1 Department of Zoology, University of Wisconsin, Madison, Madison, WI 53706,
USA
2 Program in Cellular and Molecular Biology, University of Wisconsin, Madison,
Madison, WI 53706, USA
* Author for correspondence (e-mail: wmbement{at}facstaff.wisc.edu )
Accepted 9 January 2002
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Summary |
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Key words: Cytoskeleton interactions, Actin, microtubules, Intermediate filaments
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Introduction |
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Xenopus ooctyes have extensive, polarized arrays of cortical
F-actin, microtubules, and intermediate filaments, and have consequently
provided a useful model system for analysis of cytoskeletal organization
(Gard, 1993;
Elinson et al., 1993
;
Klymkowsky et al., 1987
;
Klymkowsky, 1995
), as well as
for analysis of functional interactions among the various cytoskeletal systems
(Canman and Bement, 1997
;
Gard et al., 1997
;
Bement et al., 1999
;
Benink et al., 2000
). By
systematically manipulating the three filament systems independently and
analyzing the results of such manipulation on the remaining systems, Gard et
al. demonstrated the existence of a hierarchy of interactions among the three
systems in Xenopus oocytes (Gard
et al., 1997
). Specifically, long-term disruption of F-actin
altered the distribution of both microtubules and cytokeratin filaments,
perturbation of microtubules altered cortical cytokeratin filament polarity,
while perturbation of intermediate filaments had little effect on either
microtubules or F-actin.
Based on their results, Gard et al. proposed that the F-actin network was
somehow responsible for maintaining the normal, polarized distribution of
cytokeratin (Gard et al.,
1997). However, as in many other cell types, the three filament
systems have mutually exclusive fixation requirements in Xenopus
oocytes (McBeath and Fujiwara,
1984
; Vielkind and Swierenga,
1989
; Gard et al.,
1995
), preventing direct demonstration of interactions among the
three systems. Therefore, it is unclear how, exactly, the three systems impact
each other.
We recently developed Xenopus egg extracts as a model system for
analysis of microtubuleF-actin interactions
(Sider et al., 1999;
Waterman-Storer et al., 2000
).
In such extracts, both microtubules and F-actin rapidly assemble following
warming to room temperature, and their relative distributions can be analyzed
at high resolution by confocal fluorescence microscopy. This approach permits
analysis of interactions between cytoskeletal systems in both fixed and
unfixed samples by allowing simultaneous visualization of F-actin,
microtubules, and cytokeratin. Further, because the extracts are totally
accessible, each of the filament systems can be perturbed by pharmacological
or immunological means separately or together, without the need for
microinjection.
Two other features of the system make it ideal for analysis of interactions
of microtubules and F-actin with intermediate filaments. First, cytokeratin is
the only cytoplasmic intermediate filament protein found in Xenopus
oocytes and eggs (Franz et al.,
1983; Franz and Franke,
1986
), which simplifies interpretation of results. Second, the
cytokeratin network is disassembled into soluble oligomers of
750 kDa in
the meiotically mature egg (the source of extracts), but reassembles following
egg activation into a complex network of mature cytokeratin filaments in the
embryo (reviewed by Klymkowsky,
1995
). This permits the contributions of the microtubule and
F-actin networks to cytokeratin assembly to be assessed in the absence of a
pre-existing cytokeratin network. Here, the assembly of the cytokeratin
network in extracts was analyzed using a combination of static and time-lapse
confocal fluorescence microscopy, and manipulation of microtubules, F-actin
and the cytokeratin system itself. The results demonstrate that, while all
three cytoskeletal networks interact in vitro, the assembly and organization
of the cytokeratin network is crucially dependent on the assembly and
organization of the F-actin cytoskeleton. Further, it is shown that normal
organization of the cytokeratin network in intact, activated eggs relies on
the actin cytoskeleton. We conclude that cytokeratin assembly and organization
in the Xenopus system is critically dependent on the actin
cytoskeleton both in vitro and in vivo.
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Materials and Methods |
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Preparation of demembranated sperm
Demembranated sperm were prepared as described
(Sider et al., 1999). Adult
male Xenopus were killed by decapitation, their testes were removed
and rinsed three times in ice-cold MBSH (110 mM NaCl, 2 mM KCl, 1 mM
MgSO4, 0.5 mM NaPO4, 2 mM NaHCO3, 15 mM
Tris-base, pH 7.6). Testes were then macerated in an ice-cold pyrex Petri
plate in 4 ml ice-cold HSPPP (250 mM sucrose, 15 mM Hepes pH 7.4, 1 mM EDTA,
0.5 mM spermidine, 0.2 mM spermine, 10 µg/ml pepstatin, leupeptin,
aprotinin and chymostatin, 1 mM Pefabloc). The macerated testes were then
filtered through cheese cloth, pelleted in a clinical centrifuge at 4°C,
resuspended in HSPPP, pelleted, resuspended in 1 ml room temperature HSPPP
with 500 µg/ml lysophosphatidylcholine (Boehringer), and incubated 5
minutes at room temperature. Sperm were then diluted in 10 ml HSPPP plus 0.3%
BSA, pelleted, resuspended in HSPPP plus BSA, pelleted again, resuspended in
HSPPP plus BSA and 30% glycerol at a final concentration of
1000
sperm/µl, frozen in liquid N2 and stored at -80°C. Prior to
use in experiments, sperm were diluted in 200 µl microtubule stabilization
(TSB) buffer (30 mM K-Pipes, pH 70, 5 mM MgCl2, 1 mM EGTA),
pelleted and resuspended in 50 µl TSB. Demembranated sperm were added to
extracts to nucleate microtubule asters, however, sperm were not required for
cytokeratin assembly in extracts nor for cytokeratinF-actin
interactions.
Manipulation of the cytoskeleton
Microtubules or F-actin were disrupted by adding 20 µM nocodazole
(Calbiochem) or 10 µM latrunculin B (Calbiochem), respectively. Cytokeratin
assembly was blocked by adding anti-cytokeratin antibody (Sigma) to 5
mg/ml.
-actinin was obtained from Cytoskeleton Inc. (Denver, CO),
reconstituted in water at a concentration of 2.5 mg/ml, frozen in liquid
nitrogen and stored at -80°C.
Flow chamber analysis
Flow chambers were constructed as described
(Mandato et al., 2000).
Polylysine (Sigma)-treated coverslips were inverted over two pieces of
parallel, double-stick tape adhered to microscope slides such that the volume
of the flow chamber was 10 µl. Demembranated sperm were pipetted into
chambers and allowed to adhere for 10 minutes. Chambers were washed five times
with TSB containing 5 mg/ml BSA (TSB/BSA). Egg extracts were prepared by
thawing on ice, adding 2 µl rhodamine tubulin (10 mg/ml; Cytoskeleton Inc.)
and 2 µl 10x ATP regenerating system (100 mM creatine kinase, 100 mM
creatine phosphate, 10 mM ATP) (Leno and
Laskey, 1991
), and incubating on ice for 2 hours
(Sider et al., 1999
).
Subsequently, 10 µl of extract were pipetted into chambers, which were then
incubated at room temperature for 15 minutes. Chambers were then washed ten
times with TSB/BSA plus 20 µM taxol (TSBT/BSA), followed by incubation with
TSBT/BSA plus 1 U/ml Alexa-488 phalloidin (Molecular Probes) for 10 minutes
and washing five times with TSBT. Samples were mounted with 10 µl 80%
glycerol in 1xPBS containing 20 mM N-propyl gallate.
Cytokeratin was labeled using anti-cytokeratin antibody at 1:250 (Sigma)
followed by Cy5 anti-mouse secondary antibody at 1:100 (Amersham). Samples
were examined via a confocal laser-scanning microscope.
Rapid freeze analysis
Rapidly frozen extract samples were prepared as described
(Mandato et al., 2000).
Briefly, clean 22x22 mm and 22x25 mm coverslips were treated with
casein (Sigma) and Rain-X (Unelko, Scottsdale, AZ), respectively. 1.5 µl
egg extract was pipetted onto large coverslips and small coverslips were
inverted onto the extract. The assembly was incubated at room temperature for
15-20 minutes and then submerged in liquid nitrogen for 2-3 minutes. The small
coverslip was rapidly pried off and immediately submerged in fix (80 mM
K-Pipes, pH 6.8, 5 mM EGTA, 1 mM MgCl2, 3.7% paraformaldehyde,
0.25% gluteraldehyde, 0.2% Triton X-100, 1.0 µM Taxol). After 5 minutes,
the samples were washed twice in PBS plus 0.1% NP-40 (PBSN). After washing,
the fix reaction was quenched for 10 minutes in PBSN plus 100 mM sodium
borohydride and washed again twice in PBSN plus 5 mg/ml BSA. This was followed
by incubation for 15 minutes at 37°C in primary and secondary antibody,
each of which was followed by washing five times in PBSN plus 5 mg/ml BSA. The
following antibodies/probes were used to label the cytoskeletal systems:
microtubules were labelled with monoclonal anti-
-tubulin at 1:250
(Amersham) or polyclonal anti-tyrosinated tubulin and anti-mouse or rabbit
rhodamine or Oregon green labeled secondary antibody at 1:100; F-actin was
labelled with Texas Red or Alexa-488-labeled phalloidin at 1:100 (Molecular
Probes); and cytokeratin with a monoclonal anti-cytokeratin antibody at 1:250
(Sigma) and anti-mouse Cy5 labeled secondary at 1:100. Subsequent to all
antibody applications and washing, samples were mounted on a slide with 5
µl 80% glycerol in PBS containing 20 mM N-propyl gallate.
4D microscopy analyses
1 µl of monoclonal anti-cytokeratin antibody (Sigma), 1.5 µl of
anti-mouse FITC-labeled Fabs (Jackson ImmunoResearch), 10 µl PBS and 0.24
U/ml Texas Red phalloidin were combined and incubated on ice for 1 hour prior
to use. 1 µl of 10x ATP regenerating system and 20 µM nocodazole
were added to 25 µl of extract, which was subsequently put into 5 µl
aliquots and incubated on ice for at least 1 hour. A 5 µl aliquot was
incubated at room temperature for 15 minutes, after which 1 µl of the
antibody solution was added to the extract, which was quickly mixed, and then
2 µl was pipetted onto a slide and a cleaned, casein-treated coverslip was
inverted onto the extract. The slide was sealed and observed for 15 minutes,
taking scans every 30 seconds.
Microtubule/F-actin assembly were analyzed by adding 2 µl 10x ATP
regenerating system, 2 µl demembranated sperm, 2 µl Oxyrase (Oxyrase
Inc.), 0.15 mg/ml rhodamine tubulin and
0.25 mg/ml Oregon
Green-conjugated g-actin to 60 µl of extract. This extract solution was
then incubated on ice for 2 hours to allow full incorporation of fluorophore
labeled proteins. Samples were prepared as above, taking scans every 15
seconds.
In vivo cytokeratin immunofluorescence
Oocyte procurement and fixation are described in detail elsewhere
(Canman and Bement, 1997).
Briefly, ovaries were recovered from mature female Xenopus laevis,
immediately placed in OR2 (82.5 mM NaCl, 2.5 mM KCl, 1 mM CaCl2, 1
mM MgCl2, 1 mM Na2HPO4, 5 mM Hepes, pH 7.4),
subjected to 1% collagenase treatment, rinsed six to eight times in OR2 and
allowed to recover. Eggs were obtained by immersion of oocytes in 5 µg/ml
progesterone (Sigma) for 1 hour, followed by overnight incubation in OR2. 10
µM latrunculin for 1 hour was used to perturb F-actin. Eggs were activated
via a 5 minute incubation in 10 µM ionomycin (Sigma) in Ca2+
followed by 40 minutes in OR2. For visualization of cytokeratin, eggs were
fixed overnight in -20°C methanol, permeabilized in PBS plus 10% glycerol
and 0.15% Triton X-100, and then washed in Tris buffered saline (TBS: 100 mM
Tris, pH 7.5, 0.9% NaCl) plus 0.1% NP-40 and 5 mg/ml BSA (TBSN/BSA). Fixed
eggs were bisected, blocked with TBSN/BSA, incubated in monoclonal
anti-cytokeratin antibody (Sigma) at 1:250 in TBSN/BSA, washed, incubated in
rhodamine anti-mouse at 1:100, and washed again. Samples were mounted and
examined on a confocal laser-scanning microscope.
Image analysis, processing, and quantification
Samples were analyzed using BioRad 1024 confocal laser-scanning
fluorescence microscopes at the Keck Center for Neuroscience and in the
Department of Zoology (NSF 9724515; James Pawley, primary investigator),
University of Wisconsin. Images were obtained with a 63x 1.4 NA
objective; higher magnification images were obtained using the zoom function.
Images were processed and analyzed using Adobe Photoshop, NIH Image v20.06,
Microsoft Office PowerPoint and Excel. For
Fig. 3C, Adobe PhotoShop was
used to quantify overlap of the green channel (representing F-actin) and the
blue channel (representing cytokeratin). This was determined in pixel number
by selecting a specific color range to represent overlap. The blue channel was
then selected and rotated 90° clockwise, and overlap of the blue and green
signal was again determined (Kaverina et
al., 1998).
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Results |
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Confocal fluorescence analysis of unfixed, triple-labeled specimens demonstrated that both F-actin and cytokeratin filaments associate with astral microtubules. Although both F-actin and cytokeratin could be seen near the base of asters, the associations of the three systems were most easily observed near the peripheries of the asters (Fig. 1). All three filament systems could be found in close proximity to each other (Fig. 1, arrows); however, it was also apparent that microtubuleF-actin association could be observed in the absence of cytokeratin (Fig. 1, arrowheads), and microtubule-cytokeratin interactions could be observed in the absence of F-actin (Fig. 1, chevrons).
|
The above results suggested that while all three systems could
co-associate, pairwise associations did not necessarily require the presence
of the third system. To test this point directly, extracts were subjected to
treatments designed to perturb each of the three cytoskeletal systems.
Inhibition of F-actin polymerization by addition of latrunculin B to extracts
prior to filament polymerization in flow chambers failed to prevent
cytokeratin association with aster microtubules
(Fig. 2, latrunculin, arrows),
indicating that microtubule-cytokeratin interactions were not strictly F-actin
dependent. However, it was observed that the cytokeratin associated with
microtubules in the absence of F-actin was somewhat less filamentous than that
seen in controls. Cytokeratin filaments were disrupted by the addition of the
monoclonal anticytokeratin antibody C11. C11 has previously been shown to
cause cytokeratin disassembly in Xenopus oocytes
(Gard et al., 1997;
Canman and Bement, 1997
). The
absence of cytokeratin filaments failed to prevent F-actin association with
aster microtubules (Fig. 2,
C11, arrows). However, in the absence of cytokeratin filaments,
fewer asters were generally found in flow chambers, and those that were found
were generally less organized than those found in control samples, suggesting
that the presence of cytokeratin filaments stabilized asters during the
washing process.
|
Assessment of potential cytokeratinF-actin interactions in the
absence of microtubules was problematic in this assay, since most of the
material that is not bound to aster microtubules is removed during the washing
process (Sider et al., 1999).
Nevertheless, occasional examples of cytokeratinF-actin interactions
could be observed in the absence of microtubules
(Fig. 2, nocodazole, arrows).
Thus, cytokeratinF-actin interaction was not strictly dependent on
microtubules.
F-actin, microtubules and cytokeratin associate in rapidly frozen
extract samples
The flow chamber analysis indicated that the three systems can interact
under native (unfixed) conditions. However, the washing process distorts the
organization of the cytoskeletal networks and, in particular, is likely to
cause closer packing of the different systems than would otherwise be
observed. As an alternative approach that does not entail the extensive
washing necessary for flow chamber analysis, rapid-freezing of thin extract
specimens was employed. Small volumes of extract were sandwiched between two
coverslips, warmed to room temperature to allow polymer formation and then
plunged into liquid nitrogen. Samples were then rapidly fixed and
fluorescently stained for the three cytoskeletal systems (see Materials and
Methods) (Mandato et al.,
2000).
In control samples, all three filament systems were observed to interact in
a lengthways fashion, as observed in the flow chamber assay. However, the lack
of washing required by the flow chamber analysis made it possible to visualize
isolated microtubules, F-actin cables and cytokeratin filament cables
(Fig. 3A,B). While some
cytokeratin was observed to associate with microtubules in the absence of
F-actin, most of the cytokeratin was associated with F-actin cables
(Fig. 3A, arrows). In many of
the images, it appeared that the distal ends of microtubules were bent at
points of contact with cables of F-actin and cytokeratin, as if the
microtubules were tracking along F-actin and/or cytokeratin cables
(Fig. 3A, arrowheads). It was
also observed that F-actin was cleared from the immediate vicinity of the
microtubule organizing centres, and was most obvious near the periphery of the
asters, consistent with previous observations
(Sider et al., 1999;
Waterman-Storer et al., 2000
).
These cytoskeletal interactions are also illustrated by a 3D reconstruction
(Fig. 3B) that shows both an en
face view of a rapid freeze experiment, and the same view at a tilt of
70°. The interaction of F-actin and cytokeratin is apparent in both views
(Fig. 3B, arrowheads), implying
that overlap seen en face does indeed represent colocalization.
To measure colocalization of F-actin and cytokeratin and to provide
evidence that this colocalization is not due to chance, we employed a
variation of a quantification method [described by Kaverina et al.
(Kaverina et al., 1998)].
Colocalization was quantified by determining the total pixel number of
overlapping signal from the two channels in the original images and then in
images in which the signal representing cytokeratin was rotated 90°
clockwise. These data are represented in
Fig. 3C and show that no more
than 6% of the observed colocalization can be accounted for by chance
(P<0.05).
Interactions among the three cytoskeletal systems were then evaluated in rapidly frozen samples following systematic perturbation of each of the three cytoskeletal systems, as described above for the flow chamber assays. When microtubule polymerization was prevented by treatment of extracts with nocodazole, cytokeratin filaments still formed in association with F-actin cables (Fig. 4, nocodazole, arrowheads). When F-actin polymerization was prevented by treatment with latrunculin, occasional cytokeratin-microtubule interactions were still observed (Fig. 4, latrunculin, arrowhead), consistent with the flow chamber results. However, much of the cytokeratin was found on the substrate in aggregates (Fig. 4, latrunculin, arrow). When cytokeratin polymerization was prevented by treatment of extracts with the C11 antibody, F-actin cables and microtubules formed and associated with each other (Fig. 4, C11, arrowheads). However, as observed for the flow chamber assays, the number of astral arrays of microtubules was reduced relative to controls, suggesting that the presence of cytokeratin filaments stabilizes the microtubule and F-actin networks during processing for immunofluorescence. When both F-actin and microtubule polymerization was prevented by the combined use of latrunculin and nocodazole, most of the cytokeratin was found in large, unorganized aggregates (Fig. 4, NOC/LAT).
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CytokeratinF-actin association changes over time
To assess the relationship between cytokeratin assembly and F-actin
assembly over time, samples were processed at increasing intervals following
warming to room temperature. In specimens fixed immediately after preparation,
F-actin was not observed, and most of the cytokeratin was found as
particulates on the substrate (Fig. 5,
0'). At 5 and 10 minute time points, F-actin cables formed,
and cytokeratin was co-distributed with such cables
(Fig. 5), suggesting that
cytokeratin assembles on the forming F-actin cables. By 20 minutes, both
cytokeratin and F-actin were assembled into networks composed of large cables
(Fig. 5). Occasionally, it
appeared as if cytokeratin networks were abandoning their association with the
F-actin network and forming an independent system
(Fig. 5, 20',
arrowheads).
|
Manipulation of F-actin organization alters the organization of the
cytokeratin network
The foregoing results suggested that F-actin cables might actually serve as
templates upon which cytokeratin filaments assemble. If this interpretation is
correct, it would be predicted that changing the organization of the F-actin
network would result in a corresponding change in the cytokeratin network. To
test this prediction, exogenous -actinin, an actin-binding protein that
crosslinks F-actin into fine meshworks, was added to extracts. After allowing
sufficient time for F-actin and cytokeratin assembly, extracts were rapidly
frozen and stained for F-actin and cytokeratin as described above.
Confocal fluorescence analysis of F-actin in control extracts and extracts
containing exogenous -actinin revealed that while large F-actin cables
were found in both types of samples, extracts containing
-actinin also
had F-actin in tight meshworks of fine cables
(Fig. 6). These fine F-actin
meshworks coincided with fine meshworks of cytokeratin, indicating that the
change in F-actin organization relative to controls resulted in the predicted
change in cytokeratin filament organization. In addition, it was apparent at
higher magnification that the F-actin-bound cytokeratin appeared punctate,
although whether that was a function of increased density of F-actin or
competition of cytokeratin with exogenous
-actinin for F-actin binding
is disputable.
|
To quantify the effects of -actinin on the spatial patterns of both
F-actin and cytokeratin, the mean distance between adjacent F-actin cables and
cytokeratin was calculated in both control and
-actinin-supplemented
samples. The addition of
-actinin significantly decreased the mean
distance between both adjacent cytokeratin and F-actin cables
(Fig. 6;
P<0.05).
Dynamic interactions between F-actin and cytokeratin
To better understand the dynamic relationship between F-actin and
cytokeratin filaments as they assemble over time
(Fig. 5), we used 4D confocal
fluorescence microscopy to observe the dynamic interactions of F-actin and
cytokeratin in Xenopus egg extracts. F-actin was labeled with Texas
Red phalloidin, and cytokeratin by addition of low concentrations of the
monoclonal anti-cytokeratin antibody C11, which previously had been bound to
anti-mouse FITC-labeled FAbs. This approach led to punctate labeling of the
cytokeratin network, permitting it to be followed via 4D microscopy. After
addition of the above reagents, extracts were warmed to room temperature for
15 minutes and then examined by 4D microscopy. Consequently, the 0' time
point in Figs 7,
8 and
9 refer to the times at which
imaging was started, in contrast to Fig.
5, where the 0' time point referred to samples prepared
before warming to room temperature.
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Consistent with the above analysis of static specimens, F-actin networks
were typically associated with cytokeratin. As the actin cables began to
contract and `zipper' (Waterman-Storer et
al., 2000), the associated cytokeratin showed both a tendency to
move with the F-actin to which it was bound
(Fig. 7B, arrowheads), and the
ability to disengage from F-actin (Fig.
7A, arrowheads and arrows). When latrunculin B was added to
extracts, cytokeratin failed to form any filamentous structures and collected
in aggregates on the substrate over time, similar to results in rapidly frozen
samples (data not shown). Conversely, addition of an inhibitory concentration
(5 mg/ml) of anticytokeratin antibody did not affect formation and contraction
of F-actin networks, despite the fact that cytokeratin was prevented from
initially binding early F-actin networks and, subsequently, more extensive
cytokeratin networks were never formed (data not shown). This indicates that
whereas F-actin is required for proper cytokeratin filament organization in
extracts, the reverse is not true.
To determine the effects of experimental manipulations of the actin
filament network organization on dynamic cytokeratin-actin filament
interactions, -actinin was added to extracts prior to analysis via 4D
microscopy. The addition of
-actinin resulted in inhibition of the
F-actin `zippering' activity typically seen in these experiments
(Fig. 8). The
-actinin
also inhibited cytokeratin movement and coalescence (data not shown).
Time-lapse confocal microscopy was also used to confirm the results in the
fixed samples that led to the conclusion that the disruption of cytokeratin
filament network formation does not interfere with microtubuleF-actin
interactions. By adding demembranated sperm, rhodamine-labeled tubulin and
Oregon Green-conjugated g-actin to extracts
(Waterman-Storer et al.,
2000), we were able to observe dynamic microtubule aster growth
and expansion and the effects of these organized microtubules on surrounding
F-actin. As shown in Fig. 9,
there was no apparent difference in the effects of microtubule aster growth
and expansion on surrounding F-actin networks with or without the addition of
inhibitory concentrations of anti-cytokeratin antibody. That is, in both cases
F-actin was cleared from the region of the aster, a process which has been
shown to be dependent on microtubule-F-actin interactions
(Waterman-Storer et al.,
2000
).
F-actin is required for cytokeratin network assembly in intact
eggs
The cytokeratin network is disassembled in Xenopus eggs, but
reassembles upon egg activation/fertilization. To analyze the role of F-actin
in cytokeratin network assembly in vivo, and to confirm the results obtained
with extracts, Xenopus eggs were treated with latrunculin or, as a
control, DMSO (the vehicle for latrunculin), and then artificially activated.
Following activation, the eggs were fixed and stained with anti-cytokeratin
antibodies. In both the control and latrunculin-treated, unactivated eggs, the
cytokeratin network was disassembled (Fig.
10A,C), although cytokeratin aggregates not present in controls
were seen in latrunculin-treated, unactivated eggs. Following activation,
control eggs developed a network of fine cytokeratin filaments in the cortex
(Fig. 10B) that appear better
organized and less particulate than those in extracts, perhaps due to the
greater amount of time allowed for assembly in vivo (45' versus
15'). By contrast, the cytokeratin network in latrunculin-treated,
activated eggs was consistently abnormal (44/44 eggs from three different
females). The abnormality ranged from moderate, in which cytokeratin was
present in unusually thick, looped arrays
(Fig. 10D) to severe, in which
the cytokeratin was present in large disassembled aggregates
(Fig. 10E). Fig. 10F is a bar graph
representing the quantification of the difference in width of cytokeratin
filaments and/or aggregates in control versus latrunculin-treated, activated
eggs in two experiments. The latrunculin treatment significantly increased
cytokeratin array thickness relative to controls
(Fig. 10F;
P<0.05). These results demonstrate that F-actin is required for
normal cytokeratin assembly and organization in vivo as well as in vitro.
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![]() |
Discussion |
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Because most of the experiments were performed using cell-free extracts,
and because it is not yet possible to preserve both F-actin and cytokeratin
simultaneously in fixed Xenopus eggs
(Gard et al., 1995), we cannot
be certain of the extent to which proper cytokeratin assembly is dependent on
F-actin in intact eggs. However, the fact that large aggregates of cytokeratin
observed in latrunculin-treated extracts were also observed in
latrunculin-treated, activated eggs indicates that cytokeratin assembly in
eggs is at least partially dependent on F-actin in vivo. An in vivo
interaction between cytokeratin and F-actin is also suggested by the
demonstration that long term culture of Xenopus oocytes in
cytochalasin results in disorganization of the cytokeratin network and
aggregation of cytokeratin filaments (Gard
et al., 1997
). In the oocyte, the changes in cytokeratin induced
by F-actin disruption are slower and much less severe than we observed in
extracts or activated eggs, but there is an important difference between the
two developmental states. That is, in the oocyte, the cytokeratin network is
already assembled, whereas in the egg it is disassembled and is reassembled
only upon egg activation. Thus, the drastic effects of F-actin disruption on
the cytokeratin network in egg extracts and activated eggs, compared with its
less severe effect on cytokeratin in oocytes, may indicate that the role
played by cytokeratin-F-actin interactions changes as the network
assembles.
In support of this possibility, the time-course analysis revealed that, in
early extract samples, cytokeratin cables were invariably associated with
F-actin cables whereas, at later time points, cytokeratin cables were found
without associated F-actin, suggesting that, after assembly, cytokeratin can
release F-actin. Further, imaging of dynamic specimens allowed us directly to
observe the dissociation of F-actin from cytokeratin. These results indicate
that the initial steps of cytokeratin assembly rely on F-actin, but as the
filaments mature, they can release F-actin. Such behavior is curious, but is
consistent with previous demonstrations of F-actinintermediate filament
associations (e.g. Cary et al.,
1994), particularly those observed in cellular regions where de
novo assembly of intermediate filament structures occurs
(Green et al., 1986
;
Green et al., 1987
). A role
for F-actin as a general mediator of intermediate filament assembly is also
suggested by the results of a recent study of vimentin-fimbrin interactions in
macrophages. Specifically, Correia et al. demonstrated colocalization of
vimentin and the actin bundling protein fimbrin in retraction fibers and foci
at the leading edges of spreading macrophages
(Correia et al., 1999
). They
demonstrated that fimbrin binds specifically to soluble (i.e. nonassembled)
vimentin and is not associated with mature vimentin filaments in the cell
interior. Further, they pointed out that the binding site for fimbrin is in
the same region of vimentin required for self assembly
(Correia et al., 1999
). Thus,
it is reasonable to suggest that cytokeratin may initially be targeted to
actin cables in the disassembled form via an actin-binding protein such as
fimbrin or calponin (Mabuchi et al.,
1997
), and then released from F-actin as a result of
assembly-dependent dissociation from that actin-binding protein. If this is
correct, it may be that at least part of the effects of
-actinin on
cytokeratin distribution in extracts results not just from changing the
organization of the F-actin network, but also from competing with an
endogenous crosslinker, since
-actinin binds to F-actin via calponin
homology domains, as do fimbrin and, of course, calponin itself (reviewed by
Matsuidaira, 1991).
This study focused primarily on interactions between cytokeratin and
F-actin; however, several other findings should be considered. First, while we
often found evidence for three-way overlap of cytokeratin, F-actin and
microtubules, systematic perturbation of the three systems clearly
demonstrated that each system could interact with the other in the absence of
the third. These results, as well as the demonstration that microtubule asters
clear F-actin in the absence of cytokeratin, demonstrate that the previously
observed interactions between microtubules and F-actin in Xenopus
extracts (Sider et al., 1999;
Waterman-Storer et al., 2000
)
do not require cytokeratin filaments. In addition, although cytokeratin
organization is critically dependent on F-actin, the reverse is not true in
extracts, as shown by the fact that actin cables assemble and undergo dynamic
zippering even when cytokeratin assembly is prevented. Nevertheless, this does
not mean that inhibition of cytokeratin assembly has no effect on the other
cytoskeletal systems in extracts. In fact, both the actin networks and
microtubules became extremely sensitive to the processing necessary to view
flow chamber samples and rapidly frozen specimens. These results indicate that
intermediate filaments provide mechanical strength not only within the context
of intact cells and tissues (Yang et al.,
1996
; Goldman et al.,
1996
), but also in the context of isolated cytoplasm.
The bending of microtubules at the sites of
microtubuleF-actincytokeratin contact was also observed
repeatedly. Direct comparisons with the literature are impossible, but the
exertion of force implied by the bent microtubules is consistent with the
observation that actomyosin-dependent contractility results in microtubule
movement, bending, and breakage both in vivo
(Waterman-Storer and Salmon,
1997; Odde et al.,
1999
; Yvon and Wadsworth,
2000
; Yvon et al.,
2001
) and in vitro (Sider et
al., 1999
). The redirection of microtubules at points of F-actin
cable contact also supports the proposal of Kaverina et al. that microtubules
are guided to focal adhesions by interaction with F-actin
(Kaverina et al., 1998
).
In summary, our results indicate that F-actin is critically important for
proper cytokeratin organization in both Xenopus extracts and
activated eggs. It will therefore be of great interest to characterize the
means by which the two systems are linked. In addition to the possible
short-term linkage via a calponin homology domain containing proteins (see
above), several proteins have been identified that provide stable linkages
between F-actin and intermediate filaments (e.g.
Karakesisoglou et al., 2000;
Yang et al., 1996
). It may be
that the cell employs both stable and dynamic linkers of the two systems, to
provide the rigidity and flexibility that would be required during the complex
process of early development. That is, our results suggest that early in the
process of cytokeratin assembly, the two systems are extensively linked,
whereas later in the process, the linkage is less extensive. It could
therefore be plausibly argued that one class of linker is responsible for
joining assembling cytokeratin to F-actin, whereas another is responsible for
keeping the two systems tethered after assembly.
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References |
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