Article |
Address correspondence to Pierre A. Coulombe, Dept. of Biological Chemistry, The Johns Hopkins University School of Medicine, 725 North Wolfe St., Baltimore, MD 21205. Tel.: (410) 614-0510. Fax: (410) 614-7567. email: coulombe{at}jhmi.edu
![]() |
Abstract |
---|
![]() ![]() ![]() ![]() ![]() ![]() ![]() |
---|
Key Words: mouse skin grafting; keratin; skin; injury; wound healing; migration
![]() |
Introduction |
---|
![]() ![]() ![]() ![]() ![]() ![]() ![]() |
---|
With its 50 members partitioned into two sequence types (Hesse et al., 2001), the epithelial-restricted keratins provide a unique handle to explore the significance of the multiplicity and differential regulation of IF genes. Type I (K9-K23; Ha1-Ha9) and type II (K1-K8; Hb1-Hb6) keratin genes are regulated in a pairwise fashion, reflecting a heteropolymerization requirement (Fuchs and Weber, 1994). Most pairs of keratin genes are regulated in a differentiation-specific fashion in epithelia (Moll et al., 1982). For instance, in interfollicular epidermis, progenitor basal cells express K5/K14 as their main pair of type II/type I keratins, whereas early differentiating keratinocytes express the K1/K10 pair. This blueprint varies depending on regional differences in epidermis, on disease, or environmental challenges (McGowan and Coulombe, 1998a). This is the case after epidermal injury, which triggers the induction of K6 isoforms and type I K16 and K17 in keratinocytes undergoing activation at the wound edge. This transcriptional event occurs at the expense of K1 and K10, and correlates with striking alterations in the morphology and other properties of keratinocytes. Expression of K6, K16, and K17 persists as wound-activated keratinocytes migrate into the site of injury, but is reversible upon wound closure (Mansbridge and Knapp, 1987; Paladini et al., 1996; Takahashi et al., 1998).
Along with their partners K16 and K17, K6 paralogues, of which there are many in mammalian genomes (Tyner and Fuchs, 1986; Ramirez et al., 1995; Takahashi et al., 1995, 1998), exhibit a complex regulation that includes constitutive expression in specific compartments within all epithelial appendages (McGowan and Coulombe, 1998a). This has complicated the assessment of their role during adult wound repair; for instance, mice null for the two functional K6 genes, keratin 6 and 6ß (K6
/K6ß), die rapidly after birth owing to the fragility of stress-bearing epithelia within the oral mucosa (Wong et al., 2000; Wojcik et al., 2001). Here, we exploit an ex vivo skin explant culture assay along with in vivo skin tissue grafting to analyze the response of K6
/K6ß null to various forms of challenges including injury. We show that K6
/K6ß and K16 profoundly impact the potential of keratinocytes for wound epithelialization, and are essential to the maintenance of keratinocyte integrity in activated epidermis. We propose a model in which these keratins function to provide a resilient cytoskeletal scaffold able to maintain adequate structural integrity while providing sufficient pliability for effective migration into the wound site. This hypothesis may also explain in part the diversity and differentiation-related distribution of IF proteins.
![]() |
Results |
---|
![]() ![]() ![]() ![]() ![]() ![]() ![]() |
---|
|
Enhanced outgrowth of K6/K6ß-null keratinocytes results from migration and is accompanied by changes in keratin and F-actin organization, and in p120ctn
Re-epithelialization of skin wounds in vivo results from increases in mitotic activity and migration of keratinocytes located at the wound margins (Martin, 1997). Likewise, over a period of 8 d in culture, migration and mitosis equally contribute to the outgrowth of keratinocytes from skin explants (Wawersik et al., 2001). We addressed the role of mitotis and migration in null explants using a three-prong approach. First, explants were treated with the nucleotide analogue BrdU 2 h before harvest to visualize cells engaged in DNA replication. The density of BrdU-positive keratinocytes at the explant tissue edge, where they are most abundant, was similar in wild-type and null explants (Fig. 2, A and B). Second, analysis of proteins from explant outgrowths revealed no change in levels of proliferating cell nuclear antigen or myc relative to actin (Fig. 2 C). Third, the outgrowth of keratinocytes was assessed in two complementary ways after treatment with the irreversible mitosis inhibitor mitomycin C (Wawersik et al., 2001). In the first experiment, explants were treated with mitomycin C at 24 h after seeding, cultured for 8 d, and processed for analysis. Relative to their wild-type controls, treated and untreated K6/K6ß-null explants show a comparable enhancement in outgrowth area (Fig. 1 D). In the second experiment, treatment with mitomycin C was performed at 48 h after seeding and the outgrowth distance calculated after treatment, and at days 4, 6, and 8 thereafter, for the same set of live explants. Under such conditions, K6
/K6ß-null keratinocytes migrated further than wild-type or hemizygous keratinocytes (Fig. 2 E). Collectively, these analyses suggest that the increased epithelialization potential manifested by K6
/K6ß-null samples in skin explant culture results mainly from enhanced migration rather than increased mitosis.
|
|
Influence of strain background affects the behavior of skin explant cultures
The genetic background can influence the consequences of keratin-null mutations in mice. Depending on the strain background, K8-null mice may exhibit colorectal hyperplasia or embryonic lethality (Baribault et al., 1993, 1994), whereas K17-null mice may exhibit striking alopecia or not (McGowan et al., 2002). K6/K6ß-null mice are able to survive their severe oral lesions in a specific strain background (compare Wong et al., 2000, with Wojcik et al., 2001). The latter finding is relevant here, considering that Wojcik et al. (2001) also did not find evidence of altered wound repair in the skin of their K6
/K6ß-null mice.
The K6/K6ß-null allele was backcrossed into the C57Bl/6 and 129/SvJ inbred strains for at least seven generations, and the epithelialization potential was evaluated in skin explant culture. Wild-type explants derived from inbred and mixed background strains showed the same amount of outgrowth suggesting equal epithelialization potential (Fig. 4). In the C57Bl/6 strain, hemizygous and homozygous null K6
/K6ß-null keratinocytes exhibit an epithelialization potential similar to the mixed genetic background (Fig. 4). However, in the 129/SvJ strain, K6
/K6ß-null keratinocytes exhibit a substantially larger (3.1-fold) increase in epithelialization potential compared with wild type (Fig. 4). In this instance, hemizygous null explants display a statistically significant increase in outgrowth compared with wild type. Therefore, genetic background influences the extent to which the epithelialization potential is enhanced in K6
/K6ß-null keratinocytes. A gene dosage effect occurs in the 129/SvJ strain, indicating that the number of K6-encoding alleles is important in at least some settings, as predicted by Takahashi et al. (1998).
|
|
We examined wild-type and K6/K6ß-null grafted skin in an alternative setting in which the expression of K6
/K6ß is induced. Grafted backskins were treated with PMA, a substance known to induce hyperproliferation and K6
/K6ß expression (Takahashi et al., 1998) without abrogating terminal differentiation (Bernot et al., 2002). Significant thickening of the epidermis occurred in the suprabasal compartment of PMA-treated wild-type and K6
/K6ß-null grafted backskin (Fig. 6, A and D), providing further evidence that loss of K6
/K6ß does not alter the mitotic potential of keratinocytes (Fig. 2). The resilience of the thickened epidermis was tested via gentle rubbing of PMA-treated skin with a pencil eraser (10 measured strokes) followed by histological analysis. In the specific case of K6
/K6ß-null skin grafts, the stress test had to be interrupted because of severe loss of integrity. Upon histological examination of these samples, cytolysis and intra-epidermal cleavage had occurred in the suprabasal layers of epidermis (Fig. 6, E and F). Wild-type (Fig. 6, B and C) and hemizygous skin grafts (not depicted) did not show any such fragility or cytolysis under these conditions. These studies demonstrate the role of K6
/K6ß as a crucial element responsible for maintaining keratinocyte integrity in challenged skin tissue, and in migrating keratinocytes found at sites of tissue injury.
|
![]() |
Discussion |
---|
![]() ![]() ![]() ![]() ![]() ![]() ![]() |
---|
Significance of altered keratin expression during wound epithelialization in skin
The type II K6/K6ß and type I K16 and K17 keratin proteins are related, respectively, in primary structure to K5 and K14, which are constitutively expressed in basal keratinocytes of the skin (Lloyd et al., 1995). Accumulation of K6
/K6ß, K16, and K17 at the wound edge runs concurrently with the down-regulation of K1 and K10 (Mansbridge and Knapp, 1987; Paladini et al., 1996), whose expression normally parallels differentiation in the suprabasal layers of epidermis. This inductive response is evolutionary conserved (Estrada et al., 1993) and occurs in a number of other complex epithelia, including the oral mucosa and cornea (Schermer et al., 1989; Takahashi and Coulombe, 1997). The enhanced migratory potential exhibited by K6
/K6ß-null skin keratinocytes, thus, appears counterintuitive considering that evolution has selected for the involvement of K6 proteins after injury. The relevance of our findings is supported by studies conducted in transgenic mice overexpressing K16 protein. In this instance, a delay is observed in the keratinocyte outgrowth produced during skin explant culture. Such mice also exhibit a delay in the closure of skin wounds in vivo (Wawersik et al., 2001).
Mixed results have been obtained in previous studies involving K6-null mouse models. On the one hand, Wojcik et al. (2000) reported that loss of K6 causes a delay in the epithelialization of partial thickness skin wounds in vivo. In such wounds, keratinocytes originating from the hair follicle outer root sheath participate to epithelialization (Pang et al., 1978). On the other hand, the response of skin tissue to full-thickness injury was not altered in the K6
/K6ß-null animals able to survive in the genetic background used (Wojcik et al., 2001). In this instance, the skin response to partial thickness injury was not assessed. Differences in both strain backgrounds and assays likely contribute to the discrepancy between those observations and ours. It could be that the same factors enabling some K6
/K6ß double-null mice to survive the life-threatening lesions affecting the oral mucosa (for review see Wojcik et al., 2001) are able to compensate during wound repair. K6hf, which is related to K5, K6
, and K6ß, is not a contributing factor given that it is not expressed at the edge of skin wounds (Wojcik et al., 2001; Wang et al., 2003). With these issues notwithstanding, much of the evidence stemming from transgenic mouse models, alongside the evolutionarily conserved regulation of K6, K16, and K17 at the wound edge, argue strongly for an important and specific role provided by these keratins during wound epithelialization.
What is the nature of this role? We propose that the induction of K6, K16, and K17 in keratinocytes located proximal to the wound edge imparts them with mechanical properties that are intermediate between those exhibited by K5/K14-expressing keratinocytes in the basal layer and by K1/K10-expressing cells in the suprabasal layers. We postulate that expression of the K5/K14 pair as primary keratins renders keratinocytes relatively more pliable and, thus, better able to undergo directed cell migration, a notion that is directly supported by our explant culture findings. However, given a lack of K6/K6ß and reduced levels of K16, keratinocytes endowed mainly with K5/K14/K17 (Fig. 1 D) are insufficiently resilient to survive the harshness of the wound site. Rather, they rupture intracellularly in a fashion similar to that seen in keratin-based fragility disorders, as shown by our skin grafting studies. Conversely, abundant expression of the K1/K10 pair in the context of normal epidermal differentiation is expected to promote stronger mechanical resilience at the expense of cellular pliability. In support of this view, differentiating keratinocytes transit more rapidly through the suprabasal compartment of epidermis in K10-null mice (Reichelt and Magin, 2002). The induction of K6
/K6ß, K16, and K17 after injury is, thus, likely to reflect a compromise between conflicting needs: retaining enough cellular pliability for migration while acquiring sufficient resilience to survive the wound environment. It is conceivable that a similar role would apply in situations involving constitutive expression of K6, K16, and K17 (e.g., epithelial appendages, palmar/plantar epidermis, and select internal epithelial tissues), although in these instances a specific degree of cellular pliability may be required for purposes other than migration (for review see Swensson et al., 1998; Bernot et al., 2002).
Although the proposed model invokes the major function shared by all types of IF polymers, it does not preclude additional roles for IF proteins during wound repair. For instance, K8/K18, as well as K17, can each promote the survival of epithelial cells under specific circumstances (Caulin et al., 2000; Gilbert et al., 2001; McGowan et al., 2002). Our hypothesis also does not preclude an important role for other keratinocyte constituents during skin wound repair (Martin, 1997). This hypothesis can be generalized to account for the rapid response exhibited by other IF genes after tissue injury, and also may help justify the diversity encountered within the IF protein superfamily. This model makes the key prediction that differential regulation of keratin genes enables epithelial cells to adjust their viscoelastic properties in order to best carry out the demands placed upon them. Whereas there is evidence that type I-type II pairing significantly influences the mechanical properties of keratin filament suspensions (Hofmann and Franke, 1997; Bousquet et al., 2001; Yamada et al., 2002), it is doubtful that such in vitro studies provide a complete account of IF properties in living cells. This said, there is as yet no direct evidence addressing the issue of whether and how IFs influence the cell's viscoelastic properties. Application of particle tracking rheology (Yamada et al., 2000) to genetically modified keratinocytes in culture offers an opportunity to examine this issue. In addition to IF protein composition, key parameters such as filament concentration, organization, posttranslational modifications, and dynamics should all influence how IFs impact on the properties of the cell, viscoelastic and otherwise (Fuchs and Cleveland, 1998; Omary et al., 1998; Coulombe et al., 2000).
Understanding adaptive changes and modifier gene effects in keratinocytes null for K6/K6ß
The increased epithelialization potential exhibited by K6/K6ß-null keratinocytes in the idealized setting of skin explant culture offers an opportunity to identify key effectors of this process. That the keratinocyte cytoskeleton is profoundly affected by the absence of K6 proteins is reflected by alterations in F-actin organization, phosphotyrosine epitopes, and p120ctn. Of relevance to our findings, adenovirus-mediated expression of p120ctn increases the motility of mouse skin keratinocytes in primary culture, and alters F-actin organization (Cozzolino et al., 2003). Changes in F-actin have been observed in fibroblasts carrying null alleles for plectin, an IF-interacting protein, or vimentin, a type III IF protein, correlating with altered migratory properties (Andra et al., 1998; Eckes et al., 2000). Keratin filament and F-actin networks are differentially affected in suprabasal null keratinocytes located at the explant edge and in matrix-attached keratinocytes the leading edge of the outgrowth. This may reflect the occurrence of distinct adaptative responses depending on the immediate surroundings of the cells, or alternatively, the existence of cellular heterogeneity within the epithelial outgrowth. Although the finding of increased stress fibers in leading edge keratinocytes may be perceived to be at odds with an enhanced migratory phenotype, we have no information about the dynamic properties of F-actin in these cells, or whether enhanced outgrowth in null explants is mediated by matrix-attached keratinocytes or by suprabasal keratinocytes.
Participation of IFs to tissue repair: a universal phenomenon with common significance?
Alterations in IF gene expression represent a conserved response after tissue injury. Such changes are manifested as either a switch in the IF genes being expressed or a stimulation of the existing program. Akin to epidermal keratinocytes, neurons respond to injury with the induction of -internexin concomitant with down-regulation of the type IV neurofilament proteins (Goldstein et al., 1988, Tetzlaff et al., 1988; Muma et al., 1990; McGraw et al., 2002). In addition to its effect on neurons, trauma to the central nervous system (CNS) also triggers a reaction in neuroglia. Surrounding wound-activated astrocytes become mobilized into the site of tissue injury (Galou et al., 1996; Pekny et al., 1999). These cells elevate their expression of type III glial fibrillary acidic protein (GFAP) and reactivate the expression of two other IF proteins, vimentin (type III) and nestin (type VI), which had been expressed as part of their development (Galou et al., 1996; Kaya et al., 1999; Shibuya et al., 2002). In addition, neurofilament-L and -M subunits are transiently expressed in Schwann cells deprived of contacts with injured neurons (Fabrizi et al., 1997). In skeletal muscle, the predominant type III IF protein desmin is quickly down-regulated after injury, whereas vimentin and nestin expression is reactivated (Vaittinen et al., 2001). In simple epithelia such as liver and pancreas, injury results in a rapid and robust stimulation of type I/II IF genes already expressed, K8/K18 (Loranger et al., 1997; Ku et al., 1998; Toivola et al., 2000). And finally, the expression of K6
/K6ß, K16, and K17 is either induced or stimulated whenever complex epithelia other than skin, such as the cornea and oral mucosa, are injured (Schermer et al., 1989; Takahashi and Coulombe, 1997).
Inactivation of various types of IF genes in mice compromises the response to injury. Regenerating myelinated neurons from neurofilament-Lnull mice exhibit retarded maturation after injury (Zhu et al., 1997). GFAP-null mice show depressed long term potentiation and increased neuronal death subsequent to transient CNS ischemia (Tanaka et al., 2002). Mice lacking both GFAP and vimentin (type III) display defects in glial scar formation after injury to the CNS (Pekny et al., 1999). In primary culture, GFAP/vimentin double-null astrocytes exhibit impaired motility (Lepekhin et al., 2001). Vimentin-null mice, embryonic and adult, are delayed in their ability to heal skin wounds owing to altered mesenchymal contraction and delayed migration of fibroblasts to the wound site, respectively (Eckes et al., 2000). Also, vimentin-null fibroblasts exhibit fragility and lysis when subjected to distending forces. Yet, vimentin-null fibroblasts are competent to migrate in ex vivo culture and exhibit altered actin networks (Eckes et al., 1998). Desmin-null skeletal muscle exhibit defects in stress bearing, proliferation, and myoblast fusion after injury (Li et al., 1997; Smythe et al., 2001). K8-null mice are more likely to die after hepatectomy and other forms of acute challenges to the liver, correlating with their unusual fragility (Loranger et al., 1997; Ku et al., 1998). Although multiple mechanisms could account for these phenotypes, these observations are also consistent with a role of IFs toward promoting an optimal balance between properties of cellular pliability and resilience. Follow-up studies testing this concept will provide insight into the basic mechanisms of tissue repair, and may lead to improved strategies for the therapeutic management of chronic wounds.
![]() |
Materials and methods |
---|
![]() ![]() ![]() ![]() ![]() ![]() ![]() |
---|
Skin explant culture
Ex vivo explant culture of 24-d-old mouse skin was performed as described previously (Mazzalupo et al., 2002). Using 4-mm punches (Acuderm, Inc.), circular skin biopsies were obtained and plated with medium in 24-well dishes (Mazzalupo et al., 2002). Explants were cultured for 8 d before fixing (3% PFA for 10 min; 100% methanol for 5 min; room temperature). Immunostaining for K17 (McGowan and Coulombe, 1998b) was used to identify keratinocytes in the cellular outgrowth. Keratinocyte outgrowths were quantitated using two methods yielding similar findings. One consisted of measuring the surface area covered by the K17-positive outgrowth using the MacBas v2.5 software. The other method measured the linear distance extending between the explant biopsy and the distal edge of the cellular outgrowth by randomly applying a pattern of eight radial lines to the K17-stained preparations (Wawersik et al., 2001). Where warranted, stratified statistical analysis was performed as a function of genotype. A subset of explants were treated with mitomycin C (10 µg/ml for 2 h; Sigma-Aldrich) at 48 h after seeding and keratinocyte outgrowth was measured immediately after treatment (day 2) and at days 4, 6, and 8 of culture. Immunofluorescence studies were done on explants cultured on coverslips (Mazzalupo et al., 2002). Primary antibodies used were: rabbit polyclonal antisera directed against K6 or K17 (McGowan and Coulombe, 1998b), K16 (Bernot et al., 2002), and K5 (Covance); and mouse monoclonals directed against K14 (LLOO1; Purkis et al., 1990), BrdU (Sigma-Aldrich), and p120ctn (Transduction Labs). Actin filaments were labeled on PFA-fixed samples using rhodamine-conjugated phalloidin (Sigma-Aldrich). Secondary antibodies used were HRP- (Sigma-Aldrich) and fluorophore-conjugated (Kirkegaard & Perry Laboratories).
Skin grafting studies
Immunocompromised CD-1 nude mice (Charles River) were used as hosts. In brief, hosts were anesthetized with 25 µl/g of body weight of avertin and prepared by scoring the area of backskin where the graft was to be placed. The top layer of the skin was gently removed, leaving the vascular bed intact. 3-d-old pups derived from K6/K6ß hemizygous matings in the C57Bl/6 strain were killed and donor skin was placed on the host vascular bed, sutured (Sofsilk 40; United States Surgical), and covered with antibacterial ointment (Alpharma USPD, Inc.). Mice were housed individually thereafter. After normalization, skin grafts (two per genotype) were subjected to full thickness incisional wounding. 3 d later, the wound site and surrounding healthy tissue was biopsied at several locations and processed for morphological analyses. Alternatively, grafted tissue (two per genotype) was subjected to chemical treatment and mechanical trauma as follows. PMA (Sigma-Aldrich) was applied topically using a cotton pad on days 1, 3, and 5, and tissue was harvested on day 7. Mice were injected with 100 µg/g of body weight of BrdU (Sigma-Aldrich) 2 h before killing. Immediately before harvesting tissue for histology, gentle trauma was applied by rubbing with a pencil eraser 10 times across treated and untreated skin graft.
Morphological and protein analyses
For routine histopathology, tissues were fixed in Bouin's, paraffin-embedded, and 5-µm sections cut and stained with hematoxylin and eosin (H&E). Confocal microscopy was done using a dual spinning disk instrument (model Axiovert 200; Carl Zeiss MicroImaging, Inc.) equipped with a 63x oil objective and a digital camera (model OrcaER; Hamamatsu Co.). Image processing was done via the acquisition software (Ultraview 5.4; Orinda). For plastic embedding, samples were fixed in 2% glutaraldehyde, 1% PFA mixture overnight at 4°C, fixed after with 1% osmium tetroxide, embedded in epoxy resin and semi-thick sections were cut (0.5 µm thick) and stained with toluidine blue. To analyze proteins, cells derived from explants were harvested after 6 d in culture in ice-cold PBS supplemented with protease and phosphatase inhibitors (1 mM EGTA, 20 µM Na3VO4, 10 mM NaF, 1 µg/ml leupeptin, 2 µg/ml antipain, 10 µg/ml aprotinin, 10 µg/ml benzamidine, 1 µg/ml cymostatin, and 1 µg/ml pepstatin-A; Sigma-Aldrich). Skin tissue was removed and outgrown cells were scraped, collected by centrifugation, and lysed in buffer (1% deoxycholate, 1% Triton X-100, 0.1% SDS, 150 mM NaCl, 50 mM Tris, pH 7.5, 0.5 mM EDTA, and 1 mM EGTA) supplemented with inhibitors. Cell scrapings from at least 20 explants from three littermates with the same genotype were pooled. Soluble and insoluble protein fractions were obtained by centrifugation for 10 min at 4°C. Pellets were resuspended in a Tris-buffered 8 M urea solution with inhibitors. 2 µg of protein (Bradford assay) was subjected to SDS-PAGE, transferred onto nitrocellulose in the presence of 1 mM of sodium orthovanadate. Additional antibodies used include mouse monoclonals against K13, K15, and K16 (K8.12; Sigma-Aldrich) or phosphotyrosine (4G10; Upstate Biotechnology). Bound primary antibodies were detected using chemiluminescence (Pierce Chemical Co.).
RT-PCR analyses
Semi-quantitative RT-PCR was performed on total RNA prepared from 6-d-old skin explant cultures using TRIzol (Invitrogen) and reverse transcribed (Advantage RT-for PCR kit; CLONTECH Laboratories, Inc.). K16 (forward, 5'-AACAGCCTAGAAGAGACCAAAGGC-3'; and reverse, 5'-GGTAGGGGAGACAGATGGGGAATGCGC-3') and ß-tubulin (forward, 5'-CAACGTCAAGACGGCCGTGTG-3'; and reverse, 5'-GACAGAGGCAAACTGAGC- ACC-3') oligonucleotide primers were used. Reactions were sampled after 22, 27, 32, and 37 cycles (94°C for 1 min; 58°C for 1 min; and 68°C for 1 min) to monitor product accumulation.
![]() |
Acknowledgments |
---|
These studies were supported by grants F32-AR088553 (to P. Wong) and R01-AR42047 (to P.A. Coulombe) from the National Institutes of Health.
Submitted: 7 May 2003
Accepted: 10 September 2003
![]() |
References |
---|
![]() ![]() ![]() ![]() ![]() ![]() ![]() |
---|
Anastasiadis, P.Z., and A.B. Reynolds. 2001. Regulation of Rho GTPases by p120ctn. Curr. Opin. Cell Biol. 13:604610.[CrossRef][Medline]
Andra, K., B. Nikolic, M. Stocher, D. Drenckhahn, and G. Wiche. 1998. Not just scaffolding: plectin regulates actin dynamics in cultured cells. Genes Dev. 12:34423451.
Baribault, H., J. Price, K. Miyai, and R.G. Oshima. 1993. Mid-gestational lethality in mice lacking keratin 8. Genes Dev. 7:11911202.[Abstract]
Baribault, H., J. Penner, R.V. Iozzo, and M. Wilson-Heiner. 1994. Colorectal hyperplasia and inflammation in keratin 8-deficient FVB/N mice. Genes Dev. 8:29642973.[Abstract]
Bernot, K., P.A. Coulombe, and K.M. McGowan. 2002. Keratin 16 expression defines a subset of epithelial cells during skin morphogenesis and the hair cycle. J. Invest. Dermatol. 119:11371149.
Bousquet, O., L. Ma, S. Yamada, C. Gu, T. Idei, K. Takahashi, D. Wirtz, and P.A. Coulombe. 2001. The nonhelical tail domain of keratin 14 promotes filament bundling and enhances the mechanical properties of keratin intermediate filaments in vitro. J. Cell Biol. 155:747754.
Caulin, C., C.F. Ware, T.M. Magin, and R.G. Oshima. 2000. Keratin-dependent, epithelial resistance to tumor necrosis factor- induced apoptosis. J. Cell Biol. 149:1722.
Coulombe, P.A., and M.B. Omary. 2002. "Hard" and "soft" principles defining the structure, function and regulation of keratin intermediate filaments. Curr. Opin. Cell Biol. 14:110122.[CrossRef][Medline]
Coulombe, P.A., O. Bousquet, L. Ma, S. Yamada, and D. Wirtz. 2000. The "ins" and "outs" of intermediate filament organization. Trends Cell Biol. 10:420428.[CrossRef][Medline]
Cozzolino, M., V. Stagni, L. Spinardi, N. Campioni, C. Fiorentini, E. Salvati, S. Alema, and A.M. Salvatore. 2003. p120 catenin is required for growth factor-dependent cell motility and scattering in epithelial cells. Mol. Biol. Cell. 14:19641977.
Eckes, B., E. Colucci-Guyon, H. Smola, S. Nodder, C. Babinet, T. Krieg, and P. Martin. 2000. Impaired wound healing in embryonic and adult mice lacking vimentin. J. Cell Sci. 113:24552462.
Eckes, B., D. Dogic, E. Colucci-Guyon, N. Wang, A. Maniotis, D. Ingber, A. Merckling, F. Langa, M. Aumailley, A. Delouvee, et al. 1998. Impaired mechanical stability, migration and contractile capacity in vimentin-deficient fibroblasts. J. Cell Sci. 111:18971907.
Estrada, C.M., C.D. Park, M. Castilla, and R.A. Tassava. 1993. Monoclonal antibody WE6 identifies an antigen that is up-regulated in the wound epithelium of newts and frogs. Limb Development and Regeneration. Wiley-Liss Inc., New York. 272282.
Fabrizi, C., B.M. Kelly, C.S. Gillespie, W.W. Schlaepfer, S.S. Scherer, and P.J. Brophy. 1997. Transient expression of the neurofilament proteins NF-L and NF-M by Schwann cells is regulated by axonal contact. J. Neurosci. Res. 50:291299.[CrossRef][Medline]
Fuchs, E., and D.W. Cleveland. 1998. A structural scaffolding of intermediate filaments in health and disease. Science. 279:514519.
Fuchs, E., and K. Weber. 1994. Intermediate filaments: structure, dynamics, function, and disease. Annu. Rev. Biochem. 63:345382.[CrossRef][Medline]
Galou, M., E. Colucci-Guyon, D. Ensergueix, J.L. Ridet, M. Gimenez y Ribotta, A. Privat, C. Babinet, and P. Dupouey. 1996. Disrupted glial fibrillary acidic protein network in astrocytes from vimentin knockout mice. J. Cell Biol. 133:853863.[Abstract]
Gilbert, S., A. Loranger, N. Daigle, and N. Marceau. 2001. Simple epithelium keratins 8 and 18 provide resistance to Fas-mediated apoptosis. The protection occurs through a receptor-targeting modulation. J. Cell Biol. 154:763774.
Goldstein, M.E., S.R. Weiss, R.A. Lazzarini, P.S. Shneidman, J.F. Lees, and W.W. Schlaepfer. 1988. mRNA levels of all three neurofilament proteins decline following nerve transection. Brain Res. 427:287291.[Medline]
Hesse, M., T.M. Magin, and K. Weber. 2001. Genes for intermediate filament proteins and the draft sequence of the human genome: novel keratin genes and a surprisingly high number of pseudogenes related to keratin genes 8 and 18. J. Cell Sci. 114:25692575.[Medline]
Hofmann, I., and W.W. Franke. 1997. Heterotypic interactions and filament assembly of type I and type II cytokeratins in vitro: viscometry and determinations of relative affinities. Eur. J. Cell Biol. 72:122132.[Medline]
Irvine, A.D., and W.H. McLean. 1999. Human keratin diseases: the increasing spectrum of disease and subtlety of the phenotype-genotype correlation. Br. J. Dermatol. 140:815828.[CrossRef][Medline]
Kaya, S.S., A. Mahmood, Y. Li, E. Yavuz, M. Goksel, and M. Chopp. 1999. Apoptosis and expression of p53 response proteins and cyclin D1 after cortical impact in rat brain. Brain Res. 818:2333.[CrossRef][Medline]
Ku, N.O., S.A. Michie, R.M. Soetikno, E.Z. Resurreccion, R.L. Broome, and M.B. Omary. 1998. Mutation of a major keratin phosphorylation site predisposes to hepatotoxic injury in transgenic mice. J. Cell Biol. 143:20232032.
Lepekhin, E.A., C. Eliasson, C.H. Berthold, V. Berezin, E. Bock, and M. Pekny. 2001. Intermediate filaments regulate astrocyte motility. J. Neurochem. 79:617625.[CrossRef][Medline]
Li, Z., M. Mericskay, O. Agbulut, G. Butler-Browne, L. Carlsson, L.E. Thornell, C. Babinet, and D. Paulin. 1997. Desmin is essential for the tensile strength and integrity of myofibrils but not for myogenic commitment, differentiation, and fusion of skeletal muscle. J. Cell Biol. 139:129144.
Lloyd, C., Q.C. Yu, J. Cheng, K. Turksen, L. Degenstein, E. Hutton, and E. Fuchs. 1995. The basal keratin network of stratified squamous epithelia: defining K15 function in the absence of K14. J. Cell Biol. 129:13291344.[Abstract]
Loranger, A., S. Duclos, A. Grenier, J. Price, M. Wilson-Heiner, H. Baribault, and N. Marceau. 1997. Simple epithelium keratins are required for maintenance of hepatocyte integrity. Am. J. Pathol. 151:16731683.[Abstract]
Mansbridge, J.N., and A.M. Knapp. 1987. Changes in keratinocyte maturation during wound healing. J. Invest. Dermatol. 89:253263.[Medline]
Martin, P. 1997. Wound healing - aiming for the perfect skin regeneration. Science. 276:7581.
Mazzalupo, S., M.J. Wawersik, and P.A. Coulombe. 2002. An ex vivo assay to assess the potential of skin keratinocytes for wound epithelialization. J. Invest. Dermatol. 118:866870.
McGraw, T.S., J.P. Mickle, G. Shaw, and W.J. Streit. 2002. Axonally transported peripheral signals regulate alpha-internexin expression in regenerating motoneurons. J. Neurosci. 22:49554963.
McGowan, K.M., and P.A. Coulombe. 1998a. Onset of keratin 17 expression coincides with the definition of major epithelial lineages during skin development. J. Cell Biol. 143:469486.
McGowan, K.M., and P.A. Coulombe. 1998b. The wound repair associated keratins 6, 16, and 17: insights into the role of intermediate filaments in specifying cytoarchitecture. Subcellular Biochemistry: Intermediate Filaments. J.R. Harris and H. Herrmann, editors. Plenum Publishing Corp., London. 141165.
McGowan, K.M., X. Tong, E. Colucci-Guyon, F. Langa, C. Babinet, and P.A. Coulombe. 2002. Keratin 17 null mice exhibit age- and strain-dependent alopecia. Genes Dev. 16:14121422.
Moll, R., W.W. Franke, D.L. Schiller, B. Geiger, and R. Krepler. 1982. The catalog of human cytokeratins: patterns of expression in normal epithelia, tumors and cultured cells. Cell. 31:1124.[Medline]
Muma, N.A., P.N. Hoffman, H.H. Slunt, M.D. Applegate, I. Lieberburg, and D.L. Price. 1990. Alterations in levels of mRNAs coding for neurofilament protein subunits during regeneration. Exp. Neurol. 107:230235.[Medline]
Omary, M.B., N.-O. Ku, J. Liao, and D. Price. 1998. Keratin modifications and solubility properties in epithelial cells and in vitro. Subcellular Biochemistry: Intermediate Filaments. Vol. 31. H. Herrman and J.R. Harris, editors. Plenum Press, New York. 105140.
Oshima, R.G. 2002. Apoptosis and keratin intermediate filaments. Cell Death Differ. 9:486492.[CrossRef][Medline]
Paladini, R.D., K. Takahashi, N.S. Bravo, and P.A. Coulombe. 1996. Onset of re-epithelialization after skin injury correlates with a reorganization of keratin filaments in wound edge keratinocytes: defining a potential role for keratin 16. J. Cell Biol. 132:381397.[Abstract]
Pang, S.C., W.H. Daniels, and R.C. Buck. 1978. Epidermal migration during the healing of suction blisters in rat skin: a scanning and transmission electron microscopic study. Am. J. Anat. 153:177191.[Medline]
Pekny, M., C.B. Johansson, C. Eliasson, J. Stakeberg, A. Wallen, T. Perlmann, U. Lendahl, C. Betsholtz, C.H. Berthold, and J. Frisen. 1999. Abnormal reaction to central nervous system injury in mice lacking glial fibrillary acidic protein and vimentin. J. Cell Biol. 145:503514.
Purkis, P.E., J.B. Steel, I.C. Mackenzie, W.B. Nathrath, I.M. Leigh, E.B. Lane. 1990. Antibody markers of basal cells in complex epithelia. J. Cell Sci. 97:3950.[Abstract]
Quinlan, R., C. Hutchinson, and E.B. Lane. 1994. Intermediate filament proteins. Protein Profile. 1:779911.[Medline]
Ramirez, A., M. Vidal, A. Bravo, F. Larcher, and J.L. Jorcano. 1995. A 5'-upstream region of a bovine keratin 6 gene confers tissue-specific expression and hyperproliferation-related induction in transgenic mice. Proc. Natl. Acad. Sci. USA. 92:47834787.[Abstract]
Reichelt, J., and T.M. Magin. 2002. Hyperproliferation, induction of c-Myc and 14-3-3sigma, but no cell fragility in keratin-10-null mice. J. Cell Sci. 115:26392650.
Schermer, A., J.V. Jester, C. Hardy, D. Milano, and T.T. Sun. 1989. Transient synthesis of K6 and K16 keratins in regenerating rabbit corneal epithelium: keratin markers for an alternative pathway of keratinocyte differentiation. Differentiation. 42:103110.[Medline]
Shibuya, S., O. Miyamoto, R.N. Auer, T. Itano, S. Mori, and H. Norimatsu. 2002. Embryonic intermediate filament, nestin, expression following traumatic spinal cord injury in adult rats. Neuroscience. 114:905916.[Medline]
Smythe, G.M., M.J. Davies, D. Paulin, and M.D. Grounds. 2001. Absence of desmin slightly prolongs myoblast proliferation and delays fusion in vivo in regenerating grafts of skeletal muscle. Cell Tissue Res. 304:287294.[CrossRef][Medline]
Stenn, K.S., and R. Paus. 2001. Controls of hair follicle cycling. Physiol. Rev. 81:449494.
Swensson, O., L. Langbein, J.R. McMillan, H.P. Stevens, I.M. Leigh, W.H. McLean, E.B. Lane, and R.A. Eady. 1998. Specialized keratin expression pattern in human ridged skin as an adaptation to high physical stress. Br. J. Dermatol. 139:767775.[CrossRef][Medline]
Takahashi, K., and P.A. Coulombe. 1997. Defining a region of the human keratin 6a gene that confers inducible expression in stratified epithelia of transgenic mice. J. Biol. Chem. 272:1197911985.
Takahashi, K., R. Paladini, and P.A. Coulombe. 1995. Cloning and characterization of multiple human genes and cDNAs encoding highly related type II keratin 6 isoforms. J. Biol. Chem. 270:1858118592.
Takahashi, K., B. Yan, K. Yamanishi, S. Imamura, and P.A. Coulombe. 1998. The two functional keratin 6 genes of mouse are differentially regulated and evolved independently from their human orthologs. Genomics. 53:170183.[CrossRef][Medline]
Tanaka, H., A. Katoh, K. Oguro, K. Shimazaki, H. Gomi, S. Itohara, T. Masuzawa, and N. Kawai. 2002. Disturbance of hippocampal long-term potentiation after transient ischemia in GFAP deficient mice. J. Neurosci. Res. 67:1120.[CrossRef][Medline]
Tetzlaff, W., M.A. Bisby, and G.W. Kreutzberg. 1988. Changes in cytoskeletal proteins in the rat facial nucleus following axotomy. J. Neurosci. 8:31813189.[Abstract]
Toivola, D.M., H. Baribault, T. Magin, S.A. Michie, and M.B. Omary. 2000. Simple epithelial keratins are dispensable for cytoprotection in two pancreatitis models. Am. J. Physiol. Gastrointest. Liver Physiol. 279:G1343G1354.
Tyner, A.L., and E. Fuchs. 1986. Evidence for posttranscriptional regulation of the keratins expressed during hyperproliferation and malignant transformation in human epidermis. J. Cell Biol. 103:19451955.[Abstract]
Vaittinen, S., R. Lukka, C. Sahlgren, T. Hurme, J. Rantanen, U. Lendahl, J.E. Eriksson, and H. Kalimo. 2001. The expression of intermediate filament protein nestin as related to vimentin and desmin in regenerating skeletal muscle. J. Neuropathol. Exp. Neurol. 60:588597.[Medline]
Wang, Z., P. Wong, L. Langbein, J. Schweizer, and P.A. Coulombe. 2003. The type II epithelial keratin 6 hf (K6hf) is expressed in the companion layer, matrix, and medulla of anagen-stage hair follicles. J. Invest. Dermatol. In press.
Wawersik, M., R.D. Paladini, E. Noensie, and P.A. Coulombe. 1997. A proline residue in the alpha-helical rod domain of type I keratin 16 destabilizes keratin heterotetramers and influences incorporation into filaments. J. Biol. Chem. 272:3255732565.
Wawersik, M.J., S. Mazzalupo, D. Nguyen, and P.A. Coulomb. 2001. Increased levels of keratin 16 alter the epithelialization potential of mouse skin keratinocytes in vivo and ex vivo. Mol. Biol. Cell. 12:34393450.
Wojcik, S.M., D.S. Bundman, and D.R. Roop. 2000. Delayed wound healing in keratin 6a knockout mice. Mol. Cell. Biol. 20:52485255.
Wojcik, S.M., M.A. Longley, and D.R. Roop. 2001. Discovery of a novel murine keratin 6 (K6) isoform explains the absence of hair and nail defects in mice deficient for K6a and K6b. J. Cell Biol. 154:619630.
Wong, P., E. Colucci-Guyon, K. Takahashi, C. Gu, C. Babinet, and P.A. Coulombe. 2000. Introducing a null mutation in the mouse K6 and K6ß genes reveals their essential structural role in the oral mucosa. J. Cell Biol. 150:921928.
Yamada, S., D. Wirtz, and S.C. Kuo. 2000. Mechanics of living cells measured by laser tracking microrheology. Biophys. J. 78:17361747.
Yamada, S., D. Wirtz, and P.A. Coulombe. 2002. Pairwise assembly determines the intrinsic potential for self-organization and mechanical properties of keratin filaments. Mol. Biol. Cell. 13:382391.
Zhu, Q., S. Couillard-Despres, and J.P. Julien. 1997. Delayed maturation of regenerating myelinated axons in mice lacking neurofilaments. Exp. Neurol. 148:299316.[CrossRef][Medline]