1 Cancer Research UK Cell Structure Research Group, School of Life Sciences, MSI/WTB Complex, University of Dundee, Dundee, DD1 5EH, UK
2 Division of Gene Regulation and Expression, School of Life Sciences, MSI/WTB Complex, University of Dundee, Dundee, DD1 5EH, UK
3 Centre for High Resolution Imaging & Processing, School of Life Sciences, MSI/WTB Complex, University of Dundee, Dundee, DD1 5EH, UK
* Author for correspondence (e-mail: e.b.lane{at}dundee.ac.uk)
Accepted 9 July 2004
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Summary |
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Key words: Keratins, Desmosomes, Hemidesmosomes, Stress response, DM-EBS, Keratin mutations
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Introduction |
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Keratins belong to the superfamily of intermediate filament proteins. Heterodimeric filaments of type I and type II keratins are expressed in a tissue- and cell-specific manner in epithelial cells (reviewed by Fuchs and Weber, 1994). Stratified epithelia such as the epidermis express K5/K14 in the basal layer of cells and K1/K10 in the suprabasal layers. The keratins form a cytoplasmic network of filaments that are connected to the plasma membrane at complex cell-cell junctions (desmosomes) and cell-substrate junctions (hemidesmosomes). Tissue integrity in the epidermis is maintained by a complex cytoskeleton interaction between the keratin filaments and desmosomes and hemidesmosomes. Keratins are linked into desmosomes through desmoplakin and into hemidesmosomes via plectin; desmosomes are connected from cell to cell by the transmembrane cadherins (desmogleins and desmocollins), and hemidesmosomes are connected to the extracellular matrix substrate by integrins (usually
6ß4) and BP180. Other proteins found in these zones include plakoglobin within the plaques of both junctions, plakophilin in desmosomes and BP230 in hemidesmosomes. The exact nature of interactions between all these components is not yet clear (for reviews see Borradori and Sonnenberg, 1996
; Jones et al., 1998
).
The mutations that cause the severe (Dowling-Meara) type of EBS are mostly found in the highly conserved ends of the -helical rod domains of keratins K5 or K14. These regions are particularly important in filament assembly (Herrmann and Aebi, 1998
). There is a particularly mutagenic codon at residue 125 of K14, in the helix initiation motif, leading to the substitution of the arginine residue at this position (Coulombe et al., 1991
); mutations at this residue account for approximately 70% of Dowling-Meara EBS cases (Porter and Lane, 2003
). Morphological effects of most keratin mutations are hardly detectable in unstressed cultured keratinocytes but spontaneous keratin aggregates are characteristically seen in cells containing a K14-R125 mutation; this is in keeping with the theory that the helix initiation motif is crucial in normal filament assembly (Herrmann and Aebi, 1998
).
To understand exactly how keratins provide stress resilience in cells, cell lines have been generated from EBS patients with mutant keratins (Morley et al., 2003) and subjected to a variety of non-mechanical stresses (Morley et al., 1995
; D'Alessandro et al., 2002
; Morley et al., 2003
). However, a direct mechanical stress is more likely to address the question of physical resilience of keratinocytes. It would also reproduce the in vivo physiological stimulus that leads to breakdown in the epidermis of EBS sufferers. In this paper we present the results of a study of the effects of mechanical stretch on the keratin cytoskeleton in EBS keratinocytes. It was observed that repeated stretch and relaxation caused compaction of the filament network and in EBS cells led to fragmentation of keratin filaments. This was accompanied by changes in the localisation of desmosome and hemidesmosome components. We show that keratin fragmentation in response to mechanical stretch in DM-EBS cells leads to the formation of novel keratin ring structures that contain many nontransmembrane components of desmosome and hemidesmosome junctions. On the basis of these results we present a model for stretch-induced progressive fragmentation of the keratin cytoskeleton and its associated junctions in DM-EBS cells, which has implications for our understanding of cell junction turnover in normal keratinocytes.
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Materials and Methods |
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Mechanical stretch
Flexplates (Flexcell International, USA) were coated with collagen (Sigma) for 60 minutes at room temperature and then incubated for 60 minutes with 5% bovine serum albumin in DMEM supplemented with 10% FCS. Cells were seeded onto six-well Flexplates, and grown to 80% confluence. Wells in flexplates not to be stretched were isolated using FlexStops (Flexcell International, USA). At appropriate times, FlexStops were removed and stretching was resumed in well. Stretching was carried out using FX-4000TTM Cell Stretcher (Flexcell International, USA). Stretch was carried out at a frequency of 4 Hz and an amplitude of 12% for times varying up to 180 minutes at 37°C in 5% CO2. Cells were then harvested or prepared for microscopy. Previous experiments have suggested that keratinocytes are resistant to shear stress (E.B.L. and G. B. Nash, unpublished data). To determine whether fluid shear stress was likely to be significant during stretching experiments we examined the dissipation of drops of a viscous coloured liquid in tissue culture medium using the stretch parameters used in our experiments. We found that when more than 4 ml of medium was placed in flexplate wells there was very little dissipation of the viscous drop. This suggests that the effect of shear force on cells during stretching is minimal and that the observed effects are due to mechanical stretch.
Immunocytochemistry
Silicone membranes were excised from Flexplate using a clean, sharp, new scalpel for each well. Cells were fixed and permeabilised using 100% methanol-acetone (1:1) for 5 minutes at 20°C. Cells were then washed twice with PBS. Cells were then treated in blocking buffer (5.5% normal goat serum in PBS) for 1 hour at room temperature. Cells were washed as before then incubated for 1 hour at room temperature with the following primary antibodies: rabbit polyclonal antibody BL-18 (anti-K5) (dilution 1:500) (Purkis et al., 1990), monoclonal primary antibodies LL001 (anti-K14) (Purkis et al., 1990
), 11-5F (anti-desmoplakin) (dilution 1:100) (Parrish et al., 1987
), HD121 (anti-plectin) (Hieda et al., 1992
), CBL175 (anti-plakoglobin) (Cymbus Biotechnology), IE5 (anti-BP230), 233 (anti-BP180) (both gifts from K. Owaribe, University of Nagoya, Japan), PP1-5C2 (anti-plakophilin) (Cymbus Biotechnology Ltd), CD104 (anti-ß4-integrin) (Novocastra Laboratories Ltd), 6D8 [anti desmoglein 2 (Dsg2)] (gift from M. Wheelock, University of Toledo, Ohio, OH). All monoclonal antibodies were used at dilution 1:100. Cells were then washed 3x5 minutes in PBS and LL001 primary antibody was detected with FITC-conjugated sheep anti-mouse serum (F3008, Sigma, dilution 1:50). Monoclonal antibodies were detected using Texas Red conjugated anti-mouse serum (N2031, Amersham Pharmacia, UK, dilution 1:50). BL-18 antibody was detected using an Alexa fluorescein-conjugated goat anti-rabbit serum (Molecular Probes, Leiden, Netherlands; dilution 1:400). All antibodies were diluted in DMEM supplemented with 10% FCS and incubations were carried out in the dark.
Fluorescence microscopy
Silicone membranes were mounted onto glass slides using CitiFluor (Sigma Aldrich, UK), air-dried and visualised with an Axiovert 200M system (Carl Zeiss). For higher resolution microscopy, images were acquired using a DeltaVision microscopy system (Applied Precision, USA), using a Nikon PlanApo100x/1.4 N.A. objective lens and a Roper Scientific Interline cooled CCD camera (5 MHz MicroMax1300YHS) using the standard DeltaVision filter set. Optical sectioning was performed at 0.2 µm intervals to encompass the entire cell. Binning was set at 1x1 to give an effective pixel size of 0.067 mm. Three-dimensional (3D) data sets were deconvolved using the SoftWorRx application (Applied Precision, USA). MediaRecorder software (Silicon Graphics Ind., USA) was used to generate TIFF files from either images of single optical sections or from 3D maximum-intensity volumetric projections (generated using SoftWoRx). TIFFS were processed for publication using Adobe Photoshop.
Transmission electron microscopy
KEB-7 cells stretched for 120 minutes were fixed using Karnovsky's fixative (5% glutaraldehyde, 4% paraformaldehyde) containing 1% osmium tetroxide, for 30 minutes. Cells were rinsed three times with phosphate buffer. Cells were then dehydrated through an ethanol series, ending with two 10-minute incubations in 100% ethanol. Cells were incubated overnight in a 100% ethanol/Durcupan (Sigma) solution (1:1). This was then replaced three times with undiluted resin over a 24-hour period. The silicone membrane was then excised and seven BEEM cups (Agar Scientific, Essex, UK) filled with resin were placed upside down onto membrane. Resin was then polymerised at 60°C for 48 hours. The silicone membrane was then peeled off leaving cells attached to resin. 70 nm sections were cut using an Ultracut UCT ultramicrotome (Leica, Vienna, Austria) using a diamond knife (Diatome, Biel, Switzerland). Sections were collected on hexagonal 100 mesh copper grids (Agar Scientific, Essex, UK) then stained with 3% uranyl acetate for 10 minutes followed by lead citrate for 10 minutes. Images were obtained using a JEOL 1200 EX transmission electron microscope (JEOL, Tokyo, Japan) equipped with a Fuji FDL5000 digital plate reader. Images were scanned using a high-resolution imaging plate scanner (DITABIS, Pforzheim, Germany) and processed for publication using Adobe Photoshop.
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Results |
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After 30 minutes of cyclical stretch, NEB-1 cells started to exhibit thickening of filaments (Fig. 1C) and compaction of bundles became increasingly evident. KEB-7 cells exhibited a dramatic increase in this phenomenon at this time (Fig. 1D). KEB-7 cells showed increased amounts of filament fragmentation, again, particularly along free edges. The keratin network appeared progressively denser around the nucleus, as the whole network retracted from the cell edge. After 120 minutes stretch there was a pronounced collapse of the network in approximately 80% of KEB-7 cells (Fig. 1F). The keratin then mainly exhibited a perinuclear localisation, but there was still a large amount of fragmentation and keratin rings were present at the cell edge. Control NEB-1 cells showed dramatic bundling of filaments at this time point (Fig. 1E) but the network was still well dispersed throughout the cytoplasm. These cells did not exhibit the network collapse that was seen in KEB-7 cells.
The position of a cell within a colony of cells appeared to have an effect on the extent that the keratin network collapses. KEB-7 cells appeared to be afforded some protection from the effects of stretch when they were in the middle of a group of cells, as cells on the edge of a colony showed faster and more dramatic effects than those in the middle. Cells positioned internally, with no free edges, nevertheless did exhibit fragmentation of the keratin network but this was only evident after longer exposure to mechanical stretch (between 1 and 2 hours).
The pattern of keratin fragmentation in KEB-7 cells was uneven around the cells suggesting that uneven forces or uneven response to force was operating in different parts of the cell. Fig. 2 shows that by 10 minutes of oscillating stretch, keratin fragmentation was particularly pronounced in regions where three cells meet and/or at free edges of cells, i.e. areas where there was a dearth of desmosomes and few tonofilament bundles at right-angles to the cell membrane. Fragmentation was also observed in desmosomal areas in response to short periods of stretch. However, the frequency of this was much less than fragmentation observed in areas lacking desmosomes. This suggests that the presence of desmosomes somehow provides the keratin network with an element of resistance to mechanical stretch. As mechanical stress was applied, keratin fragmentation was seen first in these regions. After only 10 minutes of stretch, two distinct types of keratin structures were observed. Solid aggregates of keratin (Fig. 2, inset, arrow) were detected as would be expected in Dowling-Meara EBS keratinocytes. Second, novel ring-like keratin structures were also observed, easily distinguishable from the solid aggregates (Fig. 2, inset, asterisk). These hollow structures have not been previously reported.
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No effect of stretch on microtubules was observed, either in control or DM-EBS cells before or during stretch. As the microtubule network is highly dynamic it is possible that recovery could have occurred between cessation of stretch and fixation. We were aware of this possibility and endeavoured to keep this time period to a minimum, less than 90 seconds. We also examined the effect of mechanical stretch on the actin cytoskeleton: all cells examined only showed typical epithelial patterns of actin distribution and there was no fragmentation of actin during the cyclical stretch. The formation of actin stress fibres in response to stretch has been previously reported (Shirinsky et al., 1989) and is particularly evident in response to unilateral stretching (Shirinsky et al., 1989
; Sugimoto et al., 1991
; Takemasa et al., 1998
). Radial stretching was used here, which may not lead to the formation of actin stress fibres and may explain the absence of stress fibres in our experiments. We conclude that cyclical stretch as imposed here has no major redistributing effect on the actin cytoskeleton. This suggests that it is unlikely that actin has a direct role in the observed keratin fragmentation.
Effect of mechanical stretch on desmosome morphology
Because keratin fragmentation always starts from the cell edge, we examined the point where keratin filaments attach to the cell membrane, the desmosome. Within the desmosome, desmoplakin is thought to form a direct link between the keratin network and the desmosome plaque (Green et al., 1990). Both NEB-1 and KEB-7 cells make abundant desmosomes and before the application of stretch, desmoplakin staining was similar in both (Fig. 3A,B). As the time of stretch was increased, changes in desmoplakin staining were seen. After 120 minutes of stretch, desmoplakin distribution in NEB-1 cells was unchanged but the shape of the desmoplakin patches at the cell-cell contacts became elongated, suggesting some elasticity in these desmosome cell-cell junctions (Fig. 3C).
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At this same time point of 120 minutes, the desmoplakin localisation in mutant keratin cells was highly abnormal. Desmoplakin staining was very dispersed in regions where keratin fragmentation was high (Fig. 3D), but remained localised to desmosomes in regions where the keratin network was well maintained on at least one cell edge (Fig. 3D, arrow). This suggests that an intact keratin network is needed to maintain desmoplakin at desmosomes. Furthermore, keratin attached to desmosomes appeared to have some initial resistance to fragmentation, and the desmoplakin staining suggests some elasticity at the desmosome during stretch. We measured desmoplakin staining in wild-type keratinocytes before and after 10 minutes of stretch, taking 40 measurements for each condition. To try and exclude the possibility of desmoplakin recruitment to desmosomes during stretch we only measured desmoplakin staining at the junction where we could see no desmoplakin on keratin filaments close to the junction. It was observed that in response to short periods of stretch, the average length of a desmoplakin staining patch increased approximately threefold (Fig. 3E). This supports the hypothesis that there is some intrinsic elasticity of desmosomes involving desmoplakin.
In these subconfluent cultured keratinocytes, hemidesmosome protein distribution is often virtually reciprocal to desmosome protein distribution, as was seen with antibodies to BP180. The hemidesmosome protein was more sparse in desmosome-rich areas but present in larger amounts where desmosomes were few (e.g. the apex between three cells; see Fig. 4A, arrows). Thus, BP180 localisation also correlates with areas of the cell where keratin fragmentation is seen earliest after stretch. If desmosomes between cells have elastic properties that provide damping of mechanical stress and initially stop filaments breaking, the situation at hemidesmosomes may be different. If there is no elastic capacity in the junction because it is linked to a less pliable surface, such as the dermis or even worse, a plastic culture dish it follows that there would be less capacity to absorb force. Thus, breakage of mutated keratin filaments would probably be initiated here.
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To test this further, KEB-7 cells were stained for either desmoplakin (desmosomes), BP180 (hemidesmosomes) or plectin (hemidesmosomes) and stretched for 10 minutes. One hundred cells were selected at random and scored for colocalisation between keratin aggregates and the above proteins. The only prerequisite for selecting cells was that they exhibited keratin aggregation. Cell scoring was repeated three times so that 300 cells were scored for each protein. The results were plotted to show the average number of cells per 100 showing colocalisation (Fig. 4B). This showed that in response to short periods of stretch an average of only 19/100 cells showed colocalisation between desmoplakin and aggregates. For the hemidesmosomal proteins this was greatly increased; 75/100 for BP180 and 77/100 for plectin. This supports the idea that keratin aggregation begins in areas of the cell rich in hemidesmosomes and that some intrinsic desmosomal elasticity may provide initial resistance to keratin fragmentation in response to stretch.
Stretch-induced ring structures contain keratins plus junction proteins
The altered location of desmoplakin during stretch was examined using deconvolution microscopy. After 120 minutes of stretch, desmoplakin can be seen to associate with small fragments of keratin at points of cell-cell contact (Fig. 5A). Looking at the cell from the cell edge towards the nucleus it is clear that the number of small keratin fragments decreases and the number of ring structures increases (Fig. 5A). This suggests that small keratin fragments are precursors of ring structures. It can be also be seen that desmoplakin is colocalised to these keratin ring structures (Fig. 5A, arrow). The colocalisation of desmoplakin and keratin rings can be seen more clearly in Fig. 6B.
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To determine whether the cells were literally being pulled apart, cells were stretched for 120 minutes and stained for desmoglein 2 (Fig. 5B). Desmoglein 2 is one of the specialised desmosome cadherins, a major desmosomal transmembrane protein in cultured epidermal cells that is involved in maintaining the desmosome connection between two cells. Complete fragmentation of the keratin network within a cell did not lead to relocalisation of Dsg2, and there was no association of Dsg2 with keratin rings. The cells were also incubated, before stretch, with a lipophilic membrane dye (DiI, Molecular Probes, Leiden, Netherlands) known to stain internal and external membrane structures. After stretch, no changes in membrane staining were observed and there was no colocalisation of the lipophilic dye with keratin rings (results not shown). Together, these observations confirmed that the cell membranes at desmosomal junctions do not separate during stretch and that keratin rings are not associated with any membrane, as would be the case if they were contained within membrane vesicles.
We examined the localisation of the hemidesmosome protein plectin during stretch. Before stretch, plectin was seen predominantly around the distal areas of the cells, again in regions where one would predict from the above results that keratin fragmentation is initiated. After 120 minutes stretch, plectin was significantly redistributed (Fig. 5C): diffuse staining was seen at the cell periphery and plectin was seen throughout the cell. Plectin was colocalised with fragmented keratin at the cell periphery and could also be seen associated with keratin rings. The images in Fig. 5 were generated as a volume projection and could not confirm that desmoplakin and plectin are directly associated with keratin rings. Single optical slices (Fig. 6) revealed that small keratin fragments were directly associated with desmoplakin (Fig. 6A). Desmoplakin was also directly associated with keratin rings (Fig. 6B). Staining shows that desmoplakin was contained within the ring, giving rings a studded appearance. As desmoplakin associates directly with small fragments of keratin, we predict that these small fragments join together by linkage of keratin and desmoplakin, resulting in a ring structure.
Plectin was also directly associated with keratin rings (Fig. 6C), but unlike desmoplakin staining the plectin appeared to be peripheral, and not integral, to the rings. Chains of rings can be seen in stretched KEB-7 cells and plectin was commonly detected between the rings. This suggests that plectin may be involved in the lateral adhesion of keratin rings. This would be in keeping with its properties as a linker protein with the ability to cross-link keratin filaments (reviewed by Steinbock and Wiche, 1999).
The location of other desmosomal and hemidesmosomal proteins was investigated for association with keratin rings. We found that BP180 (Fig. 6D), plakoglobin (Fig. 6E), BP230 (Fig. 6F) and plakophilin (Fig. 6G) all associate with keratin rings, and staining suggests that these proteins too may interact mainly with the surfaces of keratin rings, unlike the integral interaction of desmoplakin. BP180 is a transmembrane component of the basal membrane of keratinocytes that has been localised to hemidesmosomes and shown to be important for the stabilisation of the hemidesmosome (Hopkinson et al., 1998; Hopkinson and Jones, 2000
).
In contrast to the cytoplasmic hemidesmosome components and similar to the desmosome membrane cadherins, staining for the hemidesmosome integrin ß4 subunit failed to show any localisation with keratin rings (Fig. 6H,I). This suggests a similar picture as seen for desmosomes: on disruption of the keratin filaments, the transmembrane ß4 integrin maintains its connection with the extracellular matrix and stays at the plasma membrane, as the desmosomal cadherins also stay at the membrane in desmosomes, while all the cytoplasmic proteins are relocalised during stretch and interact with fragmented keratin. BP180 was the only transmembrane protein of the junctions to become associated with keratin rings, suggesting that its association with the plasma membrane is less strong than that of the integrins and cadherins. We examined whether both desmosomal and hemidesmosomal proteins were present in the same rings. However, it was difficult to resolve these images to gain useful information. While our observations (not shown) suggest they do co-exist within rings it was difficult to prove this conclusively.
Keratin rings are identifiable by transmission electron microscopy
Transmission electron microscopy of cells stretched for 2 hours revealed structures similar to those seen by immunofluorescence. Rings and whorls of keratin bundles were observed as well as small fragments of keratin (Fig. 7A). More solid aggregates of keratin close to the basal membrane of the cells were observed within stretched cells (Fig. 7B,C), which may be homologous to the aggregates previously observed in EBS keratinocytes. Examination of single keratin rings revealed that they are often angular, as if composed of stiffer segments of greater electron density, which could reflect the areas associated with junctional proteins (Fig. 7D-F). This would fit with the small fragments being precursors of keratin rings. Keratin rings were observed with regions dipping out of the plane of section confirming that keratin rings are not hollow cylindrical structures or hollow spheres of keratin that appear as rings after sectioning.
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The keratin filament network of DM-EBS keratinocytes recovers rapidly
After 2 hours of stretch cycles, wild-type and mutant cells were released and allowed to recover, and then fixed and examined at various time points. Even the mutant keratin filament network recovered quickly. After only 2.5 minutes, the mutant cells showed bands of keratin running parallel with the cell edge (Fig. 8A, arrows); these bands were still present after 10 minutes recovery (Fig. 8B, arrows). After 30 minutes (Fig. 8C), a sparse network was already rebuilt; this network continued to develop and after 1 hour, cells appeared fully recovered (Fig. 8D,E). By 24 hours after cessation of mechanical stress, the cells appeared to have a network almost indistinguishable from that of the wild-type cells before stretch (Fig. 8F). Also, there were very few cells containing the low-level aggregates normally seen in mutant keratin keratinocytes. There was also little evidence of the bundling of keratin filaments that we observed in these cells before stretch. Mechanical stretching of both wild-type (NEB-1) and DM-EBS (KEB-7) cells resulted in increased levels of apoptosis (D. Russell, unpublished). Both cell lines exhibited similar levels of apoptosis before stretch, and stretch resulted in increased levels of apoptosis in both cell lines. Interestingly, KEB-7 cells show delayed induction of apoptosis during stretch and show reduced numbers of apoptotic cells during recovery compared with NEB-1 cells (D. Russell, unpublished).
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We examined cells after 10 minutes recovery from stretch to attempt to determine the nature of the keratin band frequently observed early in recovery (Fig. 8G). This band is usually found close to the cell edge and the filaments leading to this band from the nuclear side are very straight. On closer examination, this band of keratin has a diffuse appearance (Fig. 8H), not consistent with normal keratin filament staining. In areas of cells where this diffuse band of keratin is present, keratin rings were not observed on the nuclear side of the band. Keratin rings could be seen within the band with many giving the appearance of `opening up' within the band. The images suggest that keratin rings are resorbed into this band of keratin, which may provide the site at which keratin fragments are `rescued' and remodelled into a filament network. Desmoplakin staining (as well as other junction proteins, not shown), was concentrated within this band of keratin, presumably brought there with the keratin rings.
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Discussion |
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DM-EBS keratin filaments fragment on mechanical stress
First, the demonstration that Dowling-Meara mutant keratins do indeed fragment as a result of mechanical stress would support the model that keratins provide internal reinforcement for cells and that the pathogenic mutations lead to filament network breakdown, hence compromising the reinforcing capacity of the keratin cytoskeleton. Cells expressing the K14-R125 mutation, but not cells with wild-type keratin, show fragmentation of the keratin network on mechanical stress in this experimental situation. Mutations at this residue are the most common type found in Dowling-Meara EBS and account for approximately 70% of all DM-EBS cases investigated to date worldwide (Porter and Lane, 2003). What is not directly seen in these experiments of course is the final cytolytic step, which in the patients' epidermis is what actually leads to epidermal blisters. Monolayer cultures of keratinocytes are widely held to mimic basal layer cells in the epidermis, but in situ these cells would be attached to the overlying layers of stratified keratinocytes, directly linked by desmosomes to the epibasal layer and thence indirectly coupled to the higher suprabasal layers, culminating in the hardened layers of the stratum corneum. These layers would provide another dimension to the physical stress acting on the basal cells, which if acting in opposition to lateral displacement in the basal layer could be the final force component that leads to cell lysis. In situ, mechanical stress to the skin would usually involve stress from displacement of the overlying cells and not just shear or compression against the underlying basal lamina and the neighbouring basal cells. In this experimental situation therefore we are only able to examine part of the stress pattern of skin, but this does make it somewhat easier to analyse the consequences of this stretch.
Stretch model allows dissection of the breakdown process
Second, the assay enables the pathogenic process of cytoskeleton breakdown to begin to be analysed. Short periods of cell stretching induce clear deformation of the keratin network irregularly around the cell periphery. The earliest stages examined of this deformation show the appearance of small fragments and aggregates at the cell periphery, to varying degrees in different areas, plus the formation of novel keratin filament rings further into the cell body. Aggregates and ring structures can be seen by electron microscopy to have different internal structures. Ring structures consist of loops or rings of small bundles of keratin filaments, often with an angularity that indicates patches of differential stiffness in the ring, possibly where desmosomal or hemidesmosomal filament-associated proteins are retained. Short fragments of keratin filament bundles are also seen in early stages or in the extreme periphery of the cell and the ring structures probably arise by the annealing of sticky ends of sheared keratin bundles or tonofilaments; keratin filaments are known to assemble extremely rapidly (reviewed by Strelkov et al., 2003).
The keratin rings are usually observed at the distal margins of the filament network; the small aggregates are observed further out than this. Electron-dense aggregates are one of the diagnostic hallmarks of Dowling-Meara EBS and aggregates can occasionally be seen in these KEB-7 cells in a resting state without subjecting the cells to any additional stress, in agreement with observations of Fuchs and colleagues (Coulombe et al., 1991; Fuchs and Weber, 1994
). The nature of these aggregates is still unclear and undetermined, but they have been shown to contain both K5 and K14. However, the aggregates generated de novo by stretching in these experiments were noticeably smaller than the spontaneous aggregates seen in biopsies of patients' skin, or spontaneously in cultures of KEB-7 cells, suggesting that they might increase in size with time. This is reinforced by our observation that following recovery of KEB-7 cells from stretch-induced keratin disruption, the newly redistributed keratin initially shows no sign of aggregate formation.
A role for keratin filament tension in maintaining desmosomes
We have observed that in the EBS cells used here with mutant keratin K14 (R125P), the initiation of breakage of the keratin filaments is followed by progressive disassembly of desmosomes and hemidesmosomes. The cytoplasmic proteins of the junctions gradually become relocalised and associate with keratin fragments in the cytoplasm, which form characteristic rings, presumably because breakage of the filaments leaves `sticky' ends. A simple model for this is presented in Fig. 9. Our observations suggest that the disassembly of junctions is initiated by the loss of tension in the filament network. Observations from other pathological conditions may support this idea. A recessive mutation in desmoplakin that disrupts the desmoplakin-intermediate filament interaction has been identified that results in cardiomyopathy with large intercellular spaces, suggesting that intermediate filament attachment is important for maintaining desmosomal junctions (Norgett et al., 2000). Such tension would also be dissipated (1) on extensive phosphorylation of keratins (as occurs during mitosis in lamins and probably also in keratins, judging by the formation of keratin aggregates during mitosis) (Lane et al., 1982
; Horwitz et al., 1981
; Chou and Omary, 1994
; Liao et al., 1997
; Toivola et al., 2002
) or (2) on breaking of the epithelial barrier (as in epidermal wounding). Both these circumstances call for cytoskeleton remodelling, cell shape change and repositioning of cells with respect to their neighbours in the epithelium. In the first case this would be necessary to accommodate the new daughter cell, and in the second case (more dramatically), to change the shape of the epithelial sheet and mobilize cells for epithelial migration and wound healing. In both cases, and in other physiological situations that can be imagined, disassembly of keratin-subtended junctions on loss of tension in the cell would be a strategy that would make disassembly of unwanted junctions a fast and automatic process.
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Desmoplakin as an elastic component of desmosomes
We have shown that keratin aggregation is initiated in areas of cells where there are fewer desmosomes. Desmosomes may offer some initial resistance to fragmentation of keratin attached to desmosomes. Stretching of wild-type keratinocytes revealed that after stretch, desmoplakin staining showed an elongated appearance. Membrane staining and staining for transmembrane components of desmosomes showed that there is no cell-cell separation in response to stretch. Mapping of desmosomal components has suggested that desmoplakin may be folded or coiled at desmosomes (North et al., 1999). Our results suggest that in response to stretch, desmoplakin may unfold or result in elongation of coiled desmoplakin, implying some elasticity across the desmosome. The increasing length of patches of desmoplakin staining observed during mechanical stretch supports this idea. It is easy to imagine that this would be physiologically advantageous. Elasticity across the desmosome would enable a force applied to the epidermis to be absorbed across the tissue and would ultimately reduce the force experienced by a single cell. Desmosomes are also abundant in heart tissue and desmosome elasticity could reduce the stress on a single cell during the contraction/relaxation cycle of the heart.
Mechanical stretch is certain to have more to teach us about the function of intermediate filaments in tissues. The availability of this stretch assay, which reproduces at least part of the pathology of EBS in a tissue culture situation, will be useful for analysing the disease process and any hypothetical measures for ameliorating symptoms of this and related disorders. These experiments stress the need for much further analysis of the role of mechanical forces in regulating biological responses at the cellular level.
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Acknowledgments |
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References |
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