F-actin serves as a template for cytokeratin organization in cell free extracts

Kari L. Weber1 and William M. Bement*,1,2

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


    Summary
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 Summary
 Introduction
 Materials and Methods
 Results
 Discussion
 References
 
The microtubule, F-actin, and intermediate filament systems are often studied as isolated systems, yet the three display mutual interdependence in living cells. To overcome limitations inherent in analysis of polymer-polymer interactions in intact cells, associations between these systems were assessed in Xenopus egg extracts. In both fixed and unfixed extract preparations, cytokeratin associated with F-actin cables that spontaneously assembled in the extracts. Time-course experiments revealed that at early time points cytokeratin cables were invariably associated with F-actin cables, while at later time points they could be found without associated F-actin. In extract samples where F-actin assembly was prevented, cytokeratin formed unorganized aggregates rather than cables. Dynamic imaging revealed transport of cytokeratin by moving F-actin as well as examples of cytokeratin release from F-actin. Experimental alteration of F-actin network organization by addition of {alpha}-actinin resulted in a corresponding change in the organization of the cytokeratin network. Finally, pharmacological disruption of the F-actin network in intact, activated eggs disrupted the normal pattern of cytokeratin assembly. These results provide direct evidence for an association between F-actin and cytokeratin in vitro and in vivo, and indicate that this interaction is necessary for proper cytokeratin assembly after transition into the first mitotic interphase of Xenopus.

Key words: Cytoskeleton interactions, Actin, microtubules, Intermediate filaments


    Introduction
 Top
 Summary
 Introduction
 Materials and Methods
 Results
 Discussion
 References
 
The three major cytoskeletal systems — microtubules, actin filaments (F-actin) and intermediate filaments — are often considered in isolation, based on their differential protein composition, different functional properties and differential distributions within cells. Nevertheless, a wealth of information indicates that the three systems are mutually interdependent and that this interdependence is critical for normal cell function (Gard 1999Go; Allen and Shah, 1999Go; Canman and Bement, 1997Go; Gard et al., 1997Go; Wang et al., 2001Go). Microtubule—F-actin interactions, for example, underlie several important cell motility phenomena (reviewed by Waterman-Storer and Salmon, 1999Go), while association with microtubules is thought to control the organization (Gard et al., 1997Go), distribution (Gurland and Gundersen, 1995Go) and translocation (Prahlad et al., 1998Go) of intermediate filaments. Whether F-actin and intermediate filaments are also interdependent is less clear, but several reports indicate that, in some cases, the F-actin cytoskeleton interacts functionally with the intermediate filament cytoskeleton (e.g. Green et al., 1986Go; Green et al., 1987Go; Tint et al., 1991Go; Svitkina et al., 1998Go; Correia et al., 1999Go) (see also Discussion).

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, 1993Go; Elinson et al., 1993Go; Klymkowsky et al., 1987Go; Klymkowsky, 1995Go), as well as for analysis of functional interactions among the various cytoskeletal systems (Canman and Bement, 1997Go; Gard et al., 1997Go; Bement et al., 1999Go; Benink et al., 2000Go). 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., 1997Go). 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., 1997Go). However, as in many other cell types, the three filament systems have mutually exclusive fixation requirements in Xenopus oocytes (McBeath and Fujiwara, 1984Go; Vielkind and Swierenga, 1989Go; Gard et al., 1995Go), 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 microtubule—F-actin interactions (Sider et al., 1999Go; Waterman-Storer et al., 2000Go). 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., 1983Go; Franz and Franke, 1986Go), 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, 1995Go). 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.


    Materials and Methods
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 Summary
 Introduction
 Materials and Methods
 Results
 Discussion
 References
 
Preparation of interphase Xenopus egg extracts
Interphase Xenopus egg extracts were prepared as described (Sider et al., 1999Go). Briefly, eggs were stripped from adult Xenopus females induced to ovulate, and washed repeatedly in 1x MMR (100 mM NaCl, 2 mM KCl, 1 mM MgCl2, 0.1 mM EDTA and 5 mM Hepes, pH 7.8). Eggs were dejellied in 0.25x MMR containing 2% cysteine, rinsed four times in 0.25x MMR, rinsed twice in ice-cold ELB buffer (100 mM KCl, 0.1 mM CaCl2, 1 mM MgCl2, 10 mM K-Hepes, pH 7.7, 1 mM DTT, and 250 mM sucrose) and twice in ice-cold ELB buffer with 10 µg/ml of the protease inhibitors pepstatin, leupeptin, chymostatin, aprotinin and 1 mM Pefabloc (Boehringer-Mannheim, Indianapolis, IN). Eggs were then transferred to clear ultracentrifuge tubes, packed by a brief centrifugation in a clinical centrifuge, and the excess buffer was removed. The above protease inhibitors and cycloheximide (Sigma) were added to a final concentration (relative to packed egg volumes) of 10 µg/ml and 100 µg/ml, respectively. Eggs were crushed by spinning at 4°C for 20 minutes at 20,000 rpm using an SW41 rotor. The cytoplasmic layer was removed with a syringe bearing a 19-gauge needle, divided into 50 µl aliquots, and frozen in liquid nitrogen. Aliquots were stored at -80°C until use.

Preparation of demembranated sperm
Demembranated sperm were prepared as described (Sider et al., 1999Go). 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 cytokeratin—F-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. {alpha}-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., 2000Go). 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, 1991Go), and incubating on ice for 2 hours (Sider et al., 1999Go). 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., 2000Go). 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-{alpha}-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, 1997Go). 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., 1998Go).



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Fig. 3. Cytoskeleton interactions in rapidly frozen, fixed samples. Triple label confocal micrographs showing interactions between microtubules (red), F-actin (green) and cytokeratin filaments (blue). (A) Most of the cytokeratin is associated with F-actin cables (arrows). Frequently, the distal ends of microtubules were bent at points of contact with cables of F-actin and cytokeratin, and appeared to track along them (arrowheads). (B) En face view of cytoskeletal interactions and the same view at a tilt of 70°. The interaction of F-actin with cytokeratin (arrowheads) is apparent in both views. (C) The interaction of cytokeratin with F-actin was further examined by quantifying overlap (in pixel number) with and without rotating the channel representing cytokeratin 90° clockwise. The error bar for colocalization with rotation was too small to appear on this graph (P<0.05).

 


    Results
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 Summary
 Introduction
 Materials and Methods
 Results
 Discussion
 References
 
F-actin, microtubules and cytokeratin associate in unfixed extract samples
In vitro interactions of the three filament systems were first addressed using the flow chamber assay (Sider et al., 1999Go). In this approach, demembranated Xenopus sperm adhered to coverslips serve as microtubule-organizing centers when incubated with Xenopus egg extracts in flow chambers. Microtubule asters that form from the demembranated sperm can then be repeatedly washed to remove the extract and materials that are not associated with the microtubules. Astral microtubules are visualized by the inclusion of rhodamine-labeled tubulin in the extract, and F-actin is visualized by staining with Alexa-488 phalloidin. To permit visualization of cytokeratin filaments, C11, a monoclonal anti-cytokeratin antibody that recognizes all three cytokeratin isoforms in Xenopus eggs (Bartek et al., 1991Go; Staskova et al., 1991Go), was included with the phalloidin stain, and subsequently labeled with a Cy5-labeled anti-mouse antibody. This approach allowed visualization of all three cytoskeletal networks in the absence of fixation.

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 microtubule—F-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).



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Fig. 1. Cytoskeleton interactions in unfixed samples. Triple-label confocal micrograph showing interactions of microtubules (red), F-actin (green), and cytokeratin (blue) in an unfixed aster. All three systems could be found in close proximity to each other (arrows). Microtubule—F-actin associations could be observed in the absence of cytokeratin (arrowheads), and microtubule-cytokeratin interactions in the absence of F-actin (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., 1997Go; Canman and Bement, 1997Go). 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.



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Fig. 2. Perturbation of any single cytoskeletal system in unfixed samples does not prevent interaction of the remaining two. Triple label confocal micrographs showing that inhibition of F-actin (green) fails to prevent cytokeratin (blue) association with aster microtubules (red) (latrunculin, arrows); inhibition of cytokeratin polymerization does not inhibit F-actin association with aster microtubules (C11, arrows); inhibition of microtubule polymerization does not prevent interactions between F-actin and cytokeratin (nocodazole, arrows).

 

Assessment of potential cytokeratin—F-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., 1999Go). Nevertheless, occasional examples of cytokeratin—F-actin interactions could be observed in the absence of microtubules (Fig. 2, nocodazole, arrows). Thus, cytokeratin—F-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., 2000Go).

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., 1999Go; Waterman-Storer et al., 2000Go). 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., 1998Go)]. 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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Fig. 4. Perturbation of any single cytoskeletal system in rapidly frozen, fixed samples does not prevent interaction of the remaining two. Triple label confocal micrographs showing that inhibition of microtubule polymerization (nocodazole) does not prevent association of cytokeratin (blue) with F-actin (green, arrowheads). Inhibition of F-actin polymerization (latrunculin) fails to prevent cytokeratin (blue) from associating with aster microtubules (red, arrowheads). However, most of the cytokeratin is found in large aggregates on the substrate (arrow). Inhibition of cytokeratin polymerization (C11) does not inhibit F-actin association with aster microtubules (arrowheads). However, microtubules were found in small bundles on the substrate rather than as large asters. Inhibition of both microtubule and latrunculin polymerization (noc/lat) resulted in cytokeratin forming large, unorganized aggregates.

 

Cytokeratin—F-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).



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Fig. 5. Cytokeratin—F-actin interactions change over time. Double-label confocal micrographs showing F-actin (red) or cytokeratin (green) rapidly frozen at increasing time intervals following warming of extract to room temperature. Immediately after warming (0'), F-actin cables are not present and cytokeratin was found as particulates on the substrate. At 5' and 10', cytokeratin filaments are present, and invariably associated with F-actin cables. By later time points (20'), both F-actin and cytokeratin cables are thicker, and cytokeratin cables are frequently found without associated F-actin (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 {alpha}-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 {alpha}-actinin revealed that while large F-actin cables were found in both types of samples, extracts containing {alpha}-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 {alpha}-actinin for F-actin binding is disputable.



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Fig. 6. {alpha}-actinin changes the organization of both the F-actin and the cytokeratin network. Low magnification (left panels) and high magnification (right panels) double-label confocal micrographs showing F-actin (red) and cytokeratin (green) in rapidly frozen extract samples prepared after the addition of {alpha}-actinin. The exogenous {alpha}-actinin results in fine, highly crosslinked meshworks of F-actin cables in extracts that colocalize with cytokeratin. At high magnification, cytokeratin localization along F-actin cables in these networks appears punctate. The bar graph shows that the addition of exogenous {alpha}-actinin significantly decreased the mean distance between both adjacent F-actin cables and cytokeratin relative to control samples (P<0.05).

 

To quantify the effects of {alpha}-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 {alpha}-actinin-supplemented samples. The addition of {alpha}-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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Fig. 7. Cytokeratin can move in concert with and release from dynamic actin cables. 4D analysis kymographs that exhibit colocalization of cytokeratin (green) with F-actin (red) in living extract samples. (A) Cytokeratin is initially associated with F-actin, but releases and remains behind (arrowheads) after the F-actin moves away (arrows). (B) Cytokeratin remains associated with moving F-actin throughout the time of imaging (arroheads). Arrows on either side each represent elapsed time of one minute for their respective kymograph.

 


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Fig. 8. {alpha}-actinin inhibits zippering and contraction of F-actin cables in extracts. In untreated extracts, networks of F-actin zipper and contract over time (control). However, in extracts containing exogenously provided {alpha}-actinin, zippering is prevented, resulting in formation of a stable network of fine cables ({alpha}-actinin).

 


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Fig. 9. F-actin is cleared by astral microtubules in the absence of the cytokeratin network. Images of living extracts showing microtubules (MTs) and F-actin (Actin) with intact (Control) and disassembled (C11) cytokeratin networks. In both the presence and absence of cytokeratin filaments, the microtubules clear F-actin from around asters, indicating that cytokeratin is not required for the microtubule-actin interactions involved in clearing.

 

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., 2000Go), 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, {alpha}-actinin was added to extracts prior to analysis via 4D microscopy. The addition of {alpha}-actinin resulted in inhibition of the F-actin `zippering' activity typically seen in these experiments (Fig. 8). The {alpha}-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 microtubule—F-actin interactions. By adding demembranated sperm, rhodamine-labeled tubulin and Oregon Green-conjugated g-actin to extracts (Waterman-Storer et al., 2000Go), 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., 2000Go).

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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Fig. 10. F-actin is required for normal cytokeratin assembly and organization in vivo. Confocal, single label micrographs showing cytokeratin distribution in Xenopus eggs. In the absence of activation, both control (A) and latrunculin-treated (C) eggs have no apparent cytokeratin network, although cytokeratin aggregates not seen in controls were seen in latrunculin-treated, unactivated eggs. Following activation, cytokeratin assembles into a fine network in control eggs (B). Cytokeratin in activated, latrunculin-treated eggs ranges from moderately disordered, forming unusually thick cables and loops (D), to completely disordered, forming extremely large aggregates (E). F is a bar graph displaying for two experiments the difference in width of filaments and/or aggregates in control versus latrunculin-treated, activated eggs. Latrunculin treatment resulted in increased thickness, as well as overall variability. The error bars for the width of filaments in control, activated eggs were too small to appear on this graph (P<0.05).

 


    Discussion
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 Summary
 Introduction
 Materials and Methods
 Results
 Discussion
 References
 
The following findings from this report indicate that F-actin cables act as templates upon which cytokeratin filaments assemble after transition into the first mitotic interphase of Xenopus. (1) In Xenopus egg extracts that are derived from activated Xenopus eggs, cytokeratin associates with F-actin cables that spontaneously assemble in the extracts. This association is seen in both fixed and unfixed extract preparations, indicating that it is not an artifact of the fixation process. (2) Time-course experiments reveal a tight temporal correlation between the formation of cytokeratin networks and F-actin cables. Specifically, at the earliest time points where cytokeratin has assembled into cables, F-actin cables are also present and associated with the cytokeratin. (3) In extract samples where F-actin assembly is prevented, cytokeratin organization is grossly perturbed, such that cytokeratin forms large irregular aggregates on the substrate rather than networks of cables. (4) Dynamic imaging reveals that cytokeratin associated with F-actin cables moves in concert with the F-actin cables. This result is especially intriguing, in that it implies that F-actin-dependent motility could contribute to the formation of cytokeratin networks by bringing adjacent cytokeratin filaments into contact with each other. (5) Experimental alteration of the F-actin network organization by addition of {alpha}-actinin results in a corresponding change in the organization of the cytokeratin network. (6) Pharmacological disruption of the F-actin network in activated Xenopus eggs disrupts the normal pattern of cytokeratin organization.

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., 1995Go), 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., 1997Go). 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-actin—intermediate filament associations (e.g. Cary et al., 1994Go), particularly those observed in cellular regions where de novo assembly of intermediate filament structures occurs (Green et al., 1986Go; Green et al., 1987Go). 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., 1999Go). 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., 1999Go). 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., 1997Go), 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 {alpha}-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 {alpha}-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., 1999Go; Waterman-Storer et al., 2000Go) 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., 1996Go; Goldman et al., 1996Go), but also in the context of isolated cytoplasm.

The bending of microtubules at the sites of microtubule—F-actin—cytokeratin 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, 1997Go; Odde et al., 1999Go; Yvon and Wadsworth, 2000Go; Yvon et al., 2001Go) and in vitro (Sider et al., 1999Go). 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., 1998Go).

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., 2000Go; Yang et al., 1996Go). 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.


    Acknowledgments
 
We thank Clare Waterman-Storer, The Scripps Research Institute, for providing reagents, as well as the Department of Zoology, University of Wisconsin, Madison. Thanks also to members of our lab for advice and input into this project. Support for this work was provided by a grant from the National Institutes of Health (GM52932-04A2) to W.M.B. The confocal microscopy is funded by a grant from the National Science Foundation (9724515) to James Pawley.


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