Department of Cellular Biology, University of Georgia, Athens, Georgia 30602, USA
Author for correspondence (e-mail:
fechheim{at}cb.uga.edu)
Accepted 14 October 2002
![]() |
Summary |
---|
![]() ![]() ![]() ![]() ![]() ![]() ![]() |
---|
Key words: Cytoskeleton, Motility, EF hands, Calcium regulation, Actin-binding protein, Dictyostelium
![]() |
Introduction |
---|
![]() ![]() ![]() ![]() ![]() ![]() ![]() |
---|
The calcium-regulated 34 kDa protein has been localized to the cell cortex,
filopodia and pseudopodia, phagocytic cup and cell-to-cell contact sites
(Fechheimer, 1987;
Fechheimer et al., 1994
;
Furukawa et al., 1992
;
Furukawa and Fechheimer, 1994
;
Johns et al., 1988
;
Okazaki and Yumura, 1995
). The
34 kDa protein has been proposed to play a role in regulating the viscoelastic
properties of the actin cytoskeleton. In vitro studies show that the 34 kDa
protein is able to crosslink F-actin into bundles in a calcium-regulated
manner. The 34 kDa protein bundles actin at low calcium
(1x10-8 M), but at elevated calcium levels
(1x10-6 M) the protein is unable to bundle F actin
(Fechheimer, 1987
;
Fechheimer and Taylor, 1984
;
Lim and Fechheimer, 1997
). The
cDNA sequence of the 34 kDa protein indicates that the protein has 295 amino
acids, with two putative EF hand regions
(Fechheimer et al., 1991
).
Mutants lacking the 34 kDa protein grow and develop normally at 20°C but
have abnormal filopodia, decreased persistence of motility and are
cold-sensitive for growth (Rivero et al.,
1996
; Rivero et al.,
1999
). Cells lacking two calcium-sensitive actin-crosslinking
proteins,
-actinin and the 34 kDa protein, grow slowly at 15°C and
20°C, endocytose fluid phase slowly, produce small cells and undergo
morphogenesis to produce aberrant fruiting bodies
(Rivero et al., 1999
).
To test the hypothesis that calcium regulation of actin crosslinking is
important, we first investigated the basis of calcium binding to the 34 kDa
protein. The EF hand regions of the 34 kDa protein were modified by
site-directed mutagenesis to change critical charged amino acids at the
coordinating X, Y, Z positions to uncharged alanines
(Fig. 1A). In vitro studies of
these proteins show that the 34 kDa protein has one high-affinity
calcium-binding site, which resides in the putative second EF hand. However,
the actin binding and crosslinking of the 34 kDa EF2 protein is not
regulated by calcium. We have expressed the 34 kDa wild-type and 34 kDa
EF2 proteins in mutant Dictyostelium cell lines lacking the 34
kDa protein or both 34 kDa protein and
-actinin
(Rivero et al., 1996
;
Rivero et al., 1999
). The
results reveal that expression of the 34 kDa
EF2 protein did not
restore function to the 34-kDa-null or 34 kDa/
-actinin-null cell lines.
These results show that calcium regulation is necessary for 34 kDa protein
function and that calcium regulation of actin filament crosslinking is
essential for normal function and dynamics of the actin cytoskeleton.
|
![]() |
Materials and Methods |
---|
![]() ![]() ![]() ![]() ![]() ![]() ![]() |
---|
Analytical methods and antibodies
Protein concentration was determined according to the bicinchoninic acid
(BCA; Pierce Chemical Co., St. Louis, MO) method
(Smith et al., 1985) using
bovine serum albumin (Sigma Chemical Co., St Louis, MO) as a protein standard.
Monoclonal anti 34 kDa antibody B2C
(Furukawa et al., 1992
) and
alkaline phosphatase-conjugated goat anti-mouse antibodies (Promega Corp.,
Madison, WI) were used for western blot analysis as described previously
(Towbin et al., 1979
).
Measurement of calcium binding by equilibrium dialysis
Ultra pure water (Continental Water Systems Corporation, San Antonio, TX)
and buffers were passed over a 2x30 cm Chelex 100 (BioRAD, Hercules, CA)
resin to remove any traces of calcium. All plastic ware was rinsed in
calcium-free water before use. The proteins were first dialyzed against
storage buffer (10 mM Tris pH 7.0, 50 mM KCl, 0.2 mM DTT, 0.1 mM EDTA) for 24
hours at 4°C. To remove EDTA, the proteins were dialyzed in storage buffer
as above but without the EDTA for 24 hours with one buffer change at 4°C.
For calcium-binding measurements, the proteins were dialyzed in 40 ml of
storage buffer without EDTA containing different concentrations of
45Ca[CaCl2] for 48 hours at 25°C. To achieve
different concentrations of calcium in the buffer, calcium was added from a
200 µM calcium stock solution that contained approximately 10
µCi/µmole 45Ca (DuPont NEN, Boston, MA). Protein
concentrations of samples were determined after dialysis. After dialysis, 100
µl of buffer from the outside and 100 µl of sample from inside the
dialysis bag was sampled in triplicate, and radioactivity was determined by
liquid scintillation counting. To determine the amount of calcium bound, the
number of counts outside was averaged and subtracted from the averaged counts
inside and divided by the specific activity of 45Ca.
Actin
G-actin was isolated from rabbit skeletal muscle acetone powder and
gel-filtered as described previously
(MacLean-Fletcher and Pollard,
1980; Spudich and Watt,
1971
). The actin was stored dialyzing in G-actin buffer for one
week with fresh buffer changes every day.
Measurement of binding by co-sedimentation with actin
High-speed actin co-sedimentation assays were performed as outlined
previously (Fechheimer, 1987;
Fechheimer and Taylor, 1984
;
Lim and Fechheimer, 1997
). 3
µM of G-actin was mixed with 3 µM of wild-type 34 kDa or 34 kDa
EF2 proteins in 20 mM PIPES, pH 7.0, 50 mM KCl, 50 µM
MgCl2, 1 mM ATP, 0.2 mM DTT, 5 mM EGTA plus or minus 4.5 mM
CaCl2, in a final volume of 130 µl. Supernatant and pellet
fractions were analyzed by polyacrylamide gel electrophoresis in the presence
of SDS (Laemmli, 1970
),
visualized with Coomassie brilliant blue. The amount of protein in the pellet
and supernatant was quantified by scanning densitometry (Molecular Dynamics,
Sunnyvale, CA). Control experiments verified that none of the 34 kDa proteins
showed any significant sedimentation in the absence of actin in either the
presence or absence of calcium (data not shown). To determine the
stoichiometry of binding of the 34 kDa proteins, high-speed F-actin
co-sedimentation was performed as described above as a function of the 34 kDa
proteins at low free calcium concentrations.
Electron microscopy
Negative staining of mixtures of actin and 34 kDa proteins was performed as
previously described (Fechheimer and
Furukawa, 1993). Briefly, 5 µM G-actin was mixed with 2.5 µM
34 kDa protein or 34 kDa
EF2 protein in 20 mM PIPES, pH 7.0, 50 mM KCl,
50 µM MgCl2, 1 mM ATP, 0.2 mM DTT, 5 mM EGTA plus or minus 4.5
mM CaCl2 and incubated overnight at 4°C. The mixture was
applied to a 300-mesh copper grid coated with 0.3% Formvar and carbon for 30
seconds, washed with the above buffer for 1 minute, stained with 2% uranyl
acetate for 30 seconds and visualized using a Phillips 400 transmission
electron microscope.
Expression of mutant 34 kDa proteins in Dictyostelium
Wild-type or mutant 34 kDa or GFP-fused proteins were expressed in
Dictyostelium after subcloning into either the BamHI site of
the vector pBORP (Ostrow et al.,
1994) or the HindIII/KpnI site of pDXA-GFP
(Levi et al., 2000
). The
coding sequences were taken from the plasmids encoding full-length 34 kDa pET
15b-F18 (Lim and Fechheimer,
1997
) and p34 kDa
EF2 by PCR using primers to modify the
coding sequences with either BamHI, HindIII or KpnI
restriction enzyme sites for the purpose of subcloning. The 34 kDa proteins
were expressed in 34-kDa-null and 34 kDa/
-actinin double null strains
of Dictyostelium described previously
(Rivero et al., 1996
;
Rivero et al., 1999
).
Electroporation and growth of cells was performed as described previously
(Maselli et al., 2002
). The
34-kDa/
-actinin-deficient cells were co-transformed with pBSK-BSR,
derived from pBsR479 (Putka and Zeng,
1998
), to impart blasticidin resistance for selection purposes as
the parental cells and pBORP both utilize G418 resistance cassettes. 24 hours
after transformation the media was removed and replaced with media containing
antibiotics required for selection. The cells were cloned, and expression of
the 34 kDa proteins was assessed by western blotting and immunofluorescence.
Cells were fixed and stained as described previously
(Fechheimer, 1987
).
Rhodamine-labeled or Oregon-green-labeled phalloidin (Molecular Probes,
Eugene, OR) was utilized to localize F-actin, and monoclonal antibody B2C
(Furukawa et al., 1992
),
followed by a rhodamine-labeled secondary antibody was utilized to localize
the 34 kDa protein.
Growth
Dictyostelium AX-2 and derivatives described below were routinely
maintained in axenic shaking cultures at 150 rpm in HL-5 media
(Loomis, 1971) at 20°C. To
assess growth rates, cells were inoculated at a starting concentration of
4x104 cells/ml at 20°C or 15°C in axenic HL-5 culture
media with shaking at 150 or 120 rpm, respectively. Each data point represents
the average of three flasks, all of which were sampled daily in duplicate and
counted on a standard hemocytometer. Growth rates are expressed as doubling
time in hours and were calculated from the daily counts from inoculation until
the first day that plateau was approached.
Cell size
Cells were induced to assume a spherical shape as described previously
(Rivero et al., 1996). The
cells were then allowed to settle in a Bio-unique chamber (Bionique
Laboratories, Saranac Lake, NY), and images were recorded using a Nikon
inverted microscope equipped with a Mighty Max Cooled CCD camera (Princeton,
Trenton, NJ). The major and minor axis of each cell was measured using IP lab
image analysis software (Scanalytics, Fairfax, VA). The diameter of the cells
was determined by averaging the major and minor axis.
Ratio imaging and analysis
Images of vegetative cells fixed and stained as described above were
recorded using a Nikon inverted microscope equipped with a Mighty Max Cooled
CCD camera (Princeton, Trenton, NJ) with the IP lab image analysis software
interface (Scanalytics, Fairfax, VA). Care was taken not to saturate any
pixels in the image. The leading and trailing edges of the amoebae were
identified in the images of either the actin or the 34 kDa protein. These
regions were outlined manually using NIH image. The background noise was
sampled in three rectangular regions close to each of the edges. The average
pixel value of the background was subtracted from the maps. The subsequent
values were averaged over all pixels in the region to obtain the average
intensity value. The ratio of actin to the 34 kDa protein in the trailing and
leading edges was obtained by simple division.
![]() |
Results |
---|
![]() ![]() ![]() ![]() ![]() ![]() ![]() |
---|
Prior studies of calcium binding to the 34 kDa protein employed a
qualitative blot overlay approach
(Fechheimer and Furukawa,
1993; Lim and Fechheimer,
1997
). Therefore, 45Ca[CaCl2] and
equilibrium dialysis were used to determine the number and affinity of
calcium-binding sites in the 34 kDa protein
(Fig. 2). The results reveal
that calcium binds saturably to a single site on the 34 kDa protein with an
affinity of 2.4 µM. By contrast, essentially no calcium binding was
observed using the 34 kDa
EF2 protein
(Fig. 2), demonstrating that
the second EF hand is the site of high-affinity calcium binding to the
wild-type 34 kDa protein.
|
The actin binding and crosslinking activities of the purified 34 kDa
EF2 protein was compared to wildtype using the F-actin co-sedimentation
assay to assess actin binding. All assays were performed in the presence or
absence of micromolar free calcium to investigate calcium regulation of actin
binding. The wild-type 34 kDa protein bound to F-actin substoichiometrically
and with moderate affinity in the co-sedimentation assay at low calcium
(Fig. 3A). In the presence of
micromolar free calcium, the amount of the wild-type 34 kDa protein bound to
F-actin decreased by 70% (Fig.
3B). The wild-type protein forms tight bundles with F-actin in the
absence, but not in the presence, of micromolar calcium
(Fig. 4A,B), in agreement with
previous reports of calcium-regulated actin binding by the native and
recombinant 34 kDa protein (Fechheimer and
Taylor, 1984
; Furukawa and
Fechheimer, 1996
; Lim and
Fechheimer, 1997
).
|
|
The actin-binding activity of the 34 kDa EF2 protein was similar to
that of the wild-type 34 kDa protein at low calcium ion concentrations
(Fig. 3A). However, the 34 kDa
EF2 protein failed to show calcium-sensitive actin binding, as expected
from the absence of calcium binding to this protein
(Fig. 2). In the presence of
micromolar free calcium, the amount of 34 kDa
EF2 protein bound to
F-actin was virtually identical to that bound at low calcium concentrations
(Fig. 3B). Similarly, the 34
kDa
EF2 protein induced formation of actin bundles similar to those
formed with the wild-type protein, and actin bundling by the 34 kDa
EF2
protein was not inhibited in the presence of micromolar calcium
(Fig. 4C,D). Thus, actin
binding by the 34 kDa
EF2 protein is similar to that of the wild-type
protein except for the absence of regulation of the interaction by micromolar
calcium.
Expression of wild-type and mutant 34 kDa proteins in
Dictyostelium
The wild-type and 34 kDa EF2 proteins were expressed in
Dictyostelium 34-kDa-null cells and 34 kDa/
-actinin-null
cells, and the level of expression of these proteins was examined by western
blot with mouse monoclonal antibody B2C to the 34 kDa protein. Expression
levels were similar to those observed for wild-type protein in AX2 cells
(Fig. 1C).
Growth and division are the result of the culmination of a number of
processes involving the actin cytoskeleton. To probe the potential
significance of the EF hands for the function of the 34 kDa protein, the
growth rates of cells expressing the wild-type or 34 kDa EF2 proteins
were examined (Fig. 5A). Under
standard laboratory growth conditions at 20°C, 34-kDa-null cells grow at a
rate similar to that for wild-type AX2 cells (12.1 hours per division) as
reported previously (Rivero et al.,
1996
; Rivero et al.,
1999
). The 34-kDa-null cells and 34-kDa-null cells expressing the
wild-type 34 kDa protein grow at similar rates (12.1 and 13.3 hours per
division, respectively). By contrast, 34-kDa-null cells expressing 34 kDa
EF2 protein grow more slowly (15.3 hours per division).
|
The 34-kDa/-actinin-null cells grow slowly at 20°C compared with
AX2 cells (16.5 compared to 12.7 hours per division) as reported previously
(Rivero et al., 1996
). When
the wild-type 34 kDa protein is expressed in these cells, the wild-type growth
rate is restored (12.3 hours per division). However, expression of the 34 kDa
EF2 protein does not restore the growth rate (15.0 hours per division)
to that observed for AX2 (Fig.
6A).
|
In its natural environment, Dictyostelium must grow in conditions
of varying temperature, osmolarity and nutrient supply that are optimal and
invariant in standard laboratory culture conditions. To provide an
environmental challenge, we grew the cells in shaking culture at 15°C. The
34-kDa-null cells grow slowly (Rivero et
al., 1999) (23.2 hours per division) when compared with wild-type
AX2 (20.1 hours per division) at 15°C. When the wild-type 34 kDa protein
is expressed, the wild-type growth rate is nearly restored (21.5 hours per
division) under these conditions. By contrast, expression of 34 kDa
EF2
protein fails to stimulate the rate of growth of 34-kDa-null cells at
15°C. These cells expressing the 34 kDa
EF2 grow more slowly than
the 34-kDa-null cells (27.7 hours per division)
(Fig. 5B). The effect of
reduced temperature on growth is more pronounced in 34 kDa/
-actinin
double null cells, which grow extremely slowly at 15°C (28.7 hours per
division). Expression of the wild-type 34 kDa protein restores wild-type
growth rate to 21.4 hours. Expression of 34 kDa
EF2 protein has no
restorative effect on growth of the 34 kDa/
-actinin double mutant at
15°C (27.5 hours per division) (Fig.
6B).
The ability of a calcium-insensitive form of the 34 kDa protein to function
in vivo was also tested by measurements of cell diameter. The
34-kDa/-actinin-null cells grow to reduced size in shaking culture at
20°C, attaining an average size of 9.3 µm compared with 12.8 µm for
AX2 (Rivero et al., 1999
).
Expression of wild-type 34 kDa restores wild-type cell size to 11.2 µm.
However, cells expressing the 34 kDa
EF2 protein have an average size
of only 9.8 µm (Table 1). Thus, expression of 34 kDa
EF2 fails to restore wild-type cell size and
does not supply function to the 34-kDa/
-actinin-null cells
|
Localization of the 34 kDa and F-actin
Failure of the 34 kDa EF2 protein to support proper function of the
actin cytoskeleton in 34-kDa-null and 34 kDa/
-actinin double null cells
suggests that its localization and/or dynamics in the cells may be abnormal.
For example, calcium regulation might be required for proper localization of
the 34 kDa protein to the cortex, the cell-to-cell contact regions or the
phagocytic cup. Alternatively, the dynamics of the actin cytoskeleton might be
impeded by the absence of calcium regulation. To address these questions, the
localization of the 34 kDa protein and F-actin were investigated in
34-kDa-null cells expressing wild-type 34 kDa protein, wild-type 34 kDa
protein fused to GFP or the calcium-insensitive 34 kDa
EF2 protein.
A default hypothesis to explain the localization of the 34 kDa protein with respect to F-actin is that the localization of the 34 kDa protein simply reflects the distribution of actin filaments in the cell. To test this hypothesis, a ratiometric approach was employed to assess quantitatively the distributions of F-actin and the 34 kDa protein in the leading and trailing edges of cells. The ratio of labeling of two fluorophores labeling F-actin and the 34 kDa protein in both the front and back of the cell was measured. A ratio of these two values indicates whether the relative amount of F-actin compared with the 34 kDa protein in the front of the cell differed from that in the trailing region.
To confirm the validity and absence of technical artifacts with this approach, a control experiment was performed in which the staining of F-actin labeled with both Oregon green 488 and TRITC-labeled phalloidins was measured in the front and back of the cells (Fig. 7A,B). The ratio of Oregon-green-labeled phalloidin to TRITC-labeled phalloidin in the front to that in the back was 1.04±0.15 (Table 2). This result reveals that there is no technical or instrumental artifact that will generate a ratio significantly different from 1.0 when this analysis is performed.
|
|
By contrast, comparison of the ratio of F-actin labeled with phalloidin to
the 34 kDa protein stained with monoclonal antibody B2C in the trailing region
(Fig. 7C,D) divided by that in
the leading edge is 1.38±0.38 (n=19). This result shows that
the trailing edge of the cell has a lower amount of 34 kDa protein compared
with F-actin than the leading edge. Furthermore, the ratio of F-actin to 34
kDa protein is significantly different from the F-actin to F-actin ratio
described above (P<0.01). To verify that this result is a true
reflection of the distribution of the 34 kDa protein and F-actin rather than
an artifact resulting from limited accessibility of the monoclonal antibody
used to detect the 34 kDa protein, the analysis was repeated to determine the
relative distribution of F-actin to a 34 kDa GFP construct
(Fig. 7E,F). This 34 kDa GFP
fusion protein is functional, since it can restore normal cell size to 34
kDa/-actinin double mutants (Table
1). The ratio of F-actin to 34-kDa-GFP in the trailing compared to
the leading edge of the cell is 1.19±0.15 (n=22)
(Table 2). The enrichment for
F-actin relative to the 34 kDa protein in the rear of the cell detected using
the GFP is significantly different from the F-actin to F-actin control
(P=0.005). Thus, the monoclonal antibody B2C and the 34-kDa-GFP probe
both reveal a relative decrease in the 34 kDa protein compared with F-actin in
the rear of the cell.
To determine whether calcium regulation of the 34 kDa protein was important
for this relative decrease in the presence of the 34 kDa protein in the
trailing portion of motile amoebae, the same analysis was performed in
34-kDa-null cells expressing the 34 kDa EF2 protein
(Fig. 7G,H). The results reveal
that the ratio of F-actin stained with phalloidin to the 34 kDa
EF2
protein stained with monoclonal antibody B2C in the trailing versus the
leading regions of cells was 1.09±0.29 (n=22)
(Table 2). This value was not
significantly different from the F-actin to F-actin control that showed no
difference in the front versus the trailing edge of the cell. Moreover, these
results with the 34 kDa
EF2 protein were significantly different from
those with the wild-type 34 kDa protein (P<0.05). Thus, calcium
regulation of the 34 kDa protein is required for regulation of the
distribution of the localization of the 34 kDa protein with respect to F-actin
leading to a decrease in the relative amount of the 34 kDa protein in the
trailing edge of the cell.
![]() |
Discussion |
---|
![]() ![]() ![]() ![]() ![]() ![]() ![]() |
---|
We assayed a range of cellular functions in the presence of expressed
wild-type and EF2 34 kDa proteins. When the 34 kDa
EF2 protein
is expressed in 34-kDa-null cells, it has a negative effect reducing the
growth rate at 20°C (Fig.
5A), and it fails to rescue the slow growth of these cells at
15°C (Fig. 5B). In
addition, the 34 kDa
EF2 protein fails to restore normal growth rates
to 34kDa/
-actinin double null cells at normal (20°C) and reduced
(15°C) temperatures (Fig.
6A,B), and it fails to rescue the small cell phenotype of these
cells that lack two actin crosslinking proteins
(Table 1). These results show
that calcium regulation of actin crosslinking is essential for normal 34 kDa
function and for the function of the actin cytoskeleton in vivo.
It is intriguing to ask why the 34 kDa EF2 protein exhibits a
dominant-negative effect in 34-kDa-null cells but not in the 34
kDa/
-actinin double null cells. A possible explanation emerges from
consideration of the growth rates of the strains. Wild-type cells, 34-kDa-null
and
-actinin-null all have at least one calcium-sensitive actin
crosslinking protein and grow at a normal rate at 20°C. The 34
kDa/
-actinin double null cells have no calcium-regulated
actin-crosslinking proteins and grow more slowly. Addition of the 34 kDa
EF2 protein to the 34 kDa/
-actinin double null cells does not
restore calcium-sensitive actin-crosslinking but neither does it slow growth
in this strain since the cells were already deficient in calcium-sensitive
actin crosslinking. By contrast, the 34-kDa-null cells contained one
calcium-sensitive actin crosslinking protein and grew normally. Addition of
the calcium-insensitive 34 kDa
EF2 protein to these cells may have
added additional calcium-insensitive crosslinks so that filament
rearrangements are impeded even though the cells contain a normal
-actinin molecule. Consistent with this interpretation is that the
growth rate of the 34 kDa/
-actinin double null cells is as slow as the
34-kDa-null cells expressing the 34 kDa
EF2 protein. Thus, calcium
sensitivity of actin crosslinks in the cytoskeleton is required for dynamic
rearrangements of the cytoskeleton needed for normal growth.
The ability of an actin crosslinking protein with compromised calcium
regulation to supply function in vivo was examined previously in studies of
modified -actinin (Rivero et al.,
1999
; Witke et al.,
1993
). When the second EF hand of
-actinin is modified
(
-A M2), the resultant protein requires 500 times more calcium than
wildtype for inhibition of actin binding and is not regulated by physiological
levels of calcium. Surprisingly, this
-actinin M2 restored the ability
to proceed through normal development to a cell line lacking both
-actinin and gelation factor (Witke
et al., 1993
). An
-actinin protein mutant in the first EF
hand (
-A M1) exhibits reduced actin binding and failed to restore
normal development to these cells (Witke
et al., 1993
). Expression of these same two modified
-actinin proteins in the 34 kDa/
-actinin double null cells
rescued the slow growth phenotype (Rivero
et al., 1999
). Expression of the modified
-actinin M1 and
M2 proteins resulted in a partial and complete rescue, respectively, of the
small cell size phenotype (Rivero et al.,
1999
). The
-actinin M2 protein is not sensitive to
physiological calcium levels and supplies
-actinin function to both the
34-kDa/
-actinin null and 120/
-actinin double null, in apparent
contrast to the results with the calcium-insensitive 34 kDa
EF2
protein. This difference in the results might be explained if the 34 kDa
protein has a higher affinity for F-actin than
-actinin. This is
because the rheological behavior of crosslinked actin structures is a function
of the shear rate with higher resistance to mechanical force observed at high
shear rates. At low shear rates, the crosslinked actin structures reorganize
rapidly, and the network exhibits less stiffness
(Sato et al., 1987
;
Wachsstock et al., 1993
). The
dynamic behavior of the 34 kDa protein may be slower than
-actinin, so
that in the absence of calcium-induced release from actin, an impedance to
actin filament rearrangement, is detected. If
-actinin is sufficiently
dynamic, a calcium-insensitive form may be able to supply crosslinking
function. This explanation is supported by studies of the localization of
-actinin and the 34 kDa protein in Dictyostelium. Both the 34
kDa protein and
-actinin are associated with cortical actin filaments.
However, the 34 kDa protein is tightly associated with cortical actin, whereas
-actinin reveals both cortical staining and a significant amount of
diffuse cytoplasmic localization (Brier et
al., 1983
; Fechheimer,
1987
). Further, biochemical studies in vitro show directly that
the 34 kDa protein has a higher affinity for actin filaments than does amoeba
-actinin (Fechheimer,
1987
; Lim et al.,
1999
; Wachsstock et al.,
1993
). Thus, the results can be reconciled and do demonstrate a
requirement for calcium-regulation of the actin-crosslinking in vivo.
Calcium fluxes, either from extracellular or intracellular stores, are
required for Dictyostelium spreading, locomotion and chemotaxis as
shown by chelation of cytoplasmic calcium
(Unterweger and Schlatterer,
1995). Elevation of calcium is most prominent at the rear of a
locomoting amoeba (Taylor et al., 1980;
Yumura et al., 1996
). These
results agree with a model of locomotion in which calcium elevation at the
rear of the cell promotes simultaneous contraction of actin and myosin II and
conversion of gel to sol at the rear of the cell
(DeLozanne and Spudich, 1987
;
Knecht and Loomis, 1987
;
Mast, 1926
;
Taylor and Fechheimer, 1982
).
Our results both support and extend this concept, which continues to provide a
working model of molecular events at the trailing region of a crawling cell.
First, the 34 kDa protein and actin filaments are highly colocalized in
Dictyostelium, and the only difference that can be reproducibly
detected is a relative lack of the 34 kDa protein at the rear of motile cells
(Table 2). The relative absence
of the 34 kDa protein at the rear of motile amoebae was noted in an
independent report (Okazaki and Yumura,
1995
). We found this difference using both monoclonal antibody B2C
and a 34-kDa-GFP probe (Table
2). This decrease in the 34 kDa protein at the rear of the cell
could be a result of calcium-induced release from actin. This explanation is
supported by the observation that the difference in localization of the 34 kDa
protein and F-actin at the trailing edge of moving cells is not observed in
cells expressing the calcium-insensitive 34 kDa
EF2 protein
(Table 2). Further, the
trailing edge of cells expressing 34 kDa
EF2 was sometimes extended and
contained large aggregates of actin filaments that appeared to have been
released from the posterior cortex but not yet disassembled
(Fig. 7G,H). These results
suggest that the calcium-insensitive 34 kDa
EF2 protein is not readily
disassociated from actin at the rear of the cell, remains bound to F-actin in
the tail and inhibits depolymerization of actin at the rear. This scenario is
supported by two independent observations. First, binding of the 34 kDa
protein to actin filaments slows the depolymerization of actin filaments in
vitro (Zigmond et al., 1992
).
Thus, filaments in the trailing edge with bound 34 kDa protein would be
impeded from disassembly during tail retraction. Second, measurements of
calcium ion fluxes in Dictyostelium cells reveal that calcium
elevation following application of chemoattractant correlates not with the
initial period of actin assembly but rather with the later phase of actin
depolymerization (Nebl and Fisher,
1997
). These data provide independent evidence for a role of
calcium in stimulation of actin depolymerization. Our results suggest that
calcium-induced release of 34 kDa protein from F-actin in the trailing region
of the cell may be required for depolymerization of F-actin and dynamic
rearrangement of actin structures.
Calcium-induced events in tail retraction during movement of mammalian
cells include calpain-mediated proteolysis to promote cell detachment from the
substrate (Huttenlocher et al.,
1997) and activation of myosin II
(Eddy et al., 2000
). Although
myosin II plays a prominent role in the locomotion Dictyostelium
amoebae (Fukui et al., 1990
;
Shelden and Knecht, 1996
;
Zhang et al., 2002
), the role
of calpain is worthy of additional study. Dictyostelium and mammalian
cells may differ in the manner of calcium-induced tail retraction either
because of the speed of motility, the strength of adhesion to the substrate or
other factors. Alternatively, calpain-mediated proteolysis and
calcium-sensitive actin crosslinking may be general features of locomotion
shared by diverse types of cells. This possibility is worthy of additional
investigation.
The finding that calcium-sensitive actin filament crosslinking is
significant in vivo provides additional support for a model of partial
redundancy for the function of the multiple actin crosslinking proteins
present in eukaryotic cells that has been proposed previously
(Rivero et al., 1996;
Witke et al., 1992
). The
unique functions of actin-crosslinking proteins are observed in single
mutants, whereas redundant functions are revealed by synthetic phenotypes in
double mutants (Rivero et al.,
1999
). Additional roles of actin-crosslinking proteins can be
discerned by observation of cells under physiologically relevant conditions
not normally encountered in the laboratory
(Ferrary et al., 1999
;
Ponte et al., 2000
;
Rivero et al., 1999
) and by
expression of altered forms of the proteins that lack sites for regulation by
binding of secondary messengers or covalent modification [e.g. this work and
Yamashiro et al. (Yamashiro et al.,
2001
)]. Additional insight is gained from the ability of the
altered forms to provide function to some mutants, but not others, as
discussed above. A thorough understanding of the structure and function of
actin-crosslinking proteins is beginning to emerge but will require much
additional investigation owing to the overlapping and partially redundant
functions of these ubiquitous and essential structural proteins.
![]() |
Acknowledgments |
---|
![]() |
Footnotes |
---|
![]() |
References |
---|
![]() ![]() ![]() ![]() ![]() ![]() ![]() |
---|
Andre, B., Noegel, A. A. and Schleicher, M. (1996). Dictyostelium discoideum contains a family of calmodulin-related EF-hand proteins that are developmentally regulated. FEBS Lett. 382,198 -202.[CrossRef][Medline]
Brier, J., Fechheimer, M., Swanson, J. and Taylor, D. L. (1983). Abundance, relative gelation activity, and distribution of the 95,000 dalton actin-binding protein from Dictyostelium discoideum.J. Cell Biol. 97,178 -185.[Abstract]
Clarke, M., Bazari, W. L. and Kayman, S. L. (1980). Isolation and properties of calmodulin from Dictyostelium discoideum. J. Bacteriol. 141,397 -400.[Medline]
Condeelis, J. S. and Taylor, D. L. (1977). The
contractile basis of amoeboid movement. V. The control of gelation, solation,
and contraction in extracts from Dictyostelium discoideum. J. Cell
Biol. 74,901
-927.
Cubitt, A. B., Firtel, R. A., Fischer, G., Jaffe, L. F. and
Miller, A. L. (1995). Patterns of free calcium in
multicellular stages of Dictyostelium expressing jellyfish
apoaquorin. Development
121,2291
-2301.
DeLozanne, A. and Spudich, J. A. (1987). Disruption of the Dictyostelium myosin heavy chain gene by homologous recombination. Science 236,1086 -1091.[Medline]
Dharamsi, A., Tessarolo, D., Coukell, B. and Pun, J. (2000). CBP1 associates with the Dictyostelium cytoskeleton and is important for normal cell aggregation under certain developmental conditions. Exp. Cell Res. 258,298 -309.[CrossRef][Medline]
Doring, V., Veretout, F., Albrecht, R., Muhlbauer, B.,
Schlatterer, C., Schleicher, M. and Noegel, A. A. (1995). The
in vivo role of annexin VII (synexin)-characterization of an annexin VII
deficient Dictyostelium mutant indicates an involvement in
calcium-regulated processes. J. Cell Sci.
108,2065
-2076.
Eddy, R. J., Pierini, L. M., Matsummura, F. and Maxfield, F.
R. (2000). Ca2+-dependent myosin II activation is
required for uropod retraction during neutrophil migration. J. Cell
Sci. 113,1287
-1298.
Fechheimer, M. (1987). The Dictyostelium discoideum 30,000-dalton protein is an actin filament-bundling protein that is selectively present in filopodia. J. Cell Biol. 104,1539 -1551.[Abstract]
Fechheimer, M. and Furukawa, R. (1993). A 27,000 dalton core of the Dictyostelium 34,000 dalton protein retains Ca+2-regulated actin cross-linking but lacks bundling activity. J. Cell Biol. 120,1169 -1176.[Abstract]
Fechheimer, M., Brier, J., Rockwell, M., Luna, E. J. and Taylor, D. L. (1982). A calcium and pH regulated actin binding protein from D. discoideum. Cell Motil. Cytoskeleton 2, 287-308.
Fechheimer, M. and Taylor, D. L. (1984).
Isolation and characterization of a 30,000-dalton calcium-sensitive actin
cross-linking protein from Dictyostelium discoideum. J. Biol.
Chem. 259,4514
-4520.
Fechheimer, M., Murdock, D., Carney, M. and Glover, C. V. C.
(1991). Isolation and sequencing of cDNA clones encoding the
Dictyostelium discoideum 30,000 dalton actin bundling protein.
J. Biol. Chem. 266,2883
-2889.
Fechheimer, M., Ingalls, H. M., Furukawa, R. and Luna, E. J.
(1994). Association of the Dictyostelium 30,000 dalton
actin bundling protein with contact regions. J. Cell
Sci. 107,2393
-2401.
Ferrary, E., Cohen-Tannoudji, M., Pehau-Arnaudet, G.,
Lapillonne, A., Athman, R., Ruiz, T., Boulouha, L., El Marjou, F., Doye, A.,
Fontaine, J. J. et al. (1999). In vivo, villin is required
for Ca(2+)-dependent F-actin disruption in intestinal brush borders.
J. Cell Biol. 146,819
-839.
Fukui, Y., de Lozanne, A. and Spudich, J. A. (1990). Structure and function of the cytoskeleton of a Dictyostelium myosin-defective mutant. J. Cell Biol. 110,367 -378.[Abstract]
Furukawa, R., Butz, S., Fleischmann, E. and Fechheimer, M. (1992). The Dictyostelium discoideum 30,000 dalton protein contributes to phagocytosis. Protoplasma 169, 18-27.
Furukawa, R. and Fechheimer, M. (1994). Differential localization of alpha-actinin and the 30 kD actin-bundling protein in the cleavage furrow, phagocytic cup, and contractile vacuole of Dictyostelium discoideum. Cell Motil. Cytoskeleton 29, 46-56.[Medline]
Furukawa, R. and Fechheimer, M. (1996). Role of the Dictyostelium 30 kDa protein in actin bundle formation. Biochemistry 35,7224 -7232.[CrossRef][Medline]
Hellewell, S. B. and Taylor, D. L. (1979). The contractile basis of amoeboid movement. VI. The solation contraction coupling hypothesis. J. Cell Biol. 83,633 -648.[Abstract]
Huttenlocher, A., Palecek, S. P., Lu, Q., Zhang, W., Mellgren,
R. L., Lauffenburger, D. A., Ginsberg, M. H. and Horwitz, A. F.
(1997). Regulation of cell migration by the calcium-dependent
protease calpain. J. Biol. Chem.
272,32719
-32722.
Janson, L. W., Kolega, J. and Taylor, D. L. (1991). Modulation of contraction by gelation/solation in a reconstituted motile model. J. Cell Biol. 114,1005 -1015.[Abstract]
Johns, J. A., Brock, A. M. and Pardee, J. D. (1988). Colocalization of F-actin and 34-kilodalton actin bundling protein in Dictyostelium and cultured fibroblasts. Cell Motil. Cytoskeleton 9, 205-218.[Medline]
Knecht, D. A. and Loomis, W. F. (1987). Antisense RNA inactivation of myosin heavy chain gene expression in Dictyostelium discoideum. Science 236,1081 -1086.[Medline]
Laemmli, U. K. (1970). Cleavage of structural proteins during assembly of the head of bacteriophage T4. Nature 227,680 -685.[Medline]
Levi, S., Polyakov, M. and Egelhoff, T. T. (2000). Green fluorescent protein and epitope tag fusion vectors for Dictyostelium discoideum. Plasmid 44,231 -238.[CrossRef][Medline]
Lim, R. W. and Fechheimer, M. (1997). Overexpression, purification, and characterization of recombinant Dictyostelium discoideum calcium-regulated 34,000-dalton F-actin bundling protein from Eschericha coli. Prot. Expr. Purif. 9,182 -190.[CrossRef][Medline]
Lim, R. W. L., Furukawa, R., Eagle, S., Cartwright, R. C. and Fechheimer, M. (1999). Three distinct F-actin binding sites in the Dictyostelium discoideum 34,000 dalton actin bundling protein. Biochemistry 38,800 -812.[CrossRef][Medline]
Loomis, W. F. (1971). Sensitivity of Dictyostelium discoideum to nucleic acid analogues. Exp. Cell Res. 64,484 -486.[Medline]
Lydan, M. A., Cotter, D. A. and O'Day, D. H. (1994). Calmodulin function and calmodulin-binding proteins during autoactivation and spore germination in Dictyostelium discoideum.Cell Signaling 6,751 -762.[Medline]
MacLean-Fletcher, S. D. and Pollard, T. D. (1980). Identification of a factor in conventional muscle actin preparations which inhibits actin filament self-association. Biochem. Biophys. Res. Commun. 96, 18-27.[Medline]
Maselli, A. G., Davis, R., Furukawa, R. and Fechheimer, M.
(2002). Formation of Hirano bodies in Dictyostelium and
mammalian cells induced by expression of a modified form of an actin
cross-linking protein. J. Cell Sci.
115,1939
-1952.
Mast, O. (1926). Structure, movement, locomotion, and stimulation in amoeba. J. Morph. Physiol. 41,347 -425.
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.
Okazaki, K. and Yumura, S. (1995). Differential association of three actin-bundling proteins with microfilaments in Dictyostelium amobae. Eur. J. Cell Biol. 66, 75-81.[Medline]
Ostrow, B. D., Chen, P. X. and Chisholm, R. L. (1994). Expression of a myosin regulatory light-chain phosphorylation site mutant complements the cytokinesis and developmental defects of Dictyostelium RMLC null cells. J. Cell Biol. 127,1945 -1955.[Abstract]
Ponte, E., Rivero, F., Fechheimer, M., Noegel, A. A. and Bozzaro, S. (2000). Severe developmental defects in Dictyostelium null mutants for actin-binding proteins. Mech. Dev. 91,153 -161.[CrossRef][Medline]
Putka, F. and Zeng, C. (1998). Blasticidin resistance cassette in symmetrical polylinkers for insertional inactivation of genes in Dictyostelium. Folia Biologica 44,185 -188.[Medline]
Rivero, F., Furukawa, R., Noegel, A. A. and Fechheimer, M. (1996). Dictyostelium discoideum cells lacking the 34,000 dalton actin binding protein can grow, locomote, and develop, but exhibit defects in regulation of cell structure and movement: a case of partial redundancy. J. Cell Biol. 135,965 -980.[Abstract]
Rivero, F., Furukawa, R., Noegel, A. A. and Fechheimer, M.
(1999). Dictyostelium mutants lacking the 34 kD bundling
protein and alpha-actinin or gelation factor. J. Cell
Sci. 112,2737
-2751.
Sato, M., Schwartz, W. H. and Pollard, T. D. (1987). Dependence of the mechanical properties of actin/alpha-actinin gels on deformation rate. Nature 325,828 -830.[CrossRef][Medline]
Shelden, E. and Knecht, D. A. (1996). Dictyostelium cell shape generation requires myosin II. Cell Motil. Cytoskeleton 35, 59-67.[CrossRef][Medline]
Smith, P. K., Krohn, R. I., Hermanson, G. T., Mallia, A. K., Gartner, F. H., Provenzano, M. D., Fujimoto, E. K., Goeke, N. M., Olson, B. J. and Klenk, D. C. (1985). Measurement of protein using bicinchoninic acid. Anal. Biochem. 150, 76-85.[Medline]
Spudich, J. A. and Watt, S. (1971). The
regulation of rabbit skeletal muscle contraction. J. Biol.
Chem. 246,4866
-4871.
Strynadka, N. C. J. and James, M. N. G. (1989). Crystal structures of the helix-loop-helix calcium binding proteins. Annu. Rev. Biochem. 58,951 -998.[CrossRef][Medline]
Tan, Z. and Boss, W. F. (1992). Association of phosphatidylinositol kinase, phosphatidylinositol monophosphate kinase, and diacylglycerol kinase with the cytoskeleton and F-actin fractions of carrot (Daucus carota L.) cells grown in suspension culture. Plant Physiol. 100,2116 -2120.
Taylor, D. L. and Fechheimer, M. (1982). Cytoplasmic structure and contractility: The solation-contraction coupling hypothesis. Philos. Trans. R. Soc. Lond. B. 299,185 -197.[Medline]
Taylor, D. L., Moore, P. L., Condeelis, J. S. and Allen, R. D. (1976). The mechanochemical basis of amoeboid movement I. Ionic requirements for maintaining viscoelasticity and contractility of amoeba cytoplasm. Exp. Cell Res. 101,127 -133.[Medline]
Taylor, D. L., Blinks, J. R. and Reynolds, G. (1980a). Contractile basis of ameboid movement VII. Aequorin luminescence during ameboid movement, endocytosis, and capping. J. Cell Biol. 86,599 -607.[Abstract]
Taylor, D. L., Wang, Y. L. and Heiple, J. M. (1980b). Contractile basis of amoeboid movement. VII. The distribution of fluorescently labeled actin in living amebas. J. Cell Biol. 86,590 -598.[Abstract]
Towbin, H., Staehelin, T. and Gordon, J. (1979). Electrophoretic transfer of proteins from polyacrylamide gels to nitrocellulose sheets: procedure and some applications. Proc. Natl. Acad. Sci. USA 76,4350 -4354.[Abstract]
Unterweger, N. and Schlatterer, C. (1995). Introduction of calcium buffers into the cytosol of Dictyostelium discoideum amoebae alters cell morphology and inhibits chemotaxis. Cell Calcium 17,97 -110.[Medline]
Wachsstock, D. H., Schwatz, W. H. and Pollard, T. D.
(1993). Affinity of -actinin for actin determines the
structure and mechanical properties of actin filament gels.
Biophys. J. 65,205
-214.[Abstract]
Witke, W., Schleicher, M. and Noegel, A. A. (1992). Redundancy in the microfilament system: Abnormal development of Dictyostelium cells lacking two F-actin cross-linking proteins. Cell 68,53 -62.[Medline]
Witke, W., Hofmann, A., Koppel, B., Schleicher, M. and Noegel, A. A. (1993). The Ca2+-binding domains in non-muscle type alpha-actinin: Biochemical and genetic analysis. J. Cell Biol. 121,599 -606.[Abstract]
Yamamoto, K., Pardee, J. D., Reidler, J., Stryer, L. and Spudich, J. A. (1982). Mechanism of interaction of Dictyostelium severin with actin filaments. J. Cell Biol. 95,711 -719.[Abstract]
Yamashiro, S., Chern, H., Yamakita, Y. and Matsumura, F.
(2001). Mutant caldesmon lacking cdc2 phosphorylation sites
delays M-phase entry and inhibits cytokinesis. Mol. Biol.
Cell 12,239
-250.
Yumura, S., Furuya, K. and Takeuchi, I. (1996).
Intracellular free calcium responses during chemotaxis of
Dictyostelium cells. J. Cell Sci.
109,2673
-2678.
Zhang, H., Wessels, D., Fey, P., Daniels, K., Chisholm, R. L.
and Soll, D. R. (2002). Phosphorylation of the myosin
regulatory light chain plays a role in motility and polarity during
Dictyostelium chemotaxis. J. Cell Sci.
115,1733
-1747.
Zhu, Q. and Clarke, M. (1992). Association of calmodulin and an unconventional myosin with the contractile vacuole complex of Dictyostelium discoideum. J. Cell Biol. 118,347 -358.[Abstract]
Zhu, Q., Liu, T. and Clarke, M. (1993).
Calmodulin and the contractile vacuole complex in mitotic cells of
Dictyostelium discoideum. J. Cell Sci.
104,1119
-1127.
Zigmond, S. H., Furukawa, R. and Fechheimer, M. (1992). Inhibition of actin filament depolymerization by the Dictyostelium 30,000 dalton actin bundling protein. J. Cell Biol. 119,559 -567.[Abstract]