Department of Biology, Washington University, One Brookings Drive, St Louis, MO 63130, USA
* Author for correspondence (e-mail: miller{at}biology.wustl.edu)
Accepted 4 September 2002
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Summary |
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Key words: Myosin VI, Actin dynamics, Cortactin, Arp2/3, Dynamin
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Introduction |
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Some clues as to the cellular functions of myosin VI have come from
phenotypic analysis of mutations in Mus musculus, Caenorhabditis
elegans and Drosophila melanogaster. The defects seen in the
deaf Snell's Waltzer mutant mouse are caused by a loss-of-function mutation in
the myosin VI gene. These mutants display disruptions in the
organization and morphogenesis of the stereocilia in the inner ear
(Avraham et al., 1995;
Self et al., 1999
). The hair
cell stereocilia in these mutants are disorganized, and the membrane between
neighboring stereocilia is uprooted, suggesting a defect in the anchoring of
stereocilia and the apical membrane (Self
et al., 1999
). In C. elegans, loss-of-function mutations
in one of two isoforms of myosin VI cause male sterility
(Kelleher et al., 2000
). These
mutants display defects in the asymmetric sorting of mitochondria and
ER/Golgi-derived organelles during spermatogenesis, suggesting defects in
organelle trafficking or localization. Consistent with a role in trafficking,
myosin VI in vertebrate tissue culture cells localizes to trafficking
membranes such as the trans-Golgi network
(Buss et al., 1998
) and
clathrin-coated pits and vesicles (Buss et
al., 2001a
), and overexpression of the tail fragment of myosin VI
reduces transferrin uptake (Buss et al.,
2001a
). From these studies and those in Drosophila
(described below) it has been proposed that myosin VI has a role in organizing
and/or trafficking membrane. However, the details of how myosin VI
participates in these functions remain unclear.
In Drosophila there is one myosin VI gene. The proposed
function for myosin IV is in membrane organization and trafficking. In
Drosophila syncytial blastoderm embryos, myosin VI protein localizes
to cortical actin in transient mitotic membrane invaginations (pseudocleavage
furrows). Inhibition of myosin VI function in the Drosophila embryo
causes defects in formation of these transient mitotic membrane invaginations
(Mermall and Miller, 1995).
Thus, myosin VI is required for membrane remodeling during embryogenesis.
Myosin VI function is also required for membrane remodeling during the
individualization step of spermatogenesis
(Hicks et al., 1999). During
individualization, a syncytial membrane encasing a bundle of 64 spermatids is
remodeled so that an individual membrane encases each of the 64 mature sperm.
Myosin VI localizes to an actin complex, the individualization complex, which
assembles at the spermatid heads at the start of individualization
(Hicks et al., 1999
). This
complex progresses from the spermatid heads to the tips of tails, remodeling
membrane as it moves. Loss of myosin VI in the testis leads to the disruption
of these actin complexes as they progress and, consequently, individualization
is not completed (Hicks et al.,
1999
).
The precise function of myosin VI in individualization remains unclear. Does myosin VI catalyze transport of membrane trafficking components at the actin individualization complex? Is it involved in anchoring membrane to actin and/or organizing actin polymerization sites at the membrane? To obtain a clearer understanding of myosin VI's role in the individualization complex, we compared the localization of myosin VI in spermatids with that of proteins known to have roles in regulating actin dynamics or membrane trafficking. We also tested for genetic interactions between mutations in myosin VI and mutations in the membrane trafficking gene, dynamin. We report that myosin VI localizes with regulators of actin dynamics and is required for the localization of these proteins. We also show that myosin VI mutations interact genetically with dynamin mutations to affect actin dynamics. We propose that myosin VI is important for actin assembly at sites of membrane remodeling.
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Materials and Methods |
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Temperature shifts to inactivate shibire function
Newly eclosed adult flies (0-12 hrs after eclosure) were shifted to the
non-permissive temperature of 30°C in a water bath for the times indicated
in each experiment. At the end of the temperature shift, flies were
immediately dissected in prewarmed (30°C) dissection buffer on prewarmed
dissection slides. A temperature-controlled rubber mat (controller and heating
mat from Thermolyne) set at low temperature was placed under the dissection
slide to maintain the non-permissive temperature during dissection. Testes
dissected in this manner were immediately fixed as described below.
Testis dissection and fixation
Newly eclosed males (3-12 hrs after eclosure) were collected and incubated
on ice until paralyzed (except with shi1 flies, in which
case they were already paralyzed by the non-permissive temperature). Testes
were dissected in testis isolation buffer (47 mM NaCl, 183 mM KCl, 10 mM Tris
pH 6.8) and prepared by a method previously described
(Hime et al., 1996) with minor
modifications. Briefly, testes were squashed, incubated in cold 100% ethanol
for 10 minutes, incubated in 4% paraformaldehyde for 7 minutes, incubated once
in PBT (PBS + 0.3% Triton X-100 + 0.3% sodium deoxycholate) for 15 minutes and
washed twice in PBST (PBS + 0.1% Triton) for 10 minutes.
Antibodies and fluorescent markers
Monoclonal anti-Drosophila myosin VI antibodies, 3c7 (1:20), and
polyclonal anti-Drosophila myosin VI antibodies (1:100) were
previously described (Kellerman and
Miller, 1992). The mouse anti-rat dynamin-1 monoclonal antibody
(1:1000) was obtained from Transduction Laboratories (Lexington, KY, catalog
number D25520). The rabbit anti-Drosophila dynamin-1 (shibire)
polyclonal antibody (1:500) was a gift from Mani Ramaswami
(Estes et al., 1996
). Both
antibodies to dynamin-1 show the identical localization in the testis. The rat
anti-Drosophila cortactin antibody (1:250) was a gift from Manabu
Takahisa (Katsube et al.,
1998
). The rabbit anti-Drosophila Arp3 (1:50) was a gift
from Bill Theurkauf and the rabbit anti-human ARPC2/p34 (1:100) was a gift
from Matt Welch (Welch et al.,
1997
). The rabbit anti-Drosophila amphiphysin (1:100) and
anti-Drosophila alpha adaptin (1:25) were gifts from Andrew Zelhof
(Zelhof et al., 2001
) and Nick
Gay (Dornan et al., 1997
;
Zelhof et al., 2001
). The
rabbit and rat anti-Drosophila capping protein ß (CP-ß)
antibodies (1:10) were described previously
(Hopmann et al., 1996
).
Immunofluorescence staining and imaging
Fixed testes were blocked in PBST + 3% BSA for at least 30 minutes at room
temperature or overnight at 4°C. All antibodies and dyes were diluted in
PBST + 3% BSA. Antibodies were incubated with tissue samples for 2 hours at
room temperature or overnight at 4°C. After incubation with primary
antibodies in a humid chamber, fixed testes were washed four times in PBST +
3% BSA for 10 min at room temperature. Then, secondary antibodies were
incubated for 2 hours in a humid chamber. Goat anti-mouse Alexa Fluor 488 and
goat anti-rabbit Alexa Fluor 488 secondary antibodies were diluted 1:500. Goat
anti-mouse Alexa Fluor 568 and goat anti-rabbit Alexa Fluor 568 secondaries
were diluted 1:1000. Alexa Fluor 488 phalloidin (1:125) and the DNA dyes
TOTO-3 (1:3000) and DAPI (1 µg/ml) were included in incubations with
secondary antibodies for 2 hours at room temperature. All Alexa-conjugated
secondary antibodies, phalloidin and TOTO-3 dyes were from Molecular Probes.
After staining, testes were washed four times in containers filled with PBST
for 10 minutes at room temperature and finally mounted in mounting media [500
mg/ml glycerol, 17 mM Tris pH 8.5, 200 mg/ml Mowiole (Calbiochem)]
All imaging of fluorescent stains was performed on a Leica laser scanning spectral confocal microscope (model TCS SP2). Images shown are either single planes or, when noted, multiple planes collapsed into a single images (a projection). Projections were obtained by collecting consecutive planes at 0.5 µm intervals in volume samples that include most of the actin individualization complex.
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Results |
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The site at which myosin VI concentrates is the junction between a moving
actin structure and a zone of active membrane remodeling. This location places
myosin VI in an ideal position to link sites of remodeling to actin dynamics.
Therefore, we examined the localization of proteins that have been implicated
in membrane/actin coordination. One such protein is the actin-binding protein
cortactin. Cortactin is thought to link membrane signaling proteins to actin
dynamics by virtue of is ability to associate with both actin polymerization
components and membrane-associated kinases
(Olazabal and Machesky, 2001;
Weed et al., 2000
).
We examined the distribution of cortactin in individualizing spermatids with anti-Drosophila cortactin antibodies. Like myosin VI, cortactin concentrated at the front of actin cones (Fig. 2B), and double labeling of spermatids with cortactin and myosin VI antibodies showed that they colocalized at the front of each actin cone (Fig. 2M,N). Cortactin was also present on the cyst membrane (Fig. 3B). The distribution of cortactin in individualization complexes indicates that the fronts of the actin cones are sites where actin polymerization might be coupled with membrane dynamics. Myosin VI colocalization with cortactin at these sites suggests myosin VI may also be involved in these dynamics.
|
|
To further demonstrate that the fronts of the actin cones are sites of
regulated actin assembly, we examined the distribution of the arp2/3 complex
and capping protein in individualizing spermatids. The arp2/3 complex is a
complex of seven proteins that binds actin filaments and nucleates new actin
filament assembly (Cooper et al.,
2001; May, 2001
).
Capping protein is a barbed-end actin-binding protein with a known role in
regulating actin polymerization at sites where the arp2/3 complex promotes
assembly (Cooper and Schafer,
2000
; Schafer and Cooper,
1995
). It is also concentrated in regions of dynamic actin
assembly in many cell types (Schafer et
al., 1998
; Waddle et al.,
1996
). In individualizing spermatids, the arp2/3 complex, as
demonstrated by arp3 (Fig. 2E) and ARPC2/p34 (Fig. 2H)
staining, and capping protein, as demonstrated by CP-ß staining
(Fig. 2K), concentrated at the
front of actin cones. In both cases, staining was also visible generally
through the cytoplasm of the cyst and along the actin cones
(Fig. 2H,K). Double labeling
experiments showed that myosin VI colocalized with concentrated arp3
(Fig. 2P,Q) and capping protein
(Fig. 2S,T) at the front of
actin cones. The accumulation of proteins involved in actin polymerization at
the front of the actin cones supports the idea that the zone where myosin VI
concentrates is a zone of active actin assembly.
Myosin VI is required for the proper distribution of cortactin and
the arp2/3 complex on individualization complexes
The colocalization of cortactin, arp2/3 complex and myosin VI on
individualization complexes prompted us to examine cortactin and arp2/3
complex distribution on individualization complexes in myosin VI
mutants. Cortactin could be detected on actin individualization complexes in
myosin VI mutants (jar1). However, its
distribution was not normal. Cortactin was not concentrated at the front of
the actin cones (Fig. 3E,Q).
Instead, it was weakly present uniformly along the cones. The complexes shown
have a disrupted morphology and reduced actin staining, as is typically
observed for progressed actin cones in myosin VI mutants. When early
individualization complexes were examined in myosin VI mutants, no
early complexes showed any concentration of cortactin at the front of cones
(data not shown). By contrast, in wild-type spermatids, some early
individualization complexes had cortactin concentrated at the front and others
did not. When doubly stained for myosin VI, those complexes with concentrated
myosin VI also showed concentrated cortactin. We conclude that myosin VI is
required for the proper asymmetrical distribution of cortactin on actin
cones.
In contrast to the localization of cortactin on actin cones, its localization to cyst membrane was unaffected in myosin VI mutants, indicating that myosin VI is not required for its proper localization to cyst membrane (Fig. 3B).
We also observed defects in arp2/3 complex localization in myosin VI mutants. Arp3 did not concentrate at the front of actin cones, either early (data not shown) or on progressed complexes (Fig. 3K), in myosin VI mutants. In addition, there appeared to be a higher level of arp3 staining in the cytoplasm of the cysts in myosin VI mutants in comparison to wild-type cysts (Fig. 3K,H). This may be because arp3 cannot concentrate on the actin cones and, instead, accumulates in the cytoplasm. Like arp3, ARPC2/p34 concentration at the front of actin cones was abolished in myosin VI mutants (data not shown). Therefore, like cortactin, asymmetric distribution of the arp2/3 complex on the actin cones is dependent on myosin VI function. These findings support a role for myosin VI in regulating actin dynamics by participating in the localization of cortactin and arp2/3 complex at the front of the individualization complex.
Dynamin is localized on actin complexes of individualizing
spermatids
The close coupling between actin assembly and membrane remodeling during
individualization prompted us to examine proteins involved in membrane
dynamics to determine if myosin VI works with these proteins. Dynamin, encoded
by the Drosophila shibire gene, is a large GTPase with known roles in
promoting the fission of clathrin-coated pits into clathrin-coated vesicles
during endocytosis (Hinshaw,
2000; McNiven,
1998
). Dynamin binds cortactin and regulates actin dynamics at
membrane sites of actin assembly (Ochoa et
al., 2000
). Thus, dynamin is a likely molecule to function in
individualization complexes.
We examined dynamin distribution on individualizing spermatids and found that it localized along the length of the actin cones (Fig. 4A-C) in a distribution most similar to actin. As expected from the distribution of dynamin on actin cones, myosin VI concentrated at the front of dynamin-stained cones (Fig. 4D-F). Dynamin localization along the actin cones suggests that this region might either be an area of high endocytic membrane trafficking or a region where actin dynamics are regulated by dynamin. Furthermore, the close juxtaposition of myosin VI and dynamin suggests that they might participate in the same process during individualization.
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To gain a clearer understanding of dynamin's function in the actin cones,
we examined the distribution of two proteins known to interact with dynamin in
vertebrates, but which appear to function in different pathways in
Drosophila. -Adaptin is the
subunit of the AP-2
adaptor complex, which is known to bind clathrin and function in early
endocytosis (Hirst and Robinson,
1998
).
-Adaptin is also required for endocytosis in
Drosophila (Gonzalez-Gaitan and
Jackle, 1997
). Amphiphysin, on the other hand, is not required for
endocytosis in Drosophila
(Leventis et al., 2001
;
Razzaq et al., 2001
;
Zelhof et al., 2001
). However,
amphiphysin can influence filamentous actin localization and has been
implicated in membrane morphogenesis and organization in Drosophila
(Razzaq et al., 2001
;
Zelhof et al., 2001
).
-Adaptin was neither concentrated on actin cones nor at the front of
the cones, as indicated by anti-Drosophila
-adaptin antibodies
(Fig. 4G-I). Instead, it
localized in a particulate fashion throughout the cystic bulge ahead of the
actin cones. By contrast, Drosophila amphiphysin antibodies localized
to actin cones in a manner similar to dynamin
(Fig. 4K,B). However, unlike
dynamin, amphiphysin also concentrated at the front of cones in a manner
similar to myosin VI, cortactin and the arp2/3 complex. Thus, its distribution
is intermediate between dynamin and cortactin/myosin VI. As amphiphysin
localized to the actin cones whereas
-adaptin did not, we conclude that
dynamin on the actin cones participates in a non-endocytic function. We
hypothesize that its function is related to actin dynamics or organization on
the basis of the amphiphysin localization and further experiments (see
below).
Myosin VI and dynamin mutations genetically interact
The temperature-sensitive mutation in Drosophila dynamin,
shibire (shi1), was used to determine if dynamin
function was required for individualization. This mutation is in the GTPase
domain of dynamin (Grant et al.,
1998) and is thought to block the GTPase cycle of dynamin
(Hinshaw, 2000
), thereby
acting as a functional null at the non-permissive temperature. Exposure of the
dynamin mutant flies to the non-permissive temperature for up to 6
hours did not have any obvious effect on actin in actin cones or organization
of individualization complexes (Fig.
5A). After dynamin inactivation, myosin VI and cortactin
localization and accumulation at the front of actin cones was not
significantly affected either, although myosin VI accumulated abnormally
elsewhere in individualizing cysts (A.D.R., unpublished). Individualization
complexes moved normally when dynamin mutants were shifted to the
non-permissive temperature as judged by observations in real time using cysts
cultured in vitro (T. Noguchi, personal communication). Thus, dynamin
inactivation alone did not have a strong effect on individualization
complexes. However, striking defects were observed when the
temperature-sensitive dynamin mutation was placed into the background
of a hypomorphic myosin VI mutation, jaguar
(jar1). When these dynamin myosin VI double
mutants were exposed to the non-permissive temperature to inactivate dynamin
function, the total number of individualization complexes per testis was
dramatically reduced. In addition, most of the individualization complexes
that could be observed stained very weakly with phalloidin in comparison with
those in flies bearing only the dynamin or myosin VI
mutations (Fig. 5). We conclude
that actin assembly or stability is dramatically reduced in the dynamin
myosin VI double mutant. Therefore, we suggest that myosin VI and dynamin
function in parallel pathways and that each pathway contributes to the
regulation of actin structures in individualizing spermatids.
|
We quantified the defects in myosin VI dynamin double mutants to demonstrate the severity of the effect on actin structures. In these experiments we exposed dynamin myosin VI double mutants, dynamin single mutants, myosin VI single mutants and wild-type flies (OreR) to the non-permissive temperature for different times then fixed and stained the testes with phalloidin to visualize actin. We counted the total number of early individualization complexes (complexes that are just assembling or that have just begun to move; Fig. 6A) and determined the proportion of individualization complexes that stained strongly or weakly with phalloidin (Fig. 6B).
|
In wild-type flies there were typically 13 early individualization complexes per testis, of which most stained strongly with phalloidin (Fig. 6). These numbers were not significantly changed even after 6 hours at the non-permissive temperature. Similar numbers were observed for the dynamin mutants even after 6 hours at the non-permissive temperature. Therefore, exposing flies to as much as 6 hours of dynamin inactivation does not, obviously, affect actin structures.
In the myosin VI single mutants there was a slight reduction in
the total number of early actin complexes relative to wild-type and
dynamin mutant flies at all time points tested
(Fig. 6A). Approximately 40% of
the early complexes stained weakly with phalloidin
(Fig. 6B). These findings are
consistent with our previous results
(Hicks et al., 1999). In
myosin VI mutants, actin complexes form, but as complexes begin to
progress they fall apart and apparently lose actin filaments. This results in
fewer total actin complexes in myosin VI mutant testes and accounts
for the larger proportion of weakly staining complexes. Thus, absence of
myosin VI alone has a slight effect on actin assembly and/or stability in
early individualization complexes.
In comparison with wild-type controls and the single mutants, the dynamin myosin VI double mutant at the non-permissive temperature displayed a large reduction in the total number of early actin complexes per testis (Fig. 6A; 3 hours and 6 hours). Moreover, in the double mutant a large proportion of the actin complexes that remained stained weakly with phalloidin (e.g. 66% after 6 hours at the non-permissive temperature; Fig. 6B), indicating that those few complexes remaining had greatly reduced F-actin levels. By contrast, when the dynamin myosin VI double mutant was exposed to the non-permissive temperature for only 2 minutes, the total number of actin complexes per testis and the number of weakly staining complexes were similar to the myosin VI single mutant (Fig. 6). This demonstrates that dynamin myosin VI double mutants at the permissive temperature are similar to myosin VI single mutants, in that actin cones can assemble. Only after inactivation of dynamin are filamentous actin structures disrupted.
Since continuous actin assembly is often required to maintain stable actin structures, the loss of individualization complexes in the double mutant could be due to a disruption of actin filament assembly. Alternatively or in addition, the loss of actin filaments could be due to a direct disruption of actin filament stability. Therefore, we conclude that myosin VI and dynamin function in pathways that regulate actin assembly or stability in individualizing spermatids. We also conclude that, although dynamin activity is not required for actin cone formation, it plays a redundant role with myosin VI in regulating actin dynamics.
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Discussion |
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We propose a model in which myosin VI acts in a structural capacity in the
individualization complex to regulate actin dynamics at sites of active
membrane remodeling. Myosin VI might participate in actin dynamics solely by
its influence on cortactin and arp2/3 complex localization. Cortactin can bind
both actin and the arp2/3 complex (Weed et
al., 2000). Cortactin also enhances actin polymerization and
stabilizes actin filaments during arp2/3-complex-dependent actin
polymerization through its actin-binding activity
(Olazabal and Machesky, 2001
;
Weaver et al., 2001
). Myosin
VI could help localize or maintain the localization of arp2/3 complex and
cortactin at the fronts of the actin cones. The localization of these
components at the front of the cones would then facilitate actin assembly.
This localized actin assembly would drive actin cone movement in a manner
similar to the leading-edge protrusion or Listeria motility. In the
absence of myosin VI, the fronts of the cones would lose assembly sites and
thus actin cones would depolymerize. It is interesting to note that another
unconventional myosin, myosin 1 from yeast and Dictyosteliyum, can
interact with the arp2/3 complex and may have a direct role in actin assembly
(Evangelista et al., 2000
;
Geli et al., 2000
;
Jung et al., 2001
).
Alternatively, myosin VI might directly stabilize actin at the front of the actin cones. Since myosin VI has a coiled-coil domain that is thought to mediate its dimerization, myosin VI itself could crosslink actin filaments and provide a stabilizing force to newly generated actin filaments. Loss of myosin VI function in this scenario would result in destabilization of actin filaments and the actin-binding proteins associated with those filaments such as arp2/3 complex and cortactin. Thus, in this case, loss of arp2/3 complex and cortactin concentration would be secondary effects owing to loss of actin filaments at the front of actin cones. At this point, our results do not distinguish between a direct or indirect effect of myosin VI on actin assembly/stability.
A role for dynamin in actin dynamics
To explain the actin defects we observe in dynamin myosin VI
double mutants we also suggest that dynamin participates in a structural
capacity by helping to organize or stabilize the actin filaments of the cones.
The genetic analysis of the double mutants suggests that these genes function
in parallel pathways. Moreover, the localization of the two proteins on actin
cones is consistent with parallel separate pathways since their localization
is distinct from one another. Since dynamin is located along the cone, but is
not concentrated at the front, its role would be to help regulate actin
dynamics or organize actin away from the front where myosin VI and cortactin
are located. In this structural model, loss of both dynamin and myosin VI
function would be predicted to have severe effects on the assembly and
stability of actin filaments in the individualization complex, because two
regulators of actin structure in the individualization complex would be lost.
This is consistent with the dynamin myosin VI double mutant
phenotype.
The temperature-sensitive mutation in dynamin that we used for these
experiments was initially characterized in neurons where neuronal phenotypes
can be detected within minutes of dynamin inactivation
(Koenig and Ikeda, 1989;
Koenig et al., 1989
). We
observed that short periods of dynamin inactivation (2 minutes) in the myosin
VI background have no effect on actin. The difference in time course of effect
of dynamin inactivation might be explained by the inherently slower dynamics
of individualization, a process that takes place over the course of many
hours, as compared to the very rapid process of synaptic vesicle release and
re-uptake in neurons that requires dynamin.
There is the formal possibility that dynamin could participate in
clathrin-based endocytosis and not actin dynamics during actin cone movement,
but several lines of evidence argue against this possibility. First, a protein
normally associated with dynamin during endocytosis, -adaptin, does not
concentrate on actin complexes nor does it concentrate at the front of the
complexes like myosin VI does. Instead
-adaptin localizes to the cystic
bulge ahead of the cones. Second, electron micrographs of individualizing
spermatids do not show evidence of high endocytic activity at the
individualization complex (Tokuyasu et
al., 1972
). Third, the robust disruption of the actin
individualization complexes observed in the dynamin myosin VI double
mutant is not easily explained by a defect in endocytosis. Fourth, amphiphysin
localizes to actin cones in a similar fashion to dynamin. However, amphiphysin
does not appear to be required for endocytosis in Drosophila
(Leventis et al., 2001
;
Razzaq et al., 2001
;
Zelhof et al., 2001
). Although
it is unclear if Drosophila amphiphysin binds directly to dynamin
(Razzaq et al., 2001
), it is
clear that amphiphysin can function in non-endocytic processes that involve
actin structures and membrane morphogenesis
(Razzaq et al., 2001
;
Zelhof et al., 2001
).
Moreover, there is precedence for dynamin involvement in actin dynamics in
other organisms. For example, Dynamin-2 concentrates in actin-based podosomes
and regulates actin assembly and turnover in these structures
(Ochoa et al., 2000).
Dynamin-2 also localizes to actin comet tails of macropinocytes,
Listeria and type I PIP-kinase-induced motile vesicles and regulates
the assembly and structure of these actin tails
(Lee and De Camilli, 2002
;
Orth et al., 2002
). The
implication of dynamin in an actin assembly process is a novel finding in
Drosophila, but bolsters the recent observations that dynamin
functions to regulate actin structures in mammalian systems. This suggests
that actin assembly regulation is another conserved function for dynamin.
A role for myosin VI in linking actin assembly to membrane
dynamics
In addition to a role for myosin VI in localizing actin assembly proteins,
we speculate that myosin VI might be important for coupling actin assembly
sites to regions where the membrane is remodeled at the front of
individualization complexes. On the basis of electron micrographs of
individualizing spermatids (Tokuyasu et
al., 1972), membrane lies in close proximity to actin along the
length of the cone, including the front of the cone, and vesicles, organelles
and ribosomes are excluded from this region. The close proximity of the actin
cones to the plasma membrane suggests they are tightly linked. The front of
the actin cones is also the region where we see concentrated myosin VI and
concentrated sites of actin-assembly-regulating proteins. Immediately in front
of the cones there is an abrupt transition to the syncytial cytoplasm in the
cystic bulge, a region of large vesicles, organelles and ribosomes. Membrane
remodeling takes place at the junction between the front of the actin cones
and the syncytial cytoplasm. During individualization complex progression, the
connection between membrane and actin must be maintained, otherwise membrane
remodeling would not progress synchronously with movements of the actin
complex. Myosin VI, perhaps through its effects on cortactin localization,
might be part of the structure that links actin and membrane as the
individualization complex moves down the spermatids.
Moreover, the detection of cortactin at the front of actin cones where
myosin VI accumulates indicates that this region is not only a site of active
actin assembly but also a region of coupling between membrane and the actin
cytoskeleton. Cortactin is thought to act as a link between
membrane-associated kinases and actin assembly since it associates with
proteins such as Src, Syk and the arp2/3 complex and concentrates in zones of
membrane/actin linkage such as lamellipodia (Wu et al., 1993;
Okamura and Resh, 1995;
Maruyama et al., 1996
;
Weed et al., 1998
).
Myosin VI may be carrying out similar functions in other cellular
contexts
Our suggestion that myosin VI facilitates actin assembly through effects on
the arp2/3 complex and cortactin and couples actin dynamics to membrane
remodeling through either a direct or cortactin-dependent mechanism may be
applicable to myosin VI function in other systems. For example, vertebrate
myosin VI has been proposed to function in endocytosis
(Buss et al., 2001a;
Buss et al., 2001b
). During
endocytosis, plasma membranes undergo dynamic morphological change in order to
invaginate and form a budding vesicle. It has been proposed that F-actin
networks help deform membrane during coated-vesicle formation and that F-actin
polymerization helps propel endocytic vesicles away from the plasma membrane
(Qualmann et al., 2000
). In
these scenarios actin assembly is required and membranetopology changes must
be coordinated with actin assembly. Given such requirements it is not
surprising that cortactin has been suggested to function in endocytosis
(Jeng and Welch, 2001
;
Kaksonen et al., 2000
). We
speculate that myosin VI's role in this process may be to link actin assembly
sites containing arp2/3 complex and cortactin to sites of membrane dynamics
involving dynamin.
It is also tempting to consider that myosin VI might facilitate actin
assembly and its coupling with membrane reorganization in other cellular
contexts. Examples include membrane invagination in syncytial blastoderm
embryos (Mermall and Miller,
1995), membrane ruffling in EGF stimulated cells
(Buss et al., 1998
) and
stereocilia morphogenesis in developing hair cells
(Self et al., 1999
).
Open questions
It is unclear if the ability of myosin VI to translocate along actin
filaments is required for actin cone movement or function. In particular, it
is not known whether the minus-end movement of myosin VI is important. The
orientation of actin filaments in individualization complexes has not been
visualized directly, although the capping protein and arp2/3 complex
localization are consistent with the barbed ends of actin filaments at the
front of actin cones oriented away from the spermatid nuclei. If this is true,
then pointed-end motility would lead to myosin VI walking away from the
advancing edge. This seems inconsistent with maintaining a high concentration
of myosin VI at the front of the cones. Until the orientation of actin
filaments is known, the mobility of membrane and myosin VI observed, and the
dynamics of actin in the actin cones determined, it is premature to formulate
specific models for the involvement of myosin VI's actin-based motility in
this process. However, our studies of spermatid individualization are
providing insight into the molecules required for actin cone assembly and
movement, as well as myosin VI's specific role in this process. Our continuing
studies will allow us to develop and test models regarding myosin VI's in vivo
mechanism of action and ultimately determine the importance of its motile
properties in vivo.
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Acknowledgments |
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References |
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