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Correspondence to Gerhard Wiche: gerhard.wiche{at}univie.ac.at
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were significantly elevated, and increased migration rates, as assessed by in vitro wound-closure assays and time-lapse microscopy, were observed. Forced expression of RACK1, which is the plectin-binding receptor protein for activated PKC, in wild-type keratinocytes elevated their migration potential close to that of plectin-null cells. These data establish a link between cytolinker-controlled cytoarchitecture/scaffolding functions of keratin IFs and specific MAP kinase cascades mediating distinct cellular responses.
Abbreviations used in this paper: EBS, epidermolysis bullosa simplex; HD, hemidesmosome; IF, intermediate filament; OA, okadaic acid; RACK, receptor for activated C kinase; SB, Sorensen's buffer.
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50 amino acid residues linking two (5 and 6) of its six C-terminal repeat domains. At its N terminus, plectin harbors a functional actin-binding domain that also serves as an integrin ß4- and an additional vimentin-binding site (Sevcik et al., 2004). Plectin's association with microtubules and their cross-linking to vimentin IFs has been previously reported (Svitkina et al., 1996); however, the molecular mechanism of this interaction is still unknown. There is increasing evidence that, apart from acting as a cytoskeletal linker protein, plectin serves an important function as a scaffolding platform of proteins involved in cellular signaling. Strong support for this idea comes from the recent identification of several novel interaction partners of plectin with links to signaling, such as the nonreceptor tyrosine kinase Fer and AMP kinase, and the observation that plectin deficiency affects their enzymatic activities (Lunter and Wiche, 2002; Gregor et al., 2006). The scaffolding function of plectin was recently confirmed when it was shown that the protein binds and sequesters the receptor for activated C kinase 1 (RACK1) to the cytoskeleton, and thereby affects PKC signaling pathways (Osmanagic-Myers and Wiche, 2004). This, together with previous studies showing that plectin is involved in the regulation of actin filament dynamics and influences Rho/Rac/cdc42 signaling (Andrä et al., 1998), suggests that plectin, and likely cytolinkers in general, provide a crucial link between cytoskeleton dynamics and signaling machineries.
The prominent localization of plectin at hemidesmosomes (HDs), desmosomes, Z-line structures and dense plaques of striated and smooth muscle, intercalated discs of cardiac muscle, and focal contacts implied a role for the protein in linking the cytoskeleton to plasma membrane junctional complexes (Wiche, 1998). This was supported by studies showing that defects in plectin expression lead to the skin disease epidermolysis bullosa simplex (EBS). In these patients, as well as in plectin-deficient mice generated by targeted gene inactivation (Andrä et al., 1997), the link of the keratin cytoskeleton to HDs was dramatically affected (McMillan et al., 1998), with keratin filaments appearing less tightly bundled at their insertion into the inner plate structure of HDs. Moreover, earlier ultrastructural studies and in vitro reconstitution of IFs had shown that plectin was primarily located at branching points of IFs (Foisner et al., 1988). However, the important question of whether plectin, indeed, affects IF network cytoarchitecture and its dynamics has not been addressed so far, except for a study showing that a recombinant fragment containing plectin's C-terminal IF-binding site inhibits IF formation in vitro in a dose-dependent manner (Steinböck et al., 2000). Investigating what impact plectin deficiency has on the organization and dynamic properties of IFs in keratinocytes, we found a direct link between plectin-controlled IF cytoarchitecture and MAP kinase signaling cascades involved in cell migration and stress response.
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60% confluence), ple (–/–) cells exhibited significantly enlarged mesh size of keratin networks at their periphery compared with wild-type cells (Fig. 1, compare B with E and H with K, along with their corresponding magnifications). A quantitative analysis of subconfluent cell populations revealed that >70% of ple (–/–), but only <8% of ple (+/+), keratinocytes displayed such a phenotype. Regarding expression levels of different keratins, no differences were observed between ple (+/+) and (–/–) keratinocytes, as revealed by immunoblotting analysis (Fig. S1, available at http://www.jcb.org/cgi/content/full/jcb.200605172/DC1). Interestingly, the vimentin filament network of plectin-deficient fibroblasts showed similar characteristics. It appeared more bundled and less delicate compared with wild-type cells, creating a network with wider spaces between the filaments (unpublished data).
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6 clusters were formed in both cell types (Fig. 1, O and R), and in ple (+/+) cells, partial colocalization with plectin was observed (Fig. S2, A–C, available at http://www.jcb.org/cgi/content/full/jcb.200605172/DC1). In addition, keratin staining partially overlapped with the ring formed by integrin clusters (Fig. S2, D–F). Transmission electron microscopy revealed a prominent lateral bundling of keratin filaments in peripheral regions of ple (–/–) cells, resulting in larger spaces between individual and bundled filaments (Fig. 2 A, b and d). In some regions, filaments showed a complete lateral collapse, appearing as one massive filamentous bundle (Fig. 2 A, b, arrow). Bundling to such an extent was never observed in ple (+/+) cells, where corresponding areas were dominated by numerous fine, and apparently short, filamentous structures, with orthogonal cross-bridges filling the space between the filament bundles (Fig. 2 A, a and c).
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Keratin filaments of ple (–/–) keratinocytes are more sensitive to hypoosmotic shock
Osmotic and heat-shock assays serve as useful tools in monitoring keratin network properties and their alterations in EBS caused by keratin mutations (Morley et al., 1995; D'Alessandro et al., 2002). Therefore, we examined whether the altered keratin filament cytoarchitecture of ple (–/–) keratinocytes affected their response to changes in osmolarity and temperature. Upon urea treatment, INT6-positive retraction fibers, which are a characteristic feature of migrating and mitotic cells (Geuijen and Sonnenberg, 2002), became very pronounced at the rear of both cell types (Fig. 3 A, a, c, d, f, h, and i).
Most probably, the general shrinkage of the cells, which started shortly after exposure to urea (D'Alessandro et al., 2002), triggered the elevated formation of these structures. INT6-positive retraction fibers of ple (–/–) cells, however, were significantly longer than those of ple (+/+) cells, possibly reflecting a stronger shrinkage of plectin-deficient cells (Fig. 3 A, compare d and i). In accordance with recent studies showing increased urea-induced bundling of keratin filaments (Werner et al., 2004), the IF network appeared less filamentous after urea treatment and accumulated around the cell center, indicating that it had collapsed onto the nuclei (Fig. 3 A, b and g). Interestingly, at the leading edge of cells, which is an area devoid of retraction fibers, keratin bundles of ple (+/+) cells displayed a regular form and were closely associated with peripheral integrin clusters, whereas in ple (–/–) cells filament bundles were conspicuously tangled and distant from the cell periphery (Fig. 3 A, compare e and j). The appearance of such "torn" filaments in ple (–/–) cells suggested a reduction in their INT6ß4 anchorage. This would be consistent with the reduced attachment of keratin filaments to HDs reported for EBS patients (McMillan et al., 1998).
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6ß4 content using antibodies to INTß4. As expected, such cell fractions were highly enriched in keratins 5 (unpublished data) and 14 (Fig. 3 B) and, in the case of wild-type cells, contained considerable amounts of plectin. INTß4 was, however, completely absent from cytokeratin fractions of ple (–/–) cells. The absence of INTß4 from the cytokeratin fraction of ple (–/–) keratinocytes correlated well with the reduced number of HD-like structures found in these cells (Fig. 1, compare O with R; Andrä et al., 2003).
Collectively, these data supported the notion that the increased susceptibility of ple (–/–) keratinocytes to urea-induced deformation (as revealed by the conspicuous tangling of filaments and dramatic increase in the length of INT6ß4 retraction fibers), was caused by a lack of filament attachment to integrin clusters. On the other hand, the response of ple (+/+) and (–/–) cells to elevated temperatures in heat-shock assays was very similar, leading to a partial granulation of keratin filaments in both cases (unpublished data).
Plectin regulates IF disassembly dynamics
If, in the absence of plectin, the rigidity of IF networks is reduced and filaments are more loosely bound, or not bound at all, to the outer membrane, one might expect the network to be disassembled more readily in ple (–/–) compared with ple (+/+) cells. To test this, we monitored by immunofluorescence microscopy the kinetics of IF disassembly upon treatment of ple (+/+) and (–/–) cells with the serine/threonine phosphatase inhibitor okadaic acid (OA), which is known to selectively cause the disruption of IFs (Strnad et al., 2001). After a 2-h treatment of keratinocytes, the well-spread keratin network had formed thick bundles of filaments that seemed to be retracting toward the nucleus (Fig. 4 A, a and d, arrowheads).
At later time points (4 and 6 h), a progressive breakdown of filaments was observed, with keratin granules forming first at the cell periphery (Fig. 4 A, e, arrowheads), followed by their collapse into a dense perinuclear ring (Fig. 4 A, e, arrow) that eventually became fragmented into numerous granules of various sizes (Fig. 4 A, f, arrow). The initial bundling of filaments appeared to occur more efficiently in ple (–/–) cells, as indicated by the increased mesh size of keratin networks visualized in the majority of these cells compared with ple (+/+) cells (Fig. 4 A, a and d, arrows). Moreover, at the 4- and 6-h time points, the proportions of cells with keratin granules were significantly higher in ple (–/–) compared with ple (+/+) cells (Fig. 4 A, b and c and e and f). For a statistical analysis of keratin filament disassembly, cells were classified into three categories (1–3), where category 1 represented cells with no granules, category 2 represented cells with granules and residual filaments (Fig. 4 A, e, cell marked with asterisk), and category 3 represented cells in which complete granulation, including that of the perinuclear ring structure, had occurred (Fig. 4 A, f, cell marked with arrow). At the 2-h time point, 15% of ple (–/–) cells already fell into category 2, whereas only a mere 4% of ple (+/+) cells did (Fig. 4 B). After 4 h, the majority of the ple (–/–) cell population had their keratin network either partially (30%) or completely (50%) disassembled, whereas 70% of ple (+/+) cells were still without any granules (Fig. 4 B). After 6 h, the difference between ple (+/+) and (–/–) cells became less pronounced, but 30% of ple (+/+) cells still did not show any granules, versus only 7% in the case of ple (–/–) cells (Fig. 4 B). Based on this, we concluded that the disassembly of IFs upon OA treatment is significantly accelerated in ple (–/–) compared with wild-type keratinocytes. In agreement with a previous study (Strnad et al., 2001), even after 6 h of OA treatment, no disruption of microtubules or microfilaments was observed. In fact, actin stress fiber formation was found to be significantly increased in ple (–/–) cells, and to some extent, also in ple (+/+) cells (unpublished data).
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Monitoring detergent-soluble keratin pools during OA treatment of keratinocytes, we found the level of soluble keratin proteins to already be elevated by approximately twofold in plectin-negative compared with wild-type cells before drug treatment (Fig. 4 C, 0 h). This difference further increased to approximately threefold within 2 h of drug treatment. Thereafter, keratin solubility in ple (–/–) cells stayed about level, while that in ple (+/+) cells further increased, approaching a level similar of that of ple (–/–) cells (Fig. 4 C, 4 h).
Plectin deficiency affects stress-activated p38 and Erk1/2 MAP kinases
Previous studies have shown stress-activated p38 MAP kinase to be one of the major candidates for mediating the effects of OA on vimentin and keratins (Cheng and Lai, 1998; Toivola et al., 2002). Therefore, we examined whether the OA-induced changes in network appearance and solubility of keratins were paralleled by changes in p38 activity. Using anti–phospho-p38 antibodies to monitor the activation status of p38 kinase, we found no significant differences between ple (+/+) and (–/–) keratinocytes under basal conditions (Fig. 5, A and B).
Upon addition of OA to ple (–/–) cells, a moderate increase in p38 activity during the first hour was followed by a steep increase during the second hour and a sharp decline thereafter. In contrast, p38 kinase activity levels in ple (+/+) keratinocytes showed only a modest increase during the first 2 h, staying constant thereafter. Thus, the maximum of p38 activity measured in plectin-negative cells was more than twice as high as in wild-type cells.
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Faster in vitro migration of plectin-deficient compared with wild-type keratinocytes
Plectin-mediated attachment of the keratin cytoskeleton to INT6ß4 has been shown to play a crucial role in stabilizing adhesion of keratinocytes to the matrix, thereby inhibiting cell migration (Geuijen and Sonnenberg, 2002). Plectin-deficient keratinocytes, showing no association of INT6ß4 with keratins (Fig. 3 B), together with their up-regulation of Erk1/2 (see previous section), which is a kinase that positively regulates keratinocyte migration (Huang et al., 2004), prompted us to compare the migratory potentials of ple (+/+) and (–/–) keratinocytes using an in vitro wound-healing assay. Average migration distances measured for ple (–/–) cells were almost twice as long as those of ple (+/+) cells (Fig. 6, A and B).
Interestingly, the mesh size of the keratin network in ple (–/–) keratinocytes along the wound edge was much larger compared with that of cells at a distance from the wound and, in these regions, differences to the keratin network of ple (+/+) cells became most prominent (Fig. 6 D, compare c with g). This was consistent with our finding that an increased keratin network mesh-size characteristic of ple (–/–) keratinocytes was particularly evident in subconfluent cell cultures (Fig. 1). Furthermore, in migrating wound-edge keratinocytes, plectin's localization changed from basal integrin cluster– to keratin filament–associated (Fig. 6 D, compare a with e), highlighting the importance of plectin in organizing keratin cytoarchitecture during cell migration.
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Importantly, although MEK1/2 inhibition decreased the migration rate of ple (–/–) keratinocytes, it had no effect on their aberrant keratin network organization (Fig. 6 D, d and h), clearly placing plectin in the MAP kinase cascade upstream of Erk1/2. These data established a causal relationship between plectin deficiency and accelerated migration of keratinocytes, showing hyperactivation of Erk1/2 to be a consequence of plectin deficiency.
PKC and c-Src both have been suggested as major players in signaling pathways responsible for migration of keratinocytes (Yamada et al., 2000; Li et al., 2002), and both have been shown to be upstream activators of Erk1/2 (Miranti et al., 1999; Gagnoux-Palacios et al., 2003). Therefore, we next investigated activation of these kinases in membrane and cytosolic fractions of ple (+/+) and (–/–) keratinocytes. As shown in Fig. 7, A and B, both, PKC and c-Src kinase, exhibited increased phosphorylation (corresponding to higher activities) in the membrane fraction of ple (–/–) keratinocytes compared with wild-type cells.
Although total c-Src levels in membrane and cytosolic fractions from both cell types were comparable, those of PKC were lower in the membrane fraction of ple (–/–) compared with ple (+/+) cells. In the cytosolic fractions, total PKC signals were hardly detectable in any of the two cell types.
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Forced expression of the plectin-binding protein RACK1 in keratinocytes leads to increased motility
In a previous study, we revealed a role of plectin as a cytoskeletal regulator of PKC signaling and possibly other signaling events (Osmanagic-Myers and Wiche, 2004). We proposed that plectin sequesters RACK1, which is a receptor and scaffolding protein of activated PKC and a direct binding partner of plectin, to the cytoskeleton when PKC is inactive. Because of the lack of its cytoskeletal docking site in the absence of plectin, in plectin-deficient cells RACK1 accumulates (together with PKC) at the periphery of cells, similar to the situation in wild-type cells after activation of PKC. According to this model, one may expect that the forced expression of RACK1 in wild-type keratinocytes mimics the situation in ple (–/–) cells, leading to their characteristic phenotypes. To test this we analyzed the migration potential of keratinocytes expressing an EGFP-RACK1 fusion protein using time-lapse video microscopy.
In accordance with migration distances measured in scratch wound closure assays (Fig. 6), ple (–/–) keratinocytes displayed a migration velocity (1.58 µm/min) approximately two times as high as that of ple (+/+) cells (0.82 µm/min), when observed 2–6 h after plating (not depicted). As shown in Fig. 7 D (controls), 14–18 h after plating, ple (–/–) cells migrated three times as fast as ple (+/+) cells (1.53 vs. 0.49 µm/min). Expression of EGFP-RACK1 led to an increase in the average migration rates of both cell types. Transfected wild-type cells (1.30 µm/min) migrated 2.6 times faster than untransfected control cells, reaching 85% of the speed of untransfected ple (–/–) cells, whereas transfected ple (–/–) cells (2.02 µm/min) migrated 1.3 times faster than their untransfected counterparts. Similar experiments were performed with the cytoplasmic nonreceptor tyrosine kinase Fer, which, like RACK1, directly binds to plectin and thereby is inhibited in its activity (Lunter and Wiche, 2002). In this case the speed of wild-type cells was increased by approximately twofold (unpublished data).
In contrast, expression of an EGFP-plectin isoform 1a (full-length) fusion protein in ple (–/–) cells led to a significant slowdown of the cells, reducing their average speed to 0.98 µm/min (Fig. 7 D). This was equivalent to a slightly >50% rescue potential of the fusion protein, taking the values of control ple (+/+) and (–/–) cells into account. The lower rescue potential of plectin 1a in this assay compared with restoration of keratin network cytoarchitecture (Fig. 2 B) may reflect the requirement of other major isoforms expressed in keratinocytes, such as plectin 1c and 1 (Andrä et al., 2003), for full phenotype restoration. Thus, whereas overexpression of plectin-controlled signaling proteins, such as RACK1, led to downstream mechanisms boosting cell motility (Fig. 7), reexpression of a major plectin isoform in ple (–/–) cells led to the partial reversal of their aberrant migration.
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Based on ultrastructural analysis, we suggest that the mechanism behind the observed phenotype is a reduction of orthogonal cross-linkages between individual keratin IFs in the absence of plectin. In cross-linking individual filaments at high angles, plectin's interaction with IFs would resemble that of filamin with microfilaments. Such a mode of action would be consistent with earlier studies showing plectin to be predominantly localized at crossover and branching points of IFs (Foisner et al. 1988). Similar to filamin, plectin was shown to exist in dimeric and tetrameric states. Tetrameric structures are assumed to be formed by antiparallel alignment of two parallel plectin dimers (Wiche, 1998). Thus, exposed C-terminal IF-binding sites on both ends of such structure have the ability to cross-link two filaments. Even individual parallel dimers may have a cross-linking capacity, as an additional N-terminal vimentin-binding site resides in the N-terminal actin-binding domain of plectin (Sevcik et al., 2004). Oligomers of plectin tetramers, which are generated by the head-to-head fusion of dumbbell-shaped plectin molecules (Foisner and Wiche, 1987), have been shown to form at IF branching points (Foisner et al. 1988). Cross-linking functions of plectin have also been clearly demonstrated by Svitkina et al. (1996), who used immunogold electron microscopy to show that plectin is organized in millipede-like structures around the core of individual IFs, with plectin sidearms frequently making Y contacts with each other. Conceivably, such Y-shaped structures may emerge from head-to-head fusion of plectin molecules, allowing cross-linking at high angles. Moreover, these authors reported that plectin was not localized regularly all along IFs, but was more concentrated at their distal ends, which is consistent with our finding that defects in IF organization were most prominent in the peripheral regions of ple (–/–) cells.
The urea-based osmotic shock assay revealed a stronger stress response of ple (–/–) versus (+/+) keratinocytes, reflected by increased cell shrinkage and considerably longer retraction fibers. Suggesting an increased plasticity of ple (–/–) cells, these data also implied a role of plectin in stabilizing membrane surfaces. By interlinking IFs into properly organized three-dimensional networks, and by connecting these to the plasma membrane, plectin probably provides cell membranes with the required resistance against deformations, such as those induced by osmotic shock. A similar model has been proposed by Flanagan et al. (2001) to explain the increased deformability of filamin-deficient melanoma cells. Thus, plectin and filamin could have similar modes of action in respect to different cytoskeletal filament systems.
What is the cause of accelerated IF network disassembly in ple (–/–) cells?
We assume that plectin increases the stiffness of IFs by introducing orthogonal cross-links between filaments and, thus, acts as a stabilizer opposing their disassembly. Moreover, the protein acts as a linker, anchoring keratin filaments to hemidesmosomal INTß4, as shown on ultrastructural and biochemical levels (Fig. 4 B; Rezniczek et al., 1998). Further support for an IF-stabilizing role of plectin stems from differences in ple (+/+) and (–/–) keratinocytes during the early stages of OA-induced disassembly of IFs, which in wild-type cells correlates with the dissociation of plectin from keratin IF networks. The events after OA treatment can be viewed to parallel those of a more physiological process, i.e., mitosis, as in both cases plectin dissociates from IFs during their disassembly (Foisner et al., 1996). Thus, it seems that the release of stabilizing proteins such as plectin is a requirement for the efficient disassembly of IFs.
However, if the faster IF disassembly in ple (–/–) cells was strictly caused by the diminished mechanical stability of IFs, why did we observe a higher increase in p38 activity in these cells after OA-induced IF disassembly? It is unlikely that faster IF disassembly was caused by elevated levels of p38 kinase activity because, in this case, increased phosphorylation of keratins in ple (–/–) compared with wild-type cells should have been detected using p38 kinase site-specific phosphokeratin antibodies (Toivola et al., 2002). This, however, was not the case (unpublished data). Furthermore, there were no differences in the activity levels of p38 kinase between wild-type and ple (–/–) cells under basal conditions. Recently, an association of simple epithelial keratins 8/18 with Raf kinase and its disruption after treatment of cells with OA has been reported (Ku et al., 2004). Based on this, the authors suggested a role of keratins in sequestering a population of Raf and thereby regulating its signaling potential. In a similar fashion, keratins might regulate either p38 directly or one of its upstream effectors. Thus, we speculate that it is the faster disassembly of IFs and larger pool of their soluble subunits proteins in ple (–/–) compared with ple (+/+) cells that, via an unknown positive feedback mechanism, affects the activity of p38, rather than the other way around.
The plectin-ERK MAP kinase link and its consequences for keratinocyte migration
It is the currently accepted view that ligation of integrins triggers the activation of Erk, via either the adaptor protein Shc or some other mechanism (Wary et al., 1998). This has been shown to be the case for integrins of fibroblasts, as well as the integrin specific for keratinocytes, 6ß4 (Mainiero et al., 1997). In some studies, the importance of integrin linkage to the cytoskeleton has been emphasized. For instance, in fibroblasts and skeletal muscle cells, it was shown that the disruption of microfilaments with cytochalasin D blocked the integrin-mediated activation of MAP kinases (Zhu and Assoian, 1995). Similarly, a mutation in INTß4 that prevents plectin–INTß4 interaction leads to accelerated migration of keratinocytes (Geuijen and Sonnenberg, 2002). Fully in line with these studies, we demonstrate that plectin-deficient keratinocytes, showing no association of INT6ß4 with keratins, have an elevated migration potential. Most importantly, the migration of plectin-deficient cells was significantly reduced when plectin was reexpressed. An involvement in migration has recently been reported for another cytolinker family member, kakapo/short-stop (Fuss et al., 2004). By regulating Notch receptor localization and activity, short-stop was shown to be essential for the movement of proventricular cells during the invagination of foregut epithelium into endodermal midgut layers.
Our study shows that the increased migration potential of ple (–/–) compared with ple (+/+) keratinocytes is directly linked to elevated states of Erk kinase phosphorylation. Erk has been implicated in the migration of numerous cell types (Huang et al., 2004), and it has been shown to be the sole kinase responsible for ECM-initiated migration of keratinocytes (Li et al., 2004). On the other hand, enhancement and directionality of growth factor signaling is mediated by both Erk and p38 kinases, whereas JNKs were reported to be uninvolved in keratinocyte motility (Li et al., 2004). By pharmacological inhibition of Erk's upstream kinases MEK1/2, we were able to restore the aberrant high migratory potential of ple (–/–) keratinocytes to normal levels, but were unable to rescue the abnormal keratin network organization of these cells. Therefore, we feel it is safe to conclude that hyperactivation of Erk1/2 is a result of keratin network alterations caused by plectin deficiency, rather than the opposite. In support of this, keratinocytes from EBS patients, with mutations in keratins leading to spontaneous formation of keratin aggregates, migrate significantly faster in comparison to control cells (Morley et al., 2003). Although Erk activities were not investigated in this study, elevated basal levels of the stress-activated kinase SAPK/JNK found in these cells (D'Alessandro et al., 2002) were implicated in their faster migration. This raises the intriguing question of whether distinct alterations in keratin network organization, such as aggregation in keratin-related EBS versus bundling in EBS caused by plectin deficiency, may lead to the up-regulation of distinct signaling pathways, such as SAPK/JNK versus MAPK. Studying human keratinocytes from an EBS-MD patient, Kurose et al. (2000) reported unaltered migration using phagokinetic track measurements. As these cells very likely expressed rodless isoforms of plectin, contrary to the plectin-null cells used in our study, it is difficult to compare both studies.
The signaling pathway leading from plectin-related keratin network alterations to hyperactivation of Erk1/2 still remains elusive. Our analysis conducted so far shows that the activities of two key proteins known to be involved in the regulation of keratinocyte migration, c-Src, and PKC are up-regulated in the membrane fraction of plectin-deficient cells, and that PP2-inhibition of c-Src indeed down-regulates Erk1/2 activities. Thus, membrane-associated c-Src and PKC are likely candidates for mediators of signals from plectin to Erk1/2. For the transduction of signals from the IF network to the membrane, our live-cell imaging data of transfected migrating keratinocytes, expressing EGFP-RACK1 fusion proteins, offer a plausible mechanistic explanation. Similar to a model proposed for PKC regulation through plectin-sequestration of RACK1 on IFs of fibroblasts (Osmanagic-Myers and Wiche, 2004), we propose that in keratinocytes regulatory (trigger) proteins of PKC, c-Src, and/or other upstream affectors of Erk1/2, are sequestered on IF-associated plectin molecules in a wild-type scenario, but are unbound to IFs and have free access to the membrane because of their missing anchor in plectin-deficient cells. The faster migration of cells overexpressing RACK1, which is shown in this study, is consistent with such a model. It will be of interest to characterize in more detail on the molecular level how c-Src and PKC become activated through signaling proteins such as RACK1 (and possibly Fer kinase) and what consequences this might have in different cell types. Fig. 8 A shows a scheme depicting the model proposed.
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Antibodies
For immunoblotting, the following primary and secondary antibodies were used: anti-K5 and -K6 antisera (PRB-160P and PRB-169P, respectively; Covance), mAbs LL001 to K14 (provided by J.M. Leigh, Royal London School of Medicine and Dentistry, London, England; Morley et al., 1995), mAbs LP34 (DakoCytomation) to K5, K6, and K18 (pan-keratin), a mixture of mAbs to K18 and K8 (Ks 18.04 and Ks 8.7, respectively; Progen), anti-plectin antiserum #9 (Andrä et al., 2003), anti-INTß4 antiserum (provided by F.G. Giancotti, Memorial-Sloane Kettering Cancer Center, New York, NY; Mainiero et al., 1997), affinity-purified goat anti-vimentin antiserum (provided by P. Traub, University of Bonn, Bonn, Germany), mouse mAbs sc-535 to p38 (Santa Cruz Biotechnology, Inc.), rabbit mAbs 3D7 to phospho-Thr180/Tyr182 p38 (Cell Signaling Technology), mAb D-2 to Erk2 (Santa Cruz Biotechnology, Inc.), mAbs E-4 to phospho-Tyr204 Erk1/2 (Santa Cruz Biotechnology, Inc.), anti–c-Src antiserum (Santa Cruz Biotechnology, Inc.), anti-phospho Y418 Src antiserum (Biozol), anti-phospho Thr505 PKC antiserum (Cell Signaling Technology), mAb P36520 to PKC (BD Biosciences), anti-caveolin antiserum (BD Biosciences), goat anti–rabbit IgG, goat anti–mouse IgG, and donkey anti–goat IgG (all from Jackson ImmunoResearch Laboratories), all conjugated to horseradish peroxidase. For immunofluorescence microscopy the following primary antibodies were used: anti-plectin antiserum #46 (Andrä et al., 2003), pan-keratin (see above), anti-K5 antiserum (see above), and rat mAbs to INT6 (CD49f; BD Biosciences), mAbs B-5-1-2 to -tubulin (Sigma-Aldrich), and affinity-purified antiserum and mAbs to actin (A 2066 and AC-40, respectively; Sigma-Aldrich). As secondary antibodies, we used goat anti–rabbit IgG Alexa Fluor 488 (Invitrogen), goat anti–rat IgG Texas red (Accurate Chemical & Scientific Corporation), goat anti–mouse IgG Texas red, donkey anti–mouse Rhodamine red-X, and donkey anti–goat Cy2 (all from Jackson ImmunoResearch Laboratories).
Immunofluorescence and electron microscopy
Cells grown overnight (12 h) were methanol-fixed, washed with PBS, mounted in Mowiol, and viewed in a laser-scanning microscope (LSM 510; Carl Zeiss MicroImaging, Inc.) at room temperature. Images were visualized with either a Plan-Apochromat 63x, 1.4 NA, or a Plan-Apochromat 100x, 1.4 NA, objective lens (Carl Zeiss MicroImaging, Inc.) using LSM software and processed using the Photoshop CS2 (Adobe) software package. For electron microscopy, cells grown on glass coverslips were washed three times with 0.15 M Sorensen's buffer (SB), pH 7.4, before a 1-h fixation in 3% glutaraldehyde in SB. Cells were then washed twice with SB and postfixed in 1% OsO4 in SB for 30 min. Subsequently, they were dehydrated in ethanol and flat-embedded in epoxy resin (Agar 100). Glass coverslips were removed from the Epon block by immersion in liquid nitrogen and subsequent warming. Thin sections (60–80 nm) were cut parallel to the plane of the cell layer, using an ultramicrotome (Leica). They were then mounted on copper grids, contrasted by uranyl acetate and lead citrate, and viewed at 60 kV in an electron microscope (JEM-1210; JEOL).
Preparation of cell fractions
After 12 h of adhesion, keratinocytes were washed twice with PBS and lysed directly with 50 mM Tris-HCl, pH 6.8, 100 mM DTT, 2% SDS, 1 mM Na2VO3, 1x phosphatase inhibitor cocktail 1 (Sigma-Aldrich), 1% bromophenol blue, and 10% glycerol (sample buffer). Aliquots of cell lysates containing equal amounts of total proteins were separated by SDS-PAGE and, after immunoblotting using peroxidase-coupled secondary antibodies, protein bands were visualized by exposure to x-ray film. Quantitation of bands was performed as previously described (Osmanagic-Myers and Wiche, 2004). Triton X-100 or high-salt extract fractions of keratinocytes were prepared according to Toivola et al. (2002). Membrane and cytosolic fractions were prepared according to a digitonin-based extraction protocol (Osmanagic-Myers and Wiche, 2004).
Scratch wound assay
ple (+/+)/p53 (–/–) and ple (–/–)/p53 (–/–) basal mouse keratinocytes were grown in parallel until reaching confluence (48 h). They were then treated with 10 µg/ml mitomycin C for 2 h. Subsequently, a scratch wound was introduced into the monolayer using a yellow Gilson pipette tip. Cells were washed three times with growth medium and further incubated for 24 h. For MEK1/2 or c-Src inhibition, 30 µM PD98059 (Cell Signaling Technology) or PP2 (Calbiochem), as indicated, were added to the growth medium 1 h before scratching and throughout the scratch closure period. Before fixation, a reference wound was inflicted to determine the original wound size. Cells were then fixed with methanol and processed for immunofluorescence microscopy using mAbs to actin, anti-plectin #46, and anti–pan-keratins. The average migration distance was calculated by subtracting the wound width after 24 h migration from that of the reference wound.
Live-cell imaging
Time-lapse video microscopy was implemented on a microscope (Axiovert S100TV; Carl Zeiss MicroImaging, Inc) equipped with phase-contrast and epiillumination optics. Cells were spread on collagen I–coated coverslips at a density of 2.8 x 105 cells/cm2 and kept in KGM during the whole period of observation. Migration was monitored in a closed POCmini cultivation system (Carl Zeiss MicroImaging, Inc) at 37°C and 5% CO2. Recordings of migrations started 14 h after plating, and frames were taken with a 10x lens in 1-min intervals over a period of 90 min. Images were obtained using a back-illuminated, cooled charge-coupled device camera (Princeton Research Instruments) driven by a 16-bit controller. The whole video microscopy system was automated by Metamorph 6.3 (Universal Imaging Corporation). The length of trajectories of migrating cells was measured by tracking the central nuclei. Cells that did not spread or remain in the field of view during the whole period of observation were not taken into account for measuring. For statistical evaluation, we used 20–30 cells per construct and genotype.
Online supplemental material
Fig. S1 shows expression levels of keratins in wild-type and plectin-deficient cells monitored by immunoblotting. Fig. S2 shows that plectin and keratins colocalize with INT6 at the cell periphery. Fig. S3 shows the dissociation of plectin from keratin filaments upon treatment of cells with OA. Online supplemental material is available at http://www.jcb.org/cgi/content/full/jcb.200605172/DC1.
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Acknowledgments |
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This work was supported by grant P17862-B09 from the Austrian Science Research Fund.
Submitted: 26 May 2006
Accepted: 11 July 2006
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References |
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, P < 0.001. Bars: (A) 1 µm; (B) 10 µm.
