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Cell Motility and the Cytoskeleton 40:368–378 (1998)
Disassembly of Actin Filaments Leads
to Increased Rate and Frequency of
Mitochondrial Movement Along Microtubules
Mira Krendel, George Sgourdas, and Edward M. Bonder*
Program in Cellular and Molecular Biodynamics, Department of Biological
Sciences, Rutgers University–Newark, Boyden Hall, Newark, New Jersey
In activated sea urchin coelomocytes, cytoplasmic organelles move along distinct
actin and microtubule dependent pathways, actin-based motility is driven by an
unconventional myosin, and microtubule disassembly does not effect actindependent organelle motility [D’Andrea et al., 1994: J. Cell Sci. 107:2081–2094].
Given the growing evidence for potential interactions between components of the
actin and microtubule cytoskeletons, we examined the effect of actin filament
disassembly on the movement of mitochondria along microtubules in activated
coelomocytes. Coelomocytes treated with cytochalasin B (CB), to disrupt actin
filaments, exhibited a thinning of the cytoplasm, enhanced lateral undulation of
microtubules, and ceased centripetal cortical flow of actin. Interestingly, the loss of
actin filaments resulted in a ,1.5-fold increase in the average velocity of outward
and inward moving mitochondria and increased the frequency of centripetal
movement. To test if enhanced motility along microtubules was a consequence of
decreased actin-myosin interaction, coelomocytes were treated with 2,3butanedione monoxime (BDM), a potent inhibitor of myosin activity [Cramer and
Mitchison, 1995: J. Cell Biol. 131:179–189]. BDM inhibited all types of
actin-based motility observed in these cells including retrograde cortical flow,
protrusion and retraction of the cell edge, and movement of intracellular
organelles. Surprisingly, BDM treatment stopped the movement of mitochondria in
CB-exposed cells, suggesting that BDM can also act as an inhibitor of organelle
movement along microtubules. Collectively, these data demonstrated that microtubule-dependent mitochondrial motility and microtubule movement were sensitive
to changes in the assembly state of the actin cytoskeleton. Cell Motil. Cytoskeleton
40:368–378, 1998. r 1998 Wiley-Liss, Inc.
Key words: actin filaments; motility; organelle velocity; motor proteins
INTRODUCTION
Motility of intracellular organelles, a phenomenon
observed in all eukaryotic cells, plays an important role in
many physiological processes including, but not limited
to, transport of secretory organelles, axonal transport, and
cellular polarization [Sheetz et al., 1989; Kelly, 1990;
Hirokawa, 1993]. Experimental evidence indicates that
organelles may be transported along microtubules by the
dynein and kinesin motor protein families and along actin
filaments by the myosin family of proteins [reviewed in
Langford, 1995]. Kinesins translocate organelles toward
the plus-end of microtubules while dyneins drive move-
r 1998 Wiley-Liss, Inc.
ment toward the minus-end [Kelly, 1990; Skoufias and
Scholey, 1993]. From a cellular perspective, kinesins
move cargo outward toward the periphery of the cell
*Correspondence to: Edward M. Bonder, Department of Biological
Sciences, Rutgers University–Newark, University Heights, 101 Warren
Street, Newark, NJ 07102. E-mail: ebonder@andromeda.rutgers.edu
Contract grant sponsor: American Heart Association, New Jersey
Affiliate; Contract grant sponsor: Charles and Johanna Busch Fund at
Rutgers University.
Received 30 January 1998; accepted 13 April 1998
Actin Modulates Microtubule-Based Motility
while dyneins escort cargo inward or centripetally [see
Vale, 1987; Hirokawa, 1996]. The actin-dependent motor
protein, myosin-II, has long been known to participate in
cytoplasmic streaming in plants where cytoplasmic organelles are transported along actin cables [Shimmen and
Tazawa, 1982; Kachar, 1985]. Additionally, organelle
motility in squid axoplasm [Langford et al., 1994; Bearer
et al., 1996], in neuronal growth cones [Evans and
Bridgman, 1995] and in axons and sea urchin coelomocytes depleted of microtubules [Morris and Hollenbeck,
1995; D’Andrea et al., 1994] has been linked to various
members of the unconventional myosin family. Individual organelles may also be associated with multiple
motor proteins. For example, evidence suggests the
presence of both microtubule- and actin-based motors on
the same subset of golgi-derived organelles in epithelial
cells [Fath et al., 1994] and individual axoplasmic
organelles have been documented to switch between actin
and microtubular tracks [Kuznetsov et al., 1992]. Furthermore, dynactin, the dynein activator protein complex,
may be associated with the actin binding protein spectrin
[Holleran et al., 1996], establishing a potential molecular
linkage between the actin cytoskeleton and microtubuledependent motor proteins. Consequently, intracellular
organelle motility is likely to be the result of coordinated
interplay between the two transport systems leading to
physiologically appropriate organelle motility.
Sea urchin coelomocytes contain abundant membranous organelles concentrated in the perinuclear region of
the cell that upon stimulation are induced to undergo
outward directed translocation [D’Andrea et al., 1994].
Organelles translocating along actin filaments move
slower than those moving along microtubules. Unconventional myosin classes I, VI, VII, and IX have been
identified in coelomocytes using biochemical, immunological, or molecular biological techniques [D’Andrea et
al., 1994; Sirotkin et al., 1996]. The microtubule motor
protein, kinesin, is localized to endosomes and ER in
non-activated coelomocytes [Henson et al., 1992].
Since coelomocytes possess functionally distinct
organelle transport pathways, an understanding of organelle motility in activated coelomocytes might provide
insight into coordinated regulation of microtubule- and
actin-dependent organelle transport. D’Andrea et al.
[1994] demonstrated that disruption of microtubules did
not result in a detectable effect on the velocity of
organelles moving along actin filaments. However, since
there appears to be reasonable evidence suggesting an
association between microtubule motors and the actin
cytoskeleton [Kuznetsov et al., 1992; Holleran et al.,
1996], disruption of actin cytoskeletal integrity might
have profound effects on organelles translocating along
microtubules. To investigate this possibility, two approaches were used to modify actin cytoskeletal function:
369
depolymerization of actin filaments using cytochalasin B
and inhibition of myosin motor activity using the pharmacological agent BDM. Disassembly of actin filaments
resulted in accelerated movement of mitochondria along
microtubules as well as enhanced lateral movement of
microtubules. Treatment of coelomocytes with BDM,
previously shown to specifically inhibit the ATPase
activity of myosins II and V without affecting kinesin
ATPase [Cramer and Mitchison, 1995], resulted in the
cessation of actin- and microtubule-dependent mitochondrial motility. These results suggest that the mechanisms
regulating microtubule-dependent organelle translocation
and microtubule dynamics are responsive to changes in
the actin cytoskeleton.
MATERIALS AND METHODS
Quantitative Analysis of Organelle Motility
Strongylocentrotus purpuratus coelomocytes were
plated onto coverslips either directly from coelomic fluid
or after isolation by sucrose gradient centrifugation
[D’Andrea et al., 1994]. Cells were washed with isotonic
buffer (0.5 M NaCl, 5 mM MgCl2, 20 mM HEPES, pH
7.4) and activated by perfusion of hypotonic buffer (0.3
M NaCl, 5 mM MgCl2, 20 mM HEPES, pH 7.4). BDM
(Sigma, St. Louis, MO) was used from a 200 mM stock in
water prepared on the day of experimentation and cytochalasin B (Sigma) was diluted from a 0.5 mg/ml stock in
DMSO.
Coelomocyte activation and organelle motility were
monitored using a Zeiss (Thornwood, NY) Axiophot light
microscope equipped with DIC optics and a Plan-Neo
x100 (NA 1.3) objective lens. Images were collected
using a Hamamatsu (Bellevue, WA) Newvicon camera,
frame averaged and digitally enhanced using a Quantex
image processor prior to time-lapse video documentation.
To track individual organelle movements, videotapes were played back through a Macintosh computer
and movie sequences were created using NIH Image
software (Version 1.44). X and Y coordinates of organelles in a 20 µm by 20 µm region of a cell were
determined over 120–150 sec, using NIH Image, and
plotted using Delta Graph software. The positions of
individual organelles, undergoing a single continuous
excursion, were mapped at three-sec intervals and the
resulting coordinates were used to calculate the average
velocity of moving organelles.
Fluorescence Labeling of Coelomocyte Organelles
Cells were exposed to Rhodamine-123 (1.66 µg/
ml), acridine orange (0.5 mg/ml), or neutral red (0.5
µg/ml) in either isotonic or hypotonic buffer for 10–15
min. Labeled cells were observed by DIC and epifluorescence microscopy using a Zeiss Axiophot microscope
370
Krendel et al.
coupled to a Dage SIT camera. Images were averaged and
recorded onto S-VHS video tape. To disrupt mitochondrial membrane potential, cells were exposed to 1 µM
FCCP before, during, or after labeling with fluorescent
vital dyes.
Immunofluorescent Labeling
Coelomocytes were fixed as described by Edds
[1984] and stained with fluorescent phalloidin (Molecular
Probes, Eugene, OR) to label filamentous actin. For
indirect immunofluorescent staining of microtubules,
cells were fixed with methanol at -20°C, rinsed with
phosphate-buffered saline (PBS), blocked with PBS containing 3% BSA, and incubated with DM1a monoclonal
antibodies against a-tubulin (Sigma). Primary antibody
staining was visualized using FITC-labeled goat antimouse secondary antibodies. Immunofluorescence images were collected using a Bio-Rad (Gaithersburg, MD)
MRC-1024 laser scanning confocal microscope.
RESULTS
Mitochondrial Motility in Coelomocytes
In non-activated sea urchin coelomocytes, the majority of intracellular organelles were located centrally,
surrounding the cell nucleus where they appeared to
undergo a slow churning movement (Fig. 1A) [see also
D’Andrea et al., 1994]. Cell activation, by exposure to
hypotonic buffer, resulted in rapid induction of movement
of slightly elongated and highly refractile organelles
outward into the periphery of the cell (see Table I and
Fig.1). By phase contrast microscopy, the outward moving organelles appeared phase dense. Organelle movement was saltatory, and individual organelles were capable of changing their direction of movement. Organelles
moved along linear radial tracks and in many cases
several organelles would follow the same path, as if
moving along the same microtubule or actin microfilament bundle (Fig. 1B) [D’Andrea et al., 1994]. Actin
filaments formed radially aligned bundles throughout the
lamella and dense meshworks in the perinuclear and
lamellar regions (Fig. 2A). Microtubules formed a loose
basket around the nucleus with radial microtubules
extending into the lamellae but seldom reaching out to the
edge of the cell (Fig. 2B).
To determine the identity of the outward translocating organelles, we labeled coelomocytes with a number
of cell-permeant, fluorescent vital dyes; rhodamine-123
to stain mitochondria and acridine orange and neutral red
to label acidic compartments. Rhodamine 123 labeled
most of the peripherally translocating organelles in cells
activated with hypotonic buffer (see Fig. 1D), whereas
acridine orange and neutral red stained only perinuclear
organelles (data not shown). Comparisons of DIC and
fluorescence images established that most of the organelles labeled with rhodamine 123 were small, slightly
elongated, and highly refractile by DIC and phase dense
by phase contrast microscopy. By contrast, perinuclear
acidic vesicles were often large, round, and phase-lucent
(data not shown). To verify the specificity of rhodamine
123 labeling, coelomocytes were treated with carbonyl
cyanide p-(trifluoromethoxy)phenylhydrazone (FCCP),
an uncoupler of mitochondrial oxidative phosphorylation,
which disrupts the membrane potential and eliminates
uptake of rhodamine 123. In all cases, cells exposed to
FCCP prior to, during, or after exposure to rhodamine
123 did not show uptake of dye into the translocating
organelles. Staining of the large, round perinuclear organelles with either acridine orange or neutral red was not
effected by exposure to FCCP. These observations establish that one hallmark of coelomocyte activation is the
rapid induction of mitochondrial motility.
Effect of Cytochalasin B Treatment
on Mitochondrial Motility
Cytochalasin B is commonly used in studies of
actin-dependent motility since it disrupts the actin cytoskeleton in vivo, presumably by slowing plus-end assembly and destabilizing filaments [Bonder and Mooseker,
1986; Sampath and Pollard, 1991]. Coelomocytes exposed to CB in isotonic buffer showed a thickening of the
outer edge of the lamellae, a loss of centripetal cortical
flow, inhibition of protrusion/retraction activity of the
lamellae, ‘‘thinning’’ of the cytoplasm, and apparent
brownian motion of perinuclear organelles (data not
shown) [see also Edds, 1993]. In agreement with Edds
[1993], CB resulted in mass depolymerization of actin
filament bundles and meshworks while leaving behind a
rim of residual actin around the edge of the cell and
occasional cytoplasmic actin aggregates (Fig 2C).
When coelomocytes were first treated with hypotonic buffer to induce outward translocation of mitochondria and then exposed to CB, the initial response was a
‘‘snapping’’ back of the cytoplasm along with many, but
not all, mitochondria into the perinuclear region, as well
as inhibition of cortical flow and protrusive activity.
Mitochondria that remained in the lamellae continued to
move along linear tracks (Fig. 3) of radially aligned
individual microtubules or bundles of microtubules present within the lamellar cytoplasm that was depleted of
actin filaments (Figs. 2D and 3). All the effects of CB
were fully reversible upon incubation in hypotonic buffer
lacking CB.
Treatment with CB resulted in several noteworthy
qualitative effects on mitochondrial motility, microtubule
organization, and microtubule motility. In control cells,
we never observed mitochondria that completely trans-
Actin Modulates Microtubule-Based Motility
371
Fig. 1. Identification of mitochondrial motility in coelomocytes. A: A
video-DIC micrograph of a single coelomocyte showing the presence
of numerous organelles in the perinuclear region of the cell. B: The
same cells as shown in A after treatment with hypotonic buffer.
Numerous refractile organelles migrated outward into the broad, thin
lamellar region, often along straight linear tracks. C: Video-DIC
micrograph of a coelomocyte after treatment with hypotonic buffer
containing the mitochondria specific vital dye, rhodamine-123. Numerous organelles have migrated out into the periphery of the cell; note the
two discrete groups of refractile organelles (arrows). D: Fluorescence
micrograph of the cell presented in C. Rhodamine-123 labeling is well
correlated with the motile refractile organelles observed using DIC
microscopy, in particular compare the organelles identified by arrows,
see C and D.
located out to the very edge of the cell whereas in
CB-treated cells mitochondria often traveled to the edge
of the lamellae where they appeared ‘‘stranded’’ or
moved tangentially along the edge of the cell on microtubule tracks visible by DIC. Additionally, mitochondria
appeared to move inward with greater frequency and
movement along microtubules appeared faster (see be-
low). In the absence of an intact actin cytoskeleton, the
microtubule network tended to collapse forming parallel
bundles or loose ‘‘knots’’ of microtubules (see Figs. 2D
and 3). Interestingly, individual microtubules or bundles
of microtubules started to undergo wild lateral undulations through the thinned lamellar cytoplasm (Fig. 3, see
arrows).
372
Krendel et al.
TABLE I. Effect of CB on Coelomocyte Mitochondrial Translocation
Control
Average velocity (µm/sec 6 SEM)
Fraction of time spent moving (%)
CB
Inward
Outward
Inward
Outward
0.098 6 0.005
7.2
0.102 6 0.003
22.5
0.145 6 0.005
17.9
0.149 6 0.004
29.5
Quantitative Analysis of Mitochondrial Motility
Effect of BDM on Mitochondrial Motility
Individual mitochondria present within 20 by 20 µm
square sectors of lamellae were tracked over a 2-min time
span to determine organelle velocity and relative frequency of inward and outward movements. The position
of each mitochondrion was mapped at 3-sec intervals and
the average velocities were calculated for those mitochondria that changed position over the 3 sec. Previously,
D’Andrea et al. [1994] identified mitochondria undergoing a continuous excursion and used the information
gathered to calculate average velocity. The two methods
for determining average velocity of mitochondria resulted
in essentially identical measurements (data not shown).
An added advantage of tracking individual moving and
stationary mitochondria within the 20 by 20 µm area is
the ability to determine both the frequency of inward
versus outward movement and the percentage of time
mitochondria spent moving.
The velocities observed in control, activated coelomocytes ranged from 0.01 µm/sec to 0.48 µm/sec, with an
average outward velocity of 0.102 µm/sec and average
inward velocity of 0.098 µm/sec (Table I). Individual
mitochondria were stationary approximately 70% of the
time and the frequency for outward movement was 3
times higher than for inward movement (Table I). Average velocity of both outward and inward organelle
movement increased approximately 1.5-fold in cells
treated with CB (Table I). This 1.5-fold increase in
average velocity was a characteristic feature of CB
treatment as reflected in individual cell-by-cell comparisons before and after exposure to CB (Fig. 4). Comparison of the velocities in control vs. CB-treated coelomocytes by an unpaired t-test showed that the calculated
difference was statistically significant with a 99.9%
confidence level. In addition to increased velocity, mitochondria in CB-treated cells spent a greater proportion of
time moving as compared to control cells (50 vs. 30%
respectively; see Table I). Further, greater than 70% of the
organelles in CB-treated cells moved at rates of 0.1
µm/sec or higher whereas in control cells less than 50% of
the organelles moved with velocities of 0.1 mm/sec or
higher. Another striking change was the doubling in
frequency of inward motility following disassembly of
the actin cytoskeleton (17.9% in CB-treated vs. 7.2% in
control cells; see Table I).
The observed increase in mitochondrial motility
after CB treatment could have resulted from the loss of
the actin filament network and/or the consequent loss of
functional actin-myosin interaction. In each case, the
presence of an intact actin cytoskeleton contributes a
‘‘braking’’ effect that would impede the progress of
organelles moving along microtubules. To examine if
myosin motor activity could influence microtubule-based
organelle transport, coelomocytes were treated with BDM,
previously shown to be a specific inhibitor of myosin-II
and V activity [Cramer and Mitchison, 1995]. Treatment
of non-activated cells with 5–40 mM BDM resulted in a
dose-dependent attenuation of cortical flow and protrusion/
retraction of lamellae. In the presence of 20 or 40 mM
BDM, cortical flow and lamellar activity were completely
stopped without any readily detectable destructive changes
in overall cell morphology, presence of actin filaments, or
presence of microtubules (see Fig. 2). One qualitative
effect of BDM was that actin filaments and microtubules
appeared ‘‘collapsed,’’ giving the impression that more
bundles were present in treated cells vs. non-treated cells
(compare Fig. 2E, F with Fig. 2A, B).
When coelomocytes were first treated with isotonic
buffer containing 20–40 mM BDM and then activated by
hypotonic buffer containing BDM, there was little or no
mitochondrial movement observed (data not shown); the
inhibitory effect was fully reversible upon incubation of
cells with hypotonic buffer lacking BDM. To assess
whether the observed effect of BDM on mitochondrial
dispersion was due to an effect on coelomocyte activation, coelomocytes were first treated with hypotonic
buffer to initiate mitochondrial translocation and then
exposed to hypotonic buffer containing BDM. Mitochondria that were translocating after treatment with hypotonic buffer would stop moving after 3–5 min of exposure
to hypotonic buffer containing BDM and mitochondria
resumed translocation after the preparations were washed
free of BDM (Fig. 5). Interestingly, mitochondria that
ceased moving after BDM treatment appeared to be
loosely tethered to the location where they stopped (Fig.
5B). Occasionally, some mitochondria would undergo
short excursions in either the outward or inward direction
that were slower and lasted for shorter time periods than
typically observed in non-treated cells (Fig. 5).
Actin Modulates Microtubule-Based Motility
373
Fig. 2. Effect of CB and BDM on the organization of actin filaments
and microtubules. A & B: Laser scanning confocal micrographs of
individual, non-activated coelomocytes labeled with rhodaminephalloidin to detect actin filaments (A) and by indirect immunofluorescence to detect microtubules (B). C & D: Laser scanning confocal
micrographs of coelomocytes stained for actin filaments (C) and
microtubules (D) after treatment with isotonic buffer containing 1
µg/ml of CB. Exposure to CB resulted in depolymerization of the
majority of actin filaments, leaving behind a narrow, peripheral ridge of
phallodin stained actin (Panel C). CB treatment led to the rearrange-
ment of microtubules into a ‘‘collapsed’’ network consisting of large
bundles of microtubules (compare panels B and D). E and F: Laser
scanning confocal micrographs of coelomocytes stained for actin (E)
and microtubules (F) after treatment with isotonic buffer containing
BDM. There appears to be no detrimental effect on the quantity of actin
filaments or microtubules after exposure to BDM. However, qualitatively, treatment with BDM appeared to have an effect on the
organization of microtubules since the microtubules in treated cells
tended to form bundles. Bar - 10 µm.
Effect of BDM on Mitochondrial Motility in CB
Treated Coelomocytes
ity could be a consequence of mitochondria being ‘‘stuck’’
to an actin filament by a poisoned myosin consequently
preventing movement along microtubules. If inhibition of
microtubule motility is a secondary effect of BDM,
Since BDM is reported to be a specific inhibitor of
myosin function, the observed lack of microtubule motil-
374
Krendel et al.
Fig. 3. Mitochondrial and microtubule motility in CB-treated cells. Four frames from a two-minute
time-lapse video sequence of a single cell after exposure to CB. In this sequence, outward and inward
movement of mitochondria (arrowheads) as well as lateral undulation of a microtubule (see arrow) can be
detailed. Bar - 10 µm.
disruption of the actin-myosin interaction should release
the mitochondria and allow movement along a microtubule until another poisoned actin-myosin interaction
forms. This attachment and release could account for the
occasional, short excursions that mitochondria take in the
presence of BDM. To investigate this possibility, coelomocytes were activated using hypotonic buffer containing
CB, followed by exposure to both CB and BDM. Prior to
BDM treatment, mitochondria were observed moving
along microtubular tracks and movement was ceased
within 5 min after exposure to BDM (Fig. 6) and the
majority of mitochondria remained stationary. Some
mitochondria occasionally performed short outward and
inward excursions along microtubular tracks but they
never translocated for distances comparable to those
measured in non-BDM-treated cells (Fig. 6). After washout of BDM using hypotonic buffer containing CB,
mitochondrial motility rapidly returned to the levels
observed prior to BDM treatment (Fig. 6).
DISCUSSION
Intracellular organelle transport is driven by distinct
families of motor proteins, the myosins that move cargo
Actin Modulates Microtubule-Based Motility
375
that the dynamic properties, distribution, and length of
microtubules are modulated by the actin cytoskeleton.
Furthermore, quantitative analysis of live cells established that the direction and velocity of mitochondria
moving along microtubules are linked to the assembly
state of cytoplasmic actin filaments. Thus, our observations indicate that microtubule dynamics and microtubuledependent organelle motility are effected by the surrounding actin cytoskeleton.
Actin Cytoskeleton Moderates the Velocity
of Mitochondria Moving Along Microtubules
Fig. 4. Quantitive analysis of mitochondrial motility in coelomocytes.
Three individual cells were analyzed for outward and inward mitochondrial velocities prior to treatment with CB (j, h) and after treatment
with CB (e, B). In each case, treatment with CB resulted in an ,50%
increase in the average mitochondrial velocity. Error bars represent
SEM.
along actin filaments and the kinesins and dyneins that
carry cargo along microtubules [Skoufias and Scholey,
1993; Fath and Burgess, 1994; Langford, 1995]. While
the two transport systems consist of distinct components
performing different functions, they must be closely
linked in order to provide coordinated organelle transport.
The existence of a coordinated interaction is plausible
given that organelles can possess both actin- and microtubule-dependent motors [Fath et al., 1994], can switch
between actin and microtubule tracks [Kuznetsov et al.,
1992], and the actin binding protein spectrin is associated
with dynactin, the dynein activator complex [Holleran et
al., 1996].
Previously, we demonstrated that coelomocytes
possess both actin- and microtubule-dependent organelle
motility that could be distinguished by the velocity of
organelle translocation [D’Andrea et al., 1994]. Fast
organelle motility in coelomocytes occurs primarily along
microtubules with an average velocity of ,0.1 µm/sec
that can be eliminated by disassembling microtubules
with nocodazole. Slow organelle motility is likely to be
driven by an unconventional myosin [D’Andrea et al.,
1994; Sirotkin et al., 1996] having a velocity less than
0.05 µm per sec. Depolymerization of microtubules has
no detectable effect on the velocity of organelles moving
along actin filaments [D’Andrea et al., 1994]. In the
present study, we used organelle motility in sea urchin
coelomocytes to investigate the consequences of actin
cytoskeletal integrity and myosin motor protein activity
on organelle translocation along microtubules. We report
Correlative analysis of fluorescence and DIC images of rhodamine 123 labeled cells led to the conclusion
that the small, oblong, highly refractile organelles imaged
by video-DIC were mitochondria and that all or almost all
motile organelles in activated coelomocytes were mitochondria. These data provide documentation that, similar
to axonal transport of mitochondria [Morris and Hollenbeck, 1995], coelomocyte mitochondria are transported
along both actin filaments and microtubules. Previously,
Henson and co-authors [1992], using anti-sea urchin egg
kinesin antibodies, demonstrated that some populations
of coelomocyte organelles, but not mitochondria, are
associated with the microtubule-dependent motor protein
kinesin. Lee and Hollenbeck [1995] found that in neurons
increased kinesin phosphorylation promoted kinesin association with membranes and correlated with an increase
in axonal transport of organelles. Thus, if coelomocyte
activation, in addition to being calcium dependent [Henson and Schatten, 1983; Hyatt et al., 1984], was also
accompanied by an increase in kinase activity there could
result elevated kinesin phosphorylation leading to mitochondrial association and motility. In support of this idea
are our unpublished observations that stimulation of
protein kinase C by the phorbol ester, TPA, can induce
organelle motility in resting coelomocytes.
Interestingly, when the actin filament network in
coelomocytes was disrupted with CB the average inward
and outward velocity of mitochondria moving along
microtubules increased. Previously, Morris and Hollenbeck [1995] found that the average velocity of mitochondria moving in axons increased after treatment with
cytochalasin. Can the observed increase in average mitochondrial velocity after CB treatment be explained by a
simple ‘‘statistical effect’’ due to the loss of slow,
actin-dependent movement? Elimination of the slow
moving contingent of mitochondria would result in a
statistically elevated average velocity, without any increase in the actual rate of translocation of organelles
moving along microtubules. In coelomocytes, unlike
axons, it is possible to distinguish between the velocities
for actin-dependent and microtubule-dependent mitochondrial movement. With this information it is possible to
eliminate slow, 0.05 µm/sec or less, actin-dependent
376
Krendel et al.
Fig. 5. Analysis of mitochondrial movements after BDM treatment.
Trajectories of individual organelles were traced in a 20-by-20 µm area
within a single lamella after activation with hypotonic buffer (Panel
A), after incubation in hypotonic buffer containing 20 mM BDM
(Panel B), and after wash-out of BDM with hypotonic buffer (Panel
C). Trajectories were generated by mapping the position of 10-15
mitochondria every 3 seconds over 120 seconds using NIH Image
software. Treatment with BDM resulted in a dramatic decrease in the
movement of mitochondria and in many cases the mitochondria
appeared to remain essentially stationary. Removal of BDM restored
typical mitochondria motility (Panel C).
Fig. 6. Analysis of mitochondrial movement after treatment with
BDM and CB. Coelomocytes were initally activated with hypotonic
buffer to induce organelle motility followed by subsequent treatment
with hypotonic buffer containing CB (Panel A), hypotonic buffer
containing CB and BDM (Panel B), and finally hypotonic buffer with
CB (Panel C). Trajectories were generated as described in Figure 5.
Interestingly, treatment with BDM greatly slowed or completely
inhibited the movement of mitochondria along microtubules (Panel B).
The inhibitory effect of BDM on microtubule-based motor activity was
fully reversable upon incubation with hypotonic buffer lacking BDM
(Panel C).
movements from the calculations to obtain a corrected
microtubule driven velocity. Consequently, if the observed increase in the calculated average velocity is a
statistical effect, the corrected velocity before CB treatment should equal the velocity measured after CB
treatment. Using this analysis, as predicted from the data
in D’Andrea et al. [1994], mitochondria moved along
microtubules with a velocity of ,0.1 µm/sec in control
cells. Interestingly, disruption of actin filaments resulted
in an ,35% increase in the average velocity for both
outward and inward mitochondrial movement on microtubules. The calculated difference in velocities was statistically significant (99.9% confidence level) as judged by an
unpaired t-test indicating that mitochondria do indeed
move faster along microtubules after actin filaments were
disassembled in the lamellae of coelomocytes. Whether
the newly observed acceleration of organelles moving
along microtubules occurs in other systems (e.g., axons)
after disruption of actin filaments remains unknown.
One possible explanation for the observed increase
in mitochondrial velocity is that actin filaments provide
mechanical resistance countering the propulsive forces of
the microtubule motor proteins present on the organelle
surface. In vitro measurements of viscoelasticity suggest
that actin filaments have a twofold higher viscosity than
equivalent solutions of nonfilamentous actin [Sato et al.,
1985]. Since, actin filament depolymerization by CB
lowers viscosity [MacLean-Fletcher and Pollard, 1980] it
Actin Modulates Microtubule-Based Motility
is reasonable to predict that the viscosity of the coelomocyte cytoplasm decreased as a consequence of CB
treatment. If treatment with CB resulted in a 2-fold
decrease in the viscosity of the cytoplasm, then based on
Stokes’ Law, there should be an acccompanying 2-fold
increase in organelle velocity. Interestingly, the measured
,1.5-fold increase in outward and inward mitochondrial
velocity is consistent with the above viscosity-based
effect of actin filaments on organelles moving along
microtubules.
Alternatively, since mitochondria in activated coelomocytes may be associated both with actin- and microtubule-dependent motors, binding interactions between
myosin-type motors and actin filaments may slow down
movements along microtubules. In this case, CB treatment eliminates myosin-actin filament interactions and
the observed velocities are solely dependent upon microtubule motor protein activity. In order to test such a
‘‘myosin-braking’’ model, cells were treated with BDM
to inhibit myosin motor activity. BDM, a specific inhibitor of myosins, has been used in studies of muscle
contraction [Higuchi and Takemori, 1989; McKillop et
al., 1994; Siegman et al., 1994], postmitotic cell spreading [Cramer and Mitchison, 1995], formation of focal
contacts [Chrzanowska-Wodnicka and Burridge, 1996],
and neuronal growth cone motility [Lin et al., 1996].
Similar to other cells [Lin et al., 1996; Waterman-Storer
and Salmon, 1997], BDM inhibited myosin-driven cortical flow in coelomocytes; however, unexpectedly, BDM
inhibited all organelle motility in coelomocytes including
organelle motility occurring along microtubules.
To date, BDM is proposed to directly inhibit the
myosin II and V ATPase [Herrmann et al., 1992; McKillop et al., 1994; Cramer and Mitchison, 1995]. While
typically used as a myosin specific inhibitor, BDM can
potentially effect other cellular activities including dephosphorylation of acetylcholinesterase [Hobbiger, 1963], ion
channel conductance [Phillips and Altschuld, 1996],
and/or reduce the level of myosin II regulatory light chain
phosphorylation [Siegman et al., 1994; ChrzanowskaWodnicka and Burridge, 1996]. Consequently, the effects
of BDM may be less specific than originally thought and
results obtained in in vivo experiments utilizing BDM
should be interpreted cautiously.
In addition to accelerated velocities, mitochondria
spent greater periods of moving after CB-treatment. This
effect was especially pronounced for inward mitochondrial movement where there was a greater than twofold
increase in the frequency of movement. While it is
tempting to speculate on the potential mechanism of this
observation, the absence of available information on the
detailed polarity of microtubules and the presence of
dynein motor proteins preclude such discussion.
377
Organization of Microtubules Is Modulated
by Actin Filaments
In addition to the observed effect on mitochondrial
velocities, depolymerization of actin filaments altered the
intracellular distribution of microtubules and microtubuleassociated organelles. After cytochalasin treatment, it
appeared that the microtubules formed tangled bundles
that often extended out to the edge of the cell. In the
presence of cytochalasin, many mitochondria accumulated at the edge of the cell while in control cells the
outermost regions of the lamellae did not contain mitochondria. Strikingly, the time-lapse video images documented the presence of wild, lateral undulations of
microtubules or microtubule bundles giving an impression that these movements were actively driven. These
observations suggest that actin filaments, by forming a
cytoplasmic matrix, can limit lateral motility, bundle
formation, and plus-end growth of microtubules at the
cell periphery. Our findings complement and extend the
recent report of Waterman-Storer and Salmon [1997] who
elegantly demonstrated that actin-dependent cortical flow
profoundly effects microtubule bending, elongation, and
breakage and support the proposal by Edds [1984] that
actin filaments may limit microtubule distribution in
coelomocytes.
Our findings show that mitochondrial transport and
redistribution in activated coelomocytes are a result of
coordinated activity of actin- and microtubule-based
transport systems. Interestingly, the structural integrity of
the actin cytoskeleton exerted profound effects on the
motility characteristics of mitochondria and microtubules. This result raises the possibility that localized
assembly/disassembly of actin filaments may participate
in spatial regulation of microtubule dynamics, microtubule-dependent motor activity, and organelle positioning.
ACKNOWLEDGMENTS
We are grateful to Rob Harrell, Volodia Sirotkin,
and Tim Mure for their help and assistance during the
course of experimentation and manuscript preparation.
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