRF microscopy. A fluorescent myosin1C tail construct either fused to YFP (Fig. 2B) or (immuno-)
labeled with rhodamine or quantum dots (supplementary material
Fig. S3B,C), bound in all examined cases along the entire length
of MTs (n=232 MTs). In addition to the previously reported F-
actin binding property of the myosin-1C tail domain (Jung and
Hammer, 1994), which we could confirm by TIRF assays (Fig. 2D,
supplementary material Fig. S3E), our experiments revealed further
that the tail is capable of binding MTs and F-actin simultaneously
(Fig. 2E, supplementary material Fig. S3D); thus crosslinking of
both filamentous structures occurs without the need for additional
factors. Consistent with this finding, we observed that in mitotic
cells, myosin-1C colocalized with astral MTs (Fig. 2A) and F-actin
in regions close to the spindle poles and the cell cortex (Fig. 2F).
In other cytoplasmic compartments they were clearly separated.
This suggests a role of myosin-1C in mediating spindle attachment
to cortical actin sites of the leading edge of dividing cells
(Hestermann et al., 2002).
To elucidate the functional relevance of the observed MT
interactions, we first analyzed the effect of tail binding on MT
dynamics by TIRF microscopy. The cold-inducible
depolymerization of reconstituted MTs occurred at rate of kdepol=0.5
m minute–1 (Fig. 3A,B). MTs pretreated with saturating
concentrations of tail construct TH1-TH2-TH3, however,
maintained their initial length (Fig. 3A). By taking advantage of
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Journal of Cell Science
depolymerization rates decreased hyperbolically yielding a half
maximal effector concentration of Ki=2.4±1 nM. The TH1-TH2
domain alone stabilized MTs with Ki=2.2±1 nM, indicating that
the TH3 domain is not essential for MT stability (Fig. 3B). The
TH1 domain had no effect on MT dynamics, although it bound to
MTs in cosedimentation assays (supplementary material Fig. S2A).
The TH2 domain prevented MT depolymerization with 100-times
lower efficiency (Ki=246±55 nM) than constructs containing both
TH1 and TH2 domains. However, an equimolar mixture of the
isolated TH1 and TH2 domains had no influence on MT
disassembly (Fig. 3B, filled circle). Taken together, the results
reveal that effective stabilization of MTs requires the action of the
entire TH1-TH2 polypeptide chain. The data imply that the TH2
domain harbors the MT stabilizing properties, whereas the TH1
domain contributes to increasing MT affinity. Because the TH1TH2 domains are composed of highly repetitive Pro-Lys-rich
motifs, it is possible that charge–charge interactions are responsible
for high-affinity binding. This is supported by our observation that
the affinity for MTs is ionic strength-dependent (KD=2±1.4 nM at
25 mM KCl versus 80±50 nM at 150 mM KCl) (supplementary
material Fig. S2B). We further assessed for a possible role of the
myosin-1C tail in MT assembly by monitoring the time-dependent
turbidity changes with increasing concentrations of tubulin.
However, MT nucleation and elongation were not affected
(supplementary material Fig. S4).
Myosin-1C tail affects kinesin motility along MTs
Fig. 3. The myosin-1C tail prevents microtubule depolymerization and
affects the binding properties of kinesin-1 to MTs. (A) TIRFM images
showing the cold-induced shortening of surface-immobilized Cy5-labeled MTs
(top panel). In the presence of 115 nM myosin-1C tail construct TH1-TH2TH3, MT depolymerization is completely suppressed (bottom panel). Scale
bars: 5 m. (B) Concentration dependence of the rate of cold-induced MT
depolymerization for different myosin-1C tail constructs. Addition of
increasing concentrations of myosin-1C tail constructs TH1-TH2-TH3 (䉬) and
TH1-TH2 (䉫) reduce kdepol hyperbolically with a half maximal effector
concentration of Ki=2.4±1 nM (dashed line) and 2.2±1 nM (solid line),
respectively. TH2 (䉭) prevents MT depolymerization with a half maximal
concentration of Ki=246±30 nM (dashed and dotted line). TH1 (䉱) and an
equimolar mixture of 115 nM TH1 and TH2 (䊉) have no effect on the thermal
stability of MTs. Error bars are s.e.m. Levels of significance are indicated with
**P<0.01 and ***P<0.001. (C) Kinesin motility along MTs decorated with
myosin-1C tail. Attachment frequencies of single mCherry-labeled kinesin-1
molecules on MTs in the presence of increasing concentrations of myosin-1C
tail. The attachment frequency of rK592 to MTs decreases hyperbolically with
increasing myosin-1C tail concentrations (I=1.2±0.5 nM). Inset shows mean
velocity (䉬) and run length () (䉫) are not affected by myosin-1C tail. Error
bars are s.e.m. or standard fit errors.
this stabilizing effect, we performed titration experiments with
different tail domain constructs to test for their ability to prevent
MT decomposition and determine the apparent equilibrium
constants of this process. In the case of the entire tail,
Because the binding sites of MAPs on the MT lattice can partially
overlap with those of kinesin (Seitz et al., 2002), we tested whether
myosin-1C binding to MTs interfered with the processive properties
of kinesin motors. Indeed, we observed a myosin-1C tail
concentration-dependent reduction of the attachment frequency of
single kinesin-1 molecules on MTs (Fig. 3C). Although velocity
and processive run length were not affected (Fig. 3C, inset), kinesin
attachment frequencies decreased hyperbolically, yielding a halfmaximal effector concentration of 1.2 nM. This value is comparable
with the apparent equilibrium constant (Ki) for tail binding to MTs.
Kinesin itself did not affect myosin-1C tail binding to MTs
(supplementary material Fig. S3F). These results indicate that the
motors do not share the same MT interaction sites. Secondary
structural analysis of the tail domain by circular dichroism (data
not shown) revealed that both TH1 and TH2 are highly unstructured
domains with low -helical and -strand contents. This predicted
intrinsically disordered tail structure might effectively hinder
kinesin molecules from efficient MT binding by blocking sterically
and/or electrostatically interaction sites on the MT lattice. Myosin1C could thus have a role in regulating kinesin function in mitotic
processes.
Myosin-1C contributes to spindle stability
To test whether the tail domain itself targets myosin-1C to the
mitotic spindle or whether motor function is an additional
prerequisite for the relocalization mechanism, we expressed YFPtagged constructs lacking either the motor domain or the entire tail
in AX2 cells. The motor domain exhibited diffuse cytoplasmic
staining (supplementary material Fig. S5B), whereas the tail
accumulated in punctuated structures of the cytoplasm, with no
obvious spindle association (supplementary material Fig. S5C). A
full-length myosin-1C construct lacking the TH3 domain, however,
displayed normal spindle association (supplementary material Fig.
S5D). The latter observation agrees with the results discussed
Function of myosin-1C in mitosis
We therefore followed a dominant-negative approach to investigate
a possible mitotic function of myosin-1C. Dominant-negative
inhibition has been a successful tool for the characterization of
myosins in vivo (Tsiavaliaris et al., 2002; Mulvihill et al., 2006).
In particular, this approach can be helpful when morphological
changes from gene deletion are not apparent because of the presence
of functionally redundant protein isoforms. The coexpression of a
full-length myosin-1C construct (YFP–myosin-1CG395A) containing
a single point mutation in the motor domain, which is known to
reduce actin affinity and perturb motor properties (Sasaki et al.,
1998) indeed induced similar defects in growth and endocytic
behavior, as seen for myosin-1CKO cells (supplementary material
Fig. S6). More interestingly, we observed morphological changes
Journal of Cell Science
earlier that the TH3 domain neither contributes to MT binding nor
affects MT stability. In summary, these observations indicate that
both a functional motor domain and the TH1-TH2 domains are
essential for the targeting mechanism of myosin-1C to MTs of the
spindle apparatus.
In agreement with a role for myosin-1C in endocytosis, myosin1C-deficient cells (myosin-1CKO) exhibited reduced efficiency of
fluid and particle uptake (supplementary material Fig. S6C,D),
which, in turn, affected their axenic (Fig. 4D) and bacteria-assisted
growth (supplementary material Fig. S6B). During mitosis, myosin1CKO cells displayed no apparent phenotype related to MT
organization. Confocal analysis revealed typical MT patterns and
normal spindle architecture (supplementary material Fig. S5A).
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Fig. 4. Dominant negative inhibition of myosin-1C. (A) Superimposed confocal image sections of mitotic wild-type (bottom row) and YFP–myosin-1CG395A
cells stained for -tubulin and chromatin. Misaligned chromosomes along partially formed spindles during mitosis (first and third row) and defects in chromosome
separation (second row) are observed in cells with increased YFP–myosin-1CG395A expression levels. Scale bars: 5 m. (B) The size of nuclei in YFP–myosin1CG395A cells follows a broad distribution with an average nuclear size of 55 m2, whereas untransfected cells and cells expressing YFP–myosin-1C display a
narrow distribution of nuclear sizes of 34 and 35 m2, respectively. Dashed lines are Gaussian fits to the data. (C) Cells producing mutant YFP–myosin-1CG395A
have increased levels of chromatin than YFP–myosin-1C producing cells and AX2 cells. Error bars represent s.d. The inset depicts DAPI-stained nuclei of YFP–
myosin-1CG395A cells with small and large nuclei. Cells with larger nuclei have increased levels of YFP–myosin-1CG395A (green). (D) Growth of wild-type (AX2),
myosin-1C-knockout cells, YPF–myosin-1C, and YFP–myosin-1CG395A cells in shaking cultures.
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in the MT patterns of these cells during mitosis, indicating that the
mutant competes with endogenous myosin-1C function. In
comparison with wild-type cells (Fig. 4A, bottom row), cells
overexpressing YFP–myosin-1CG395A showed irregular spindle
patterns with misaligned chromosomes (Fig. 4A, first row). In
some cases, cells underwent cytokinesis before chromosomal
separation was completed (Fig. 4A, second row), and the formation
of interzonal MTs in prometaphase cells was strongly affected
(Fig. 4A, third row). As a consequence of impaired spindle function,
we observed that YFP–myosin-1CG395A cells contained nuclei that
were on average twice as large as those in non-transfected cells or
in cells producing wild-type myosin (Fig. 4B). In agreement with
potential defects in chromosome separation, the nuclear chromatin
content of these cells was increased twofold (Fig. 4C). Compared
with the wild type, mitosis of dominant-negative cells was
prolonged approximately threefold (supplementary material Fig.
S7A). In particular, the progression from prometaphase to
metaphase, where the assembly of interzonal spindle MTs takes
place, was delayed in the mutant cells (supplementary material
Movies 1, 2) compared with the wild type (supplementary material
Movie 3). When grown in suspension, the fraction of myosin1CKO and myosin-1CG395A cells with one nucleus was decreased in
favor of two and more nuclei (supplementary material Fig. S7B),
indicating an involvement of myosin-1C in the temporal coupling
of mitosis to cytokinesis. Multiple nuclei were not observed in
surface-attached cells, because traction-mediated cytoplasmic
fission and motile activity of the cells support the process of
cytokinesis (Neujahr et al., 1997).
Because Dictyostelium class-1 myosins share overlapping
interphase functions (Jung et al., 1996), members of the same
subclass might also compensate for depletion or dominant-negative
impairment of myosin-1C during mitosis. Two potential candidates
are myosin-1B and myosins-1D. Myosin-1D binds and stabilizes
MTs as efficiently as myosin-1C does (data not shown). Moreover,
the localization pattern of myosin-1D throughout the cell cycle
resembles in many aspects that of myosin-1C. Myosin-1D decorates
the cortical membrane during interphase (supplementary material
Fig. S8E), and is concentrated at the central spindles and nuclear
envelope during mitosis (supplementary material Fig. S8B). This
association is observed throughout mitosis, indicating additional
roles of this motor, for example, in nuclear membrane fission to
daughter cells (data not shown). By contrast, myosin-1B and the
short-tailed isoform myosin-1E clearly lack an association with
MTs during mitosis (supplementary material Fig. S8A,C), but
distribute throughout the cytoplasm (supplementary material Fig.
S8D,F).
It is well established that the actin cytoskeleton, MTs and their
regulated interplay are required for successful chromosome
segregation (Rodriguez et al., 2003; Woolner et al., 2008). Recent
studies demonstrated that myosin-10 is an essential integrator of
spindle MTs with cortical F-actin required for nuclear anchoring
and spindle assembly in Xenopus laevis oocytes (Weber et al.,
2004). Other examples include the association of nonmuscle
myosin-2 (MHC-A) with MTs of the spindle apparatus (Kelley et
al., 1996) and the localization of myosin-5 at the MT organizing
center during melanocyte cell division (Wu et al., 1998). Diffusive
interactions of myosin-5 with MTs are assumed to be required for
efficient and targeted cargo delivery from the cell center to the
periphery (Ali et al., 2007).
Here, we characterized myosin-1C as the first class-1 myosin
that undergoes dynamic and cell-cycle-specific changes from
cortical regions in interphase to the MT-rich spindle during mitosis.
Our in vitro results show that myosin-1C might mediate direct
interactions between the F-actin and the MT cytoskeleton. With its
unique ability to effectively stabilize MTs and potentially serve as
a dynamic linker between both cytoskeletal networks, myosin-1C
appears to be important for normal spindle function to ensure high
fidelity of cell division. Both head and tail interactions are important
for the cell-cycle-dependent relocalization of the protein.
Coordinated mechanisms that switch motile activity on and off
(Fujita-Becker et al., 2005; Dürrwang et al., 2006) and modulate
tail-mediated MT–F-actin–lipid interactions, e.g. by a fold-back
mechanism of the tail (Stöffler and Bähler, 1998; Hwang et al.,
2007), might regulate the temporal association of myosin-1C with
cortical membranes during endocytosis, and with the mitotic spindle
during cell division. Our study reveals that the functions of myosin1 are not limited to interphase processes that occur at the F-actinmembrane interface, but extend to the MT network and its dynamic
organization during mitosis.
Materials and Methods
Plasmid construction
The sequence encoding myosin-1C comprising 1182 amino acids (aa) was amplified
from D. discoideum genomic DNA by PCR (using the primers myoCfw and myoCrv
and cloned in pDXA-3H-eYFP-mcs (Knetsch et al., 2002). Mutation G395A was
introduced by PCR-directed mutagenesis using primer myoG395A. Sequences
encoding myosin-1C tail domain constructs TH1-TH2-TH3 (aa 770–1182), TH1TH2 (aa 770–1136), TH1 (aa 770–982) and TH2 (aa 975–1136) were amplified by
PCR and cloned in pGEX-6P2 for expression in E. coli as N-terminal GST fusions
and in pDXA–YFP–mcs as N-terminal YFP fusion for the expression in D.
discoideum cells, respectively. The myosin-1C tail construct YFP–TH1-TH2-TH3
with N-terminal His6 tag fused to YFP was obtained from pDXA–YFP–MIC-tail by
PCR and ligated into the pProExHta vector using restriction sites EcoRI and XbaI.
All constructs were confirmed by sequencing.
Cell lines, protein expression and purification
Cultivation of D. discoideum cells was performed as described previously (Dürrwang
et al., 2006). Synchronization experiments were performed as described (Nellen and
Saur, 1988; Tuxworth et al., 1997). GST fusion constructs were expressed in E. coli
Rosetta pLys-S cells (Merck, Darmstadt). Cells were grown at 30°C in YT medium
(16 g Tryptone, 10 g yeast extract, 5 g NaCl per liter pH 7.0) and induced with 0.5
mM IPTG at OD600=0.6, grown at 21°C for 16 hours, harvested by centrifugation
(4°C, 4000 g). Cell lysis was performed at 4°C for 30 minutes in Buffer (50 mM
Tris-HCl, pH 8.0, 300 mM NaCl, 1 mM MgCl2, 1 mM EDTA, 5 mM benzamidine,
5 mM DTT, 4 mM PMSF, 0.5 mg/ml lysozyme, four pellets Complete inhibitor
cocktail (Roche, Penzberg, Germany), 1000 U benzonase (Merck, Darmstadt,
Germany) and lyzed with Triton X-100 (1% final concentration). After centrifugation
at 20,000 g at 4°C (Avanti J-30I, Beckmann Coulter) the supernatants were applied
to a glutathione-Sepharose column, washed with buffer (50 mM Tris-HCl, pH 7.5,
300 mM NaCl, 1 mM EDTA, 2 mM DTT, 2 mM benzamidine). A linear gradient of
Buffer A (50 mM Tris-HCl, pH 8.0, 300 mM NaCl) and Buffer B (Buffer A
containing 10 mM reduced glutathione) eluted the protein containing fractions,
which were dialyzed against Storage Buffer (25 mM Tris-HCl, pH 7.5, 50 mM
arginine, 50 mM glutamate, 300 mM NaCl, 2 mM DTT, 3% Sucrose), snap-frozen
in liquid nitrogen, and stored at –80°C.
Fluorescence microscopy
Phagocytosis and pinocytosis assays were performed as described previously (Rivero
and Maniak, 2006). For live cell imaging, cells were seeded and washed twice with
Mes buffer (20 mM Mes, pH 6.8, 0.2 mM CaCl2, 2 mM MgCl2). For
immunofluorescence, cell fixation was performed in 20 mM PIPES (pH 6.8)
containing 3% paraformaldehyde. Cells were permeabilized in PBS containing 0.1%
Triton X-100 for 5 minutes, washed with PBS, and blocked with Image-iT Signal
Enhancer. Cells were incubated with 4 g/ml anti--tubulin antibody in PBS
containing 3% BSA for 1 hour and washed with PBS followed by incubation with
Alexa-Fluor-633-labeled secondary antibody (1:500 in PBS) for 1 hour. Chromatin
was stained with 0.1 g/ml 4⬘,6⬘-diamidino-2-phenylindole (DAPI) for 5 minutes.
Coverslips were mounted on glass slides with Slow Fade Gold. Images were recorded
with an IX-81 inverted microscope (Olympus, Germany). The microscope set-up
ScanR (Olympus, Munich) was used for the statistical analysis of the chromatin
content in wild-type and mutant cell lines. For confocal microscopy Leica TCS-SP2
AOBS and Olympus FV-1000 systems equipped with 63⫻, 1.4 NA oil objectives
were used.
Function of myosin-1C in mitosis
Journal of Cell Science
Protein interactions and functional analyses using TIRF microscopy
A self-made objective type total internal reflection fluorescence (TIRF) microscope
with single-fluorophore sensitivity was used for in vitro microscopy (Amrute-Nayak
et al., 2008). For Cy5 excitation, a HeNe laser (633 nm, 35 mW, Coherent) was used.
Emitted light was recorded with a back-illuminated Andor iXon DV887 EMCCD
camera (Andor Technology, Belfast, UK). MT immobilization was performed as
described (Helenius et al., 2006) and incubated for 1 minute with 0.1–1.4 M YFP–
myosin-1C tail in buffer BRB12 (12 mM PIPES, 2 mM MgCl2, 1 mM EGTA, pH
6.8). After washing with buffer BRB12 containing 0.5 mg/ml BSA and an oxygen
scavenger system (800 U/ml catalase, 10 mg/ml glucose, 10 U/ml glucose oxidase,
and 10 mM DTT), images of myosin-1C-decorated MTs were recorded. F-actin
interaction assays were performed by binding TRITC–Phalloidin-labeled F-actin to
100 nM surface-immobilized myosin-1C tail pre-incubated with buffer BRB12
containing 5 mg/ml BSA. Myosin-1C-tail-mediated F-actin–MT crosslinking was
performed with taxol-stabilized Cy5–MTs that were pre-incubated for 1 minute with
58 nM GST–TH1-TH2-TH3 in buffer BRB12. After blocking with 0.5 mg/ml BSA,
TRITC–phalloidin-labeled F-actin was injected into the chamber and incubated for
1 minute. MT–F-actin assembly was recorded after rinsing again. MT
depolymerization assays were performed using GMPCPP-treated Cy5-labeled MTs
(Hyman et al., 1992). After incubation for 1 minute with various concentrations of
different myosin-1C tail constructs at 35°C, MT depolymerization was induced by
the pre-cooled (10°C) objective stage of the TIRF microscope and recorded for 10
minutes at time intervals of 1 frame every 10 seconds at a light exposure time of 100
ms per frame. The rate of MT depolymerization was determined by measuring the
reduction in length of individual MTs over time using Andor Solis software. Fits to
plots of depolymerization versus myosin-1C tail construct concentration were
performed using Origin 7.0 (OriginLab, Northampton, MA). Motility assays of
mCherry-tagged rat kinesin-1 molecules (592 aa of rat kinesin tagged at the Cterminus with mCherry) along GMPCPP-stabilized Cy5-labeled MTs were performed
at 23°C in the presence of 1 mM ATP at different myosin-1C tail concentrations. Run
lengths of single kinesin-1 molecules distributed according to an exponential decay
and were, therefore, fitted to yield the run length decay constant  (Seitz et al.,
2002).
Tubulin polymerization assays
Tubulin polymerization (nucleation and elongation) was followed by turbidity
measurements at 340 nm (Johnson and Borisy, 1975) using a NanoDrop 2000c
Spectrophotometer (Thermo Scientific) pre-heated to 37°C in buffer BRB80 (80
mM PIPES, 2 mM MgCl2, 1 mM EGTA, pH 6.8) containing 1 mM GTP and 5%
glycerol. To monitor MT nucleation, different concentrations of tubulin in BRB80
containing 1 mM GTP were incubated at 37°C with and without 100 nM myosin1C tail.
We thank C. Waßmann for excellent technical assistance, Margaret
A. Titus (University of Minnesota, MN) for the myosin-1C null cells,
Ralph Gräph (University of Potsdam, Germany) and Thierry Soldati
(Université de Genève, Switzerland) for antibodies, W. Kammerloher
(Olympus, Munich) for support with the ScanR system, B. Brenner and
D. J. Manstein for helpful discussions. This work was supported by
DFG grants SCHO-1213/1 to T.S. and TS-169-3/1 to G.T.
Supplementary material available online at
http://jcs.biologists.org/cgi/content/full/124/15/2521/DC1
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