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J. Biol. Chem., Vol. 282, Issue 11, 8175-8187, March 16, 2007
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1
2
From the
Departments of Molecular Cell Biology and Biomolecular Structural Chemistry, Max F. Perutz Laboratories, University of Vienna, A-1030 Vienna, Austria
Received for publication, September 5, 2006 , and in revised form, December 20, 2006.
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ABSTRACT |
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 TOP ABSTRACT INTRODUCTIONEXPERIMENTAL PROCEDURESRESULTSDISCUSSIONREFERENCES |
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INTRODUCTION |
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TOPABSTRACTINTRODUCTIONEXPERIMENTAL PROCEDURESRESULTSDISCUSSIONREFERENCES |
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Plectin belongs to the plakin family of cytolinker proteins, which also includes desmoplakin, BPAG1, ACF7/microtubule actin cross-linking factor, envoplakin, periplakin, and epiplakin (8). Like most of the other proteins in this family, plectin has a multidomain structure consisting of a globular N-terminal domain; a central rod domain, and a globular C-terminal domain, which itself is multimodular. Electron microscopy of purified plectin molecules confirmed their three-domain structure (9). Functionally, the N-terminal domain harbors an actin-binding domain; the central rod domain mediates dimerization; and the C-terminal domain contains the IF-binding site. The building block of the plectin C terminus is a repeat (R) domain known as the plakin or plectin repeat domain (8, 10). R domains consist of a conserved core domain or module built from 4.5 tandemly repeated copies of a 38-amino acid motif. The modules are separated from each other by linker sequences of variable lengths. The concept of the module as the basic structure of the C-terminal domain of plakin proteins has been validated when the crystal structure of two desmoplakin R domains was solved (11). The C terminus of plectin (
1900 amino acids) features six highly homologous R domains (R1–6) of 300 amino acids each. Accommodation of the six R domains of plectin within its globular C-terminal domain, which has an estimated diameter of 9 nm (9), requires tight packing. Janda et al. (10) proposed a circular arrangement of antiparallel-oriented plectin R1–5 domains with the R6 domain in their center; such a structure could be stabilized by hydrophobic interactions between residues on the surface of each R domain and/or through cysteines via disulfide bond formation due to physiological bursts of oxidative stress.
The C-terminal part of plectin contains from 13 (mouse) to 17 (human) cysteines (12, 13). Four of these generally highly conserved residues are clustered in the plectin R5 domain, which harbors also the major IF-binding site of the protein (14). This provocative localization might serve a functional purpose, as these cysteines could form intra- and inter-repeat disulfide bridges, providing more structural rigidity to the protein itself. In addition, they may mediate interactions of the protein with various binding partners with docking sites in this region, such as various IF subunit proteins (reviewed in Ref. 1) and proteins involved in cellular signaling, including RACK1 (3), a regulator of protein kinase C, and AMP-activated protein kinase (4), a key regulatory enzyme of energy homeostasis. Intracellular disulfide bridge formation is likely to be of particular importance in situations in which cells have to respond to mechanical, oxidative, or other types of stress. Because full-length plectin or even entire subdomains, such as the C-terminal globular domain with its six R domains (>500 and >200 kDa, respectively), are too large to be recombinantly expressed for biochemical analyses, this study was restricted to the plectin R5 domain.
The following questions regarding R5 cysteines were addressed. Does their oxidation change the biochemical and conformational properties of the protein? Are they accessible to disulfide cross-linking within R5 and between R5 and R4? Is the binding affinity of R5 for vimentin affected by their oxidative state? Are they targets of nitrosylation, and if so, does nitrosylation occur in vivo, and does it have consequences for plectin-regulated cellular functions?
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EXPERIMENTAL PROCEDURES |
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TOPABSTRACTINTRODUCTIONEXPERIMENTAL PROCEDURESRESULTSDISCUSSIONREFERENCES |
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Expression and Purification of Proteins—Recombinant proteins were expressed in Escherichia coli strain BL21(DE3) after induction with 1 mM isopropyl 
-D-thiogalactopyranoside. Unless otherwise indicated, His-tagged recombinant proteins were prepared under reducing conditions and affinity-purified on nickel columns (His·Bind, Novagen) following the protocol of the manufacturer. The bacterial cell pellets were resuspended in a lysis buffer containing 10 mM piperazine (pH 11.0), 1% Triton X-100, and 10 mM -mercaptoethanol, and the binding, washing, and elution column buffers (20 mM Tris-HCl (pH 9.0) containing 5, 20, and 250 mM imidazole, respectively) were supplemented with 5 mM -mercaptoethanol. Proteins were kept in elution buffer at 4 °C and dialyzed against the required buffer prior to being assayed. All solutions were degassed and supplemented with -mercaptoethanol until the dialysis step, except in europium binding assays and CD spectroscopy, in which no -mercaptoethanol was added. Recombinant mouse full-length vimentin was purified by sequential ion-exchange column chromatography on DEAE-Sepharose and CM-Sepharose (GE Healthcare) as described previously (15). Also in this case, all buffers were degassed. Protein concentrations were estimated by the Bradford or BCA (Pierce) method.
SDS-PAGE and Immunoblotting—Protein samples (10 µg/20 µl) were analyzed by SDS-10% PAGE under reducing and nonreducing conditions. The reduction of samples before application to the gel was achieved by the addition of 0.2 M dithiothreitol (DTT) to 2x sample buffer. After electrophoresis, the gels were stained with 0.25% Coomassie Brilliant Blue, 45% methanol, and 10% acetic acid and scanned in an HP Scan-Jet 8250 scanner. Densitometric analysis was performed with Gel Doc 2000 gel documentation system (Bio-Rad) and Quantity One image analysis software (Bio-Rad). Transfer to 0.2-µm nitrocellulose membranes (Protran®, Schleicher & Schüll) and immunoblotting were done following standard procedures.
Spectroscopy and Urea-induced Unfolding—The details of far-UV CD and fluorescence emission spectroscopy, including sample preparation, are provided in supplemental "Experimental Procedures."
Oxidative Cross-linking—Purified recombinant proteins at a concentration of 0.5 mg/ml were dialyzed overnight at 4 °C against 10 mM Tris-HCl (pH 7.9) while being concurrently exposed to oxidation by air (no precautions were taken to exclude air from the solutions). The reaction was quenched by the addition of iodoacetamide (final concentration of 50 mM) to block free sulfhydryl groups. Samples were then resolved by SDS-10% PAGE under nonreducing or reducing conditions. Alternatively, aliquots of two different recombinant proteins were mixed 1:1 at a concentration of 0.5 mg/ml each; dialyzed against 10 mM Tris-HCl (pH 7.9), 6 M urea, and 1 mM DTT for 1.5 h at room temperature; and subsequently oxidized by air while being dialyzed at 4 °C into 10 mM Tris-HCl (pH 7.9) without urea. The reactions were then quenched and analyzed as described above.
Blot Overlay Binding Assay—Purified proteins (0.5 µg/lane) were subjected to SDS-10% PAGE and blotted onto nitrocellulose membranes. The membranes were blocked for 1 h with 3% bovine serum albumin (BSA) in phosphate-buffered saline; incubated for 1 h with purified samples of vimentin (10 µg/ml) in 137 mM NaCl, 2.7 mM KCl, 4.3 mM Na2HPO4·7H2O, 1.4 mM KH2PO4, 1 mM EGTA, 2 mM MgCl2, 1 mM DTT, and 0.1% (v/v) Tween 20 (pH 7.5); and washed several times with phosphate-buffered saline containing 0.05% (v/v) Tween 20. Bound vimentin was detected using affinity-purified goat anti-mouse antibodies to vimentin (diluted 1:5000; kindly provided by Peter Traub, University of Bonn, Bonn, Germany) (16), followed by incubation with horseradish peroxidase-conjugated anti-goat IgG (diluted 1:5000; Jackson ImmunoResearch Laboratories, Inc.). Immune complexes were detected by chemiluminescence (West Pico ECL, Pierce).
Co-sedimentation Assay—Co-sedimentation assays were carried out essentially as described by Choi et al. (11) with minor modifications regarding protein preparation (see above) and buffer composition. To prepare vimentin filaments, recombinant vimentin was dialyzed stepwise at room temperature against 5 mM Tris acetate (pH 8.3), 1 mM EDTA, and 10 mM -mercaptoethanol with decreasing concentrations of urea (6 to 0 M). The dialyzed vimentin solution was centrifuged in a Beckman benchtop ultracentrifuge (OptimaTM TLX) at 100,000 x g for 30 min at room temperature, and soluble vimentin (8.9 µM) was polymerized in 10 µl of 20 mM HEPES (pH 7.5), 150 mM NaCl, 1 mM EDTA, and 0.1% (v/v) Tween 20 (buffer A) for 30 min at 37 °C.
Recombinant plectin R5 domains (wild-type and cysteine-free mutant) were heated in elution buffer supplemented with 0.1% (v/v) Tween 20 for 10 min at 37 °C and dialyzed sequentially against 20 mM Tris-HCl (pH 8.1) and 0.1% (v/v) Tween 20 (buffer B), buffer B supplemented with 1 mM EDTA, and buffer A. All solutions were degassed. Recombinant R4 was dialyzed against the same set of buffers without an initial heating step. R4 and wild-type and cysteine-free R5 were then added to polymerized vimentin (8.9 µM in 10 µl of buffer A) at different concentrations (1–10:1 molar ratios of vimentin to plectin). After the addition of buffer A to a total volume of 50–100 µl, the mixtures were incubated for 1 h at 37 °C. Vimentin filaments and associated protein were pelleted by centrifugation at 100,000 x g for 30 min at room temperature using the Beckman OptimaTM TLX ultracentrifuge. Supernatants were withdrawn immediately after centrifugation, and an equal volume of 2x SDS-PAGE sample buffer was added. Pellets were dissolved in 50 µl of 2x SDS-PAGE sample buffer and 50 µl of buffer A. The supernatant and pellet fractions were analyzed by SDS-10% PAGE. For controls without vimentin, 8.9 µM R4, wild-type and cysteine-free R5, and BSA were treated in the same manner.
Europium Binding Assay—Recombinant vimentin dialyzed stepwise against 50 mM NaHCO3 (pH 8.5) and labeled with Eu3+ was overlaid in serial dilutions onto 96-microtiter plates coated with wild-type and cysteine-free R5 (both at 100 nM). Bound protein was determined by releasing the vimentin-bound Eu3+ with enhancement solution and measuring fluorescence with a Wallac time-resolved microtiter plate fluorometer (excitation wavelength of 340 nm and emission wavelength of 615 nm). Further details of this assay have been described previously (14, 15). The Scatchard method was used for analysis of binding data, and fluorescence values were converted to concentrations by comparison with an Eu3+ standard.
Cell Culture and Immunofluorescence Microscopy—The details of the procedures used for the isolation and immortalization of endothelial cells from mouse kidneys and of cell processing for immunofluorescence microscopy are given under supplemental "Experimental Procedures." Endothelial cells to be analyzed by immunofluorescence microscopy after NO donor-mediated nitrosylation were grown on gelatin-coated glass coverslips; treated with 100 µM S-nitroso-N-acetylpenicillamine (SNAP) in the dark for 2, 4, or 6 h; washed with phosphate-buffered saline; and then fixed.
S-Nitrosylation Assays—S-Nitrosylated proteins were detected using the biotin-switch assay as described by Jaffrey and Snyder (17). For in vitro assays, purified wild-type R5 and R5-C4S (80 µg each) in 250 mM HEPES (pH 7.7), 1 mM EDTA, and 0.1 mM neocuproine (HEN buffer) were incubated in the dark with 100 µM SNAP for 2 h at room temperature, and the NO donors were then removed by passing the samples twice through a desalting column (Micro Bio-Spin P-6, Bio-Rad). Proteins in the flow-through were blocked with 20 mM methyl methanethiosulfonate (Sigma) for 20 min at 50 °C, precipitated with acetone for 20 min at -20 °C, and collected by centrifugation at 10,000 x g for 10 min at 4 °C. Pellets were resuspended in 500 µl of HEN buffer containing 1% SDS (HENS buffer) and incubated with 1 mM ascorbic acid to release NO from thiol groups, which were subsequently biotinylated with 1 mM N-(6-(biotinamido)hexyl)-3'-(2'-pyridyldithio)propionamide (Pierce). Proteins were again acetone-precipitated and resuspended in 300 µl of HENS buffer, and biotinylated proteins were recovered by streptavidin affinity chromatography. Eluted proteins were separated by SDS-10% PAGE; transferred to nitrocellulose membranes; analyzed by immunoblotting using antibodies to plectin (diluted 1:2500; No. 9) (18), endothelial nitric-oxide synthase (NOS; diluted 1:1000; BD Transduction Laboratories), and His tag (diluted 1:1000; Qiagen Inc.); and visualized by chemiluminescence. For the identification of S-nitrosylated proteins in cultured cells, confluent mouse renal endothelial cells (2.5 x 107) were incubated with 100 µM SNAP for 2 h in the dark or with 100 nM phorbol 12-myristate 13-acetate (PMA) for 20 h, washed thoroughly with phosphate-buffered saline, scraped off, lysed in 350 µl of HEN buffer supplemented with 2.5% SDS, blocked with methyl methanethio-sulfonate, and subjected to the biotin-switch assay as described above.
Homology Modeling—A model was generated using an automated homology modeling server (ExPASy proteomics server using SWISS-MODEL ProModII) running at the Swiss Institute of Bioinformatics (Geneva, Switzerland) and Geno3D (Lyon, France). The structural template used for modeling was the crystal structure of human desmoplakin repeat domain B (Protein Data Bank code 1LM7) (11), which shares 73% sequence identity with the plectin R5 module. Modeling by satisfaction of spatial restraints was performed by the method of Combet et al. (19). In brief, the alignment of the target (plectin R5 module) and the template (desmoplakin domain B) was obtained using the Needleman-Wunsch global alignment algorithm on the EMBL-EBI server (www.ebi.ac.uk/emboss/align/). Restraints on various distances, angles, and dihedral angles in the sequence were derived from the alignment of the target with the template structure. Finally, the three-dimensional model was refined by applying distance geometry, simulated annealing, and energy minimization procedures. Visualization of the three-dimensional structure was performed using the molecular graphics package Yasara (www.yasara.com).
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RESULTS |
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TOPABSTRACTINTRODUCTIONEXPERIMENTAL PROCEDURESRESULTSDISCUSSIONREFERENCES |
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-sheets (forming a -hairpin) and two antiparallel 
-helices (Fig. 1, A and B). The fold is remarkably similar to that of the ankyrin repeat, as it was predicted by threading analysis (10). Conserved hydrophobic residues in the -hairpin contribute to the packing within each PLEC repeat and promote the adoption of a globular, cylinder-like structure that is 45 Å long with a 25-Å diameter. Cys1 is located in the -hairpin of the first PLEC repeat, whereas Cys2 and Cys3 are in the terminal part of the fifth PLEC repeat, which interacts with the first PLEC repeat (Fig. 1B). Cys2 and Cys3 are on the surface of the structure, whereas Cys1 is partially buried in a groove. The crystallized desmoplakin fragment does not include the linker region containing the corresponding Cys4. There is evidence, however, that this cysteine, residing in the loop connecting R domains B and C, is exposed, as partial chymotryptic digestion of the bacterially expressed C-terminal domain of desmoplakin results in cleavage of the polypeptide within this R domain linker region (11). Secondary structure predictions for this region suggest that Cys4 is located in a short unstructured sequence connecting an -helical segment and a -sheet (Fig. 1A). This linker region has also one of the highest sequence conservations among different plakin family members (22). An alignment of the corresponding linker regions of plectin and desmoplakin is shown in Fig. 1C. Expression of Functional Wild-type and Mutant R5 Domains—To assess the potential of individual R5 cysteines to form disulfide bridges, we generated recombinant versions of R5 in which all four cysteines were replaced with serines either individually or in different combinations. Cys1 and Cys4 were mutated alone and in various combinations with Cys2 and Cys3, including a mutant without any cysteine (Cys-free R5). In addition, we generated a mutant with native Cys1 and Cys4, but without Cys2 and Cys3 (R5-C2S,C3S), and its counterpiece with Cys2 and Cys3, but without Cys1 and Cys4 (R5-C1S,C4S). Schematics of wild-type R5 and the eight mutant versions generated and their assigned names are shown in Fig. 1D. The wild-type and mutant proteins were expressed to a similar extent and were equally well soluble, except for the cysteine-free mutant, which showed a slight tendency to aggregate. Proteins were purified to homogeneity, and their purity and functional competence were verified by SDS-PAGE and co-sedimentation with in vitro assembled vimentin IFs (data not shown).
The overexpression of R5 leads to the collapse of IF networks in different types of cells (14, 15). To further demonstrate that the mutations had not caused significant changes in R5 function, we transfected mammalian expression plasmids encoding Myc-tagged versions of wild-type and Cys-free R5 domains into PtK2 and mouse fibroblast cells. As monitored by immunofluorescence microscopy, overexpression of either protein led to IF collapse in both cell types after 2 days (supplemental Figs. S1 and S2), without any detectable phenotypic differences. Similarly, when immortalized keratinocytes (18) were subjected to this type of analysis, in agreement with previous results (14), a decoration but not a collapse of keratin networks was observed with both versions of R5 (data not shown). There was also no detectable difference between the two R5 variants regarding effects on actin filament or focal adhesion contact organization in fibroblasts (data not shown). Thus, the specific replacement of cysteines with serines did not appear to alter the basic association patterns of plectin with its cellular partners, strongly suggesting that the mutants were functional.
Biochemical and Conformational Properties of Wild-type R5 Compared with Its Cysteine-free Variant—Optical spectroscopic methods were used to reveal structural differences and changes in thermodynamic stability associated with the elimination of all four cysteines in R5. In particular, far-UV CD spectroscopy was used to obtain global information about the secondary structure content, whereas the intrinsic fluorescence of the single tryptophan located in close vicinity of Cys4 (Fig. 1C) was exploited as a local probe of the tertiary structure.
The far-UV CD spectrum of wild-type R5 (Fig. 2A, upper panel) measured at pH 7.5 showed two minima at 222 and 208 nm of almost equal negative ellipticity, typical for a high content of -helical structure. In contrast, the second minimum in the spectrum of cysteine-free R5 (Fig. 2A, upper panel) not only was shifted toward 205 nm, but showed an increased negative ellipticity, indicating a higher content of random coil structure. These findings were verified by estimation of the secondary structure content using three different algorithms (SELCON, CONTIN, and CDSSTR), which yielded consistently 22% -helical, 27% -sheet, and 50% random coil content for wild-type R5, whereas for cysteine-free R5, the -helical content was reduced to 17% in conjunction with an increased random coil content of 55%. Most important, the far-UV CD spectra of cysteine-free R5 recorded at pH 9 and 11 (Fig. 2A, lower panel) became essentially identical to those of wild-type R5, which were pH-insensitive (middle panel). In sum, the CD spectra of the wild-type and cysteine-free mutant R5 proteins were essentially identical, indicating that the substitution had a negligible effect on the overall secondary structure and did not cause large structural perturbation. Additionally, it can be concluded that the contribution of the four R5 cysteine residues to the far-UV CD signal is negligible.
The fluorescence spectrum of the single tryptophan at pH 7.5 exhibited an emission maximum at 335 nm for wild-type R5, whereas for the cysteine-free mutant, the maximum was shifted to 341 nm (data not shown), indicating a less buried conformation of the environment-sensitive tryptophan residue (23). Incubation of both proteins in 8 M urea resulted in red shifts to 348.5 nm for the wild-type protein and to 350.5 nm for the mutant protein, indicating, in both cases, an exposure of the tryptophan residue to the polar solvent. SDS-PAGE of samples of wild-type R5 in its native state (no urea) showed that the vast majority of wild-type R5 used for the measurement was in the oxidized monomeric form (data not shown), resulting from disulfide bond formation between Cys1 and Cys4 (see below). As a consequence, the tryptophan residue neighboring Cys4 became buried within the protein structure.
The thermodynamic stability was determined by urea-induced unfolding at pH 7.5 monitored by far-UV CD and tryptophan fluorescence (Fig. 2B). The unfolding of wild-type R5 showed sigmoidal transition curves for both far-UV CD and fluorescence with transition midpoints at 3.1 and 2.9 M urea, respectively. In contrast, the unfolding profiles for the cysteine-free mutant were clearly not coincident. The transition midpoint of the tryptophan fluorescence signal was at 1.5 M urea, whereas the far-UV CD unfolding profile reflected a very broad transition without a distinct base line for the native state. This behavior indicates the existence of intermediate states, where the tryptophan fluorescence reports only the unfolding of the C-terminal part of R5. Free energies of unfolding were not calculated, as these calculations require the proteins to follow a two-state transition. However, the observed transition midpoints allowed a semiquantitative comparison of the thermodynamic stabilities of wild-type R5 and the cysteine-free mutant.
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41 kDa) and oxidized (38 kDa) monomer and was able to form dimers, trimers, tetramers, and other oligomers (Fig. 3A, lane 1). The faster migration of the oxidized form compared with the reduced form of the monomer is typical for intramolecular disulfide bonding, resulting in a more tightly folded structure (24, 25). R5-C2S,C3S gave a cross-linking pattern similar to that of wild-type R5 (Fig. 3A, lane 8). Because the only cysteines available in the R5-C2S,C3S mutant are Cys1 and Cys4, we concluded that the oxidized forms of the monomer were presumably formed by disulfide bond formation between these two residues. Mutants R5-C1S and R5-C1S,C2S,C3S displayed two prominent bands corresponding to the reduced form of the monomer (41 kDa) and dimers with an abnormal mobility of 100 kDa (Fig. 3A, lanes 2 and 5). We believe that these dimers were formed by disulfide pairing of two Cys4 residues in two different R5 molecules. Cys4 resides in the linker region between modules 5 and 6 at a relatively large distance from the core region of R5. Disulfide bond formation between two Cys4 residues may therefore generate dimers with a larger hydrodynamic dimension, thus migrating slower (apparent molecular mass of 100 kDa instead of 82 kDa) upon SDS-PAGE due to their extended conformation (26, 27). In contrast, the cross-linked R5-C4S mutant formed a dimer migrating at the expected mobility of 82 kDa (Fig. 3A, lane 7). Cys-free R5 occurred only as the reduced form of the monomer (41 kDa) (Fig. 3A, lane 6). In addition to a protein band with an apparent molecular mass of 100 kDa, mutant R5-C1S,C3S yielded a band corresponding to a higher oligomer (Fig. 3A, lane 3). The rest of the mutants, R5-C1S,C2S, and R5-C1S,C4S, remained in the monomeric form after oxidation (41 kDa) (Fig. 3A, lanes 4 and 9).
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We speculated earlier that plectin R domains may bind to each other not only because of hydrophobic and electrostatic interactions, but also because of disulfide bond formation between the repeats (10). To test this hypothesis, we first examined whether the single cysteine residue present in R4 is able to form a disulfide bond between two R4 molecules. When R4 oxidatively cross-linked under nonreducing conditions was analyzed, two bands were found, one corresponding to the monomer and the other to the dimer, both with the expected mobilities (Fig. 3B, lane 1), whereas a single monomeric band was observed under reducing conditions (data not shown). Next, we tested whether R4 and R5 could be linked via a disulfide bridge. Heterodimer formation was examined by incubating R4 with wild-type R5 under reducing conditions for 1.5 h in the presence of 6 M urea, followed by dialysis to remove urea and by SDS-PAGE, both without reducing agents. Because of the size difference of R4 and R5 (30 and 41 kDa, respectively), a band of a size between that of the homodimers (61 and 82 kDa, respectively) would be indicative of heterodimer formation. Such a band was indeed observed (Fig. 3B, lane 4). This band was absent when R4 was incubated with the cysteine-free R5 mutant (Fig. 3B, lane 5). Thus, the single cysteine in R4 as well as cysteines in R5 formed disulfide bonds not only between their own molecular entities (R4/R4 and R5/R5 homodimers), but also between each other (R4/R5 heterodimers).
Increased Vimentin Binding Affinity of Plectin R5 in the Reduced Compared with the Oxidized Form—The major vimentin-binding site of plectin is located in the linker region connecting the R5 and R6 domains, where also Cys4 is found (Fig. 1C). It was therefore of special interest to examine whether this cysteine plays any role in vimentin-R5 interaction. In a first series of experiments, vimentin binding was assessed by co-sedimentation of wild-type or cysteine-free R5 with IFs assembled from recombinant vimentin in vitro. After incubation of mixtures containing vimentin and R5 proteins at different molar ratios (1:1, 5:1, and 10:1), filaments were sedimented, and supernatants and pellets were analyzed by SDS-PAGE, followed by densitometric quantification of protein bands. Samples containing R4 or BSA (at 1:1 molar ratios to vimentin) instead of R5 and additional controls containing just vimentin, BSA, R4, or R5 (without the corresponding binding partner) were processed and analyzed under similar conditions. At a vimentin/R5 ratio of 1:1, nearly 90% of R5 co-sedimented with vimentin, whereas no plectin remained unbound at a ratio of 5:1 or 10:1 (Fig. 4A), indicating optimal binding at molar ratios somewhere in between 1:1 and 5:1. No significant differences in the vimentin binding abilities of wild-type R5 and its cysteine-free variant were detectable, however, using this assay. Regarding the controls, BSA and plectin R4 did not co-sediment with vimentin (Fig. 4A, lanes 1 and 2), and when vimentin, BSA, both versions of R5, and R4 were incubated and processed alone, only polymerized vimentin was found in the pellets (Fig. 4B), except for a minor apparently self-aggregating fraction of Cys-free R5 (Fig. 4, compare B, lane 4, and A, lane 4P).
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41 kDa) of monomeric wild-type R5 and its disulfide-linked (oxidized) higher order oligomers (Fig. 4C, lanes 3 and 4), as well as to Cys-free R5 (lane 5). In contrast, vimentin showed no binding to the oxidized monomeric form (38 kDa) of wild-type R5 (Fig. 4C, lane 3). This suggested that the IF-binding site of plectin was not accessible for vimentin upon disulfide bond formation between Cys1 and Cys4. The binding of vimentin to the BSA control or R4 was not observed (Fig. 4C, lanes 1 and 2). The differential binding of vimentin to the reduced and oxidized forms of R5 prompted us to measure the binding affinities of R5 and its variants for vimentin. To this end, we titrated the binding of Eu3+-labeled vimentin to immobilized wild-type and cysteine-free mutant R5 proteins under reducing and nonreducing conditions. The dissociation constants obtained showed that wild-type R5 bound with 2-fold higher affinity to vimentin under reducing (Kd = 0.155 µM) compared with nonreducing conditions (Kd = 0.312 µM), whereas an even higher binding affinity (Kd = 0.096 µM) was observed for the mutant protein (Fig. 4D). Evidently, these differences in binding affinity reflect distinct structures of the reduced and nonreduced forms of R5 on the one hand and of wild-type R5 and its cysteine-free mutant on the other.
Nitrosylation of R5 Cys4 in Vitro—S-Nitrosylation is a reversible post-translational modification with a potential role in the regulation of protein function in response to oxidative stress. Mechanistically, S-nitrosylation is the reversible covalent binding of NO to a sulfhydryl group of a reactive cysteine, and it is precisely targeted to residues flanked by basic and acidic amino acids (28). As Cys4 in R5 meets these criteria, an in vitro S-nitrosylation assay was carried out using wild-type R5 and R5-C4S as the protein substrates, SNAP and S-nitrosoglutathione as the NO donors, and the biotin-switch method for detection of S-nitrosylated cysteines. In this assay, after blocking non-nitrosylated free thiol groups by methylation, S-nitrosylated cysteines are selectively identified by the cleavage of S-NO by ascorbate, followed by biotinylation of the free thiols, pull down of biotinylated proteins with streptavidin beads, and immunoblotting of eluates. As shown in Fig. 5A, we observed extensive nitrosylation of wild-type R5 preincubated with SNAP, whereas hardly any signal was detected with R5-C4S. Similar results were obtained with S-nitrosoglutathione as the NO donor (data not shown).
Nitrosylation of Plectin in Endothelial Cell Cultures—Nitric oxide plays a key regulatory role in endothelial cell function (29). To investigate whether plectin S-nitrosylation occurs in vivo, we incubated immortalized mouse renal endothelial cells in culture with SNAP and assayed plectin S-nitrosylation by the biotin-switch assay. As shown in Fig. 5B, a strong signal corresponding to full-length plectin could be detected in the streptavidin beads eluate, indicating S-nitrosylation of endogenous endothelial cell plectin. When ascorbate (1 mM) was added to the samples prior to blocking free (non-nitrosylated) thiol groups, no plectin signal was detectable in the eluate (Fig. 5B, Control), validating the assay. As an additional control, the membrane was stripped and overlaid with antibodies to endothelial NOS, which itself is a target of nitrosylation (30), revealing, as expected, the presence of the enzyme in both the starting cell lysate and the eluate recovered from the streptavidin beads (Fig. 5B, eNOS). We next examined whether plectin can be S-nitrosylated by an endogenous mechanism of NO generation. For this purpose, we treated cultured endothelial cells with PMA and subjected the cell lysates to the biotin-switch assay. PMA, an agonist of protein kinase C, has been shown to increase the expression and enzymatic activity of endothelial NOS in endothelial cells (31, 32). As shown in Fig. 5C, plectin was indeed S-nitrosylated by endogenously generated NO, whereas at basal NO levels (no PMA treatment), neither plectin nor endothelial NOS was nitrosylated.
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97:3:0 and 95:4:1 in untreated and 6-h SNAP-exposed wild-type cells, respectively, the corresponding values were 93:6:1 and 1:8:91 in plectin-deficient cells (Fig. 6, K-N). This strongly suggested an antagonistic role of plectin in S-nitrosylation-mediated vimentin filament collapse. |
DISCUSSION |
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TOPABSTRACTINTRODUCTIONEXPERIMENTAL PROCEDURESRESULTSDISCUSSIONREFERENCES |
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-helical content) and more stable (higher resistance to urea denaturation) than the mutant without any cysteines, which resembles wild-type R5 in the reduced form. Thus, we infer that the formation of disulfide bridges could indeed contribute to the stabilization of the protein. Furthermore, our data revealed that R5 cysteines can form intramolecular disulfide bridges between Cys1 and Cys4 and that this oxidized R5 species migrates faster than its reduced counterpart, suggesting conformational alterations of R5 upon oxidation. As the modeling of the R5 module revealed Cys1 to be partially buried in the core of the protein, the presence of the linker in the cross-linked R5 species probably caused a change in the local conformation, exposing the surface in the vicinity of Cys1 and promoting the formation of a disulfide bridge. Choi et al. (11) proposed that, in desmoplakin, each R domain represents an independent structure, with the three of them arranged as "beads-on-a-string." In a model proposed by Janda et al. (10) for the six R domains of plectin, the repeats are packed against each other in a way in which plectin R1–5 are arranged around a central R6 domain, forming a compact C-terminal globular or disk-like domain. It was suggested that, in addition to reciprocal hydrogen bond interactions mediated by hydrophobic residues on the surface of the repeats, linked cysteine residues could contribute to stabilizing this structure (10). Evidence in support of a compact packing of plectin R domains comes from electron microscopy, where the C-terminal domain was visualized as a globule 9 nm in diameter (9). Homology modeling of each of the six modules of plectin based on the known desmoplakin structure (50–73% sequence identity) yielded a cylinder-like configuration with an average size of 45 x 25 Å. Accommodation of six repeats within a 9-nm globule would indeed require tight packing of the repeats rather than a loose beads-on-a-string arrangement. R4 has a single cysteine in a position equivalent to Cys2 of R5; this cysteine can form disulfide bridges with cysteines of R5, linking these repeats together (Fig. 3B). We presume that corresponding cysteines in the other repeats could behave similarly. These findings lend further support to the model advanced by Janda et al. (10).
Comparison of Vimentin Binding of Plectin and Desmoplakin—In this study, the results of two experimental approaches, vimentin co-sedimentation and dot-blot overlay assays, clearly showed that R4 does not bind to vimentin, whereas R5 does. Furthermore, R4 did not co-localize with vimentin when overexpressed in PtK2 cells, nor did it cause the collapse of vimentin filaments (data not shown). This behavior is in full agreement with the results of a previous study in which the plectin vimentin-binding site was mapped to the linker region of R5 (14). IF binding of the corresponding linker regions has been shown for other plakin protein family members, such as envoplakin and periplakin (22). It should be noted, however, that an additional vimentin-binding site of plectin has been localized in its N-terminal actin-binding domain (33). On the other hand, Choi et al. (11) reported that each of the R domains of desmoplakin was able to bind to vimentin independently of the linker region. However, binding was very weak and occurred only if the R domains were present at a large molar excess, with a cooperative effect when all three repeats were present. There are two possible explanations for this apparent discrepancy. One is based on the assumption that proteins in the family are not redundant, but that each has a unique function. Thus, despite the overwhelming sequence similarity, desmoplakin could bind to vimentin in a different fashion or with different efficiency compared with plectin. In fact, plectin seems to have a higher affinity for vimentin than does desmoplakin, as concluded from the observation that plectin R5 co-sedimented with vimentin at a molar ratio of 1:1. The other explanation could be the different conditions under which the assays were performed. As we have shown, single repeats of plectin bind to vimentin only in their reduced state. However, it appears that oxidation of the preparations of desmoplakin single repeats was not prevented (11), and as a consequence, binding was probably not optimal.
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Our study has shown shows that Cys4 in R5 is highly reactive and that plectin is a target for nitrosylation in vitro and in vivo. S-Nitrosylation of cysteines leading to nitrosothiols is a biochemical post-translational modification of proteins that allows cells to respond to environmental changes in specific subcellular compartments and can modulate protein activity in a fashion analogous to phosphorylation (34). Despite the existence of cysteine residues in virtually all proteins, proteomic analysis of S-nitrosylated proteins in extracts from brain (35), mesangial cells (36), and intact endothelial cells (37) revealed that S-nitrosylation is restricted to a small number of proteins. Furthermore, S-nitrosylation is not a function of total cysteine content because, generally, only one single cysteine thiol in the protein is modified. The specificity is determined by structural motifs around the cysteine and the intracellular proximity to a NO generator, such as the enzyme NOS, which produces NO through the conversion of L-arginine into L-citrulline. Both requirements seem to be fulfilled by Cys4 in R5. First, the residues upstream (Gln-Glu) and downstream (Glu) of Cys4 perfectly match the consensus motif for S-nitrosylation. Second, although a comprehensive co-localization study of endogenous NOS and plectin in different cell types and tissues has not been reported to date, at least for muscle, there is strong evidence for intracellular proximity. The majority of NOS activity in skeletal muscle is associated with neuronal NOS, one of three isoenzymes differentially expressed in different cell types. Being targeted to 1-syntrophin, a dystrophin-associated protein, neuronal NOS is found at specialized structures on the surface membrane of muscle fibers, such as neuromuscular and myotendinous junctions and costameres (38). As a prominent component of neuromuscular and myotendinous junctions and costameres, plectin is also associated with the sarcolemma and has been found to co-localize with dystrophin (39, 40). Furthermore, there is evidence that plectin directly interacts with dystrophin and the dystrophin-associated transmembrane laminin receptor -dystroglycan.4
Role for Plectin R5 Cys4 in NO-induced IF Collapse?—Assessing whether S-nitrosylation stimulated in endothelial cells by exposure to NO donors has any effects on cellular cytoarchitecture, we found an interesting link between plectin deficiency and dynamic properties of the IF network, which is built up of vimentin in these cells. Whereas well spread cellular vimentin networks largely persisted in cells treated with the NO donor SNAP for up to 6 h in wild-type cells, we observed a progressive collapse of vimentin filament networks into perinuclear bundles in plectin-deficient cells. One explanation for this phenomenon could be that plectin protects vimentin against oxidation. Like all other type III IFs, vimentin contains a single cysteine residue at a highly conserved position (position 328 in the human protein). Several reports suggest that this residue is oxidized in vivo. In fact, upon exposure of fibroblast-like synoviocytes to H2O2, it is oxidized in preference to other cytoskeleton proteins (41). Furthermore, these cells show an increased susceptibility to vimentin collapse around the nucleus upon exposure to oxidative stress, and vimentin IFs in rheumatoid synoviocytes are more susceptible than those in normal synoviocytes, bearing a possible pathological significance (42). The proposal that synoviocyte vimentin becomes glutathionylated (41) has been confirmed recently by proteomics approaches showing that vimentin is glutathionylated in human T lymphocytes and rat aortic smooth muscle cells (43, 44). Furthermore, prostanylation of the vimentin thiol has been shown in mesangial cells (45) and its nitrosylation in endothelial cells (46).
By binding to vimentin, plectin may either sterically hinder access of oxidants and thus prevent or impede IF collapse or prevent vimentin molecules from assuming a confirmation enabling vimentin-vimentin disulfide bridge formation. There is evidence, at least from in vitro experiments, that disulfide-cross-linked vimentin homodimers or desmin/vimentin heterodimers are filament assembly-incompetent under nonreducing conditions (47–49). The proposed shielding effect of plectin on the vimentin thiol group may be further enhanced by the multitude of proteins bound to its surface as a consequence of its scaffolding function (3).
Alternatively, transient nitrosylation of Cys4 in R5 of plectin may promote its subsequent disulfide bond formation with the unique cysteine of vimentin, thereby preventing its bond formation with another vimentin molecule and eventually filament collapse. In fact, preliminary results indicate that R5 Cys4 of plectin can form disulfide bridges with the unique cysteine of vimentin in vitro.4
Another conceivable way that plectin might affect NO-induced IF network collapse involves protein phosphorylation. The reversible phosphorylation/dephosphorylation of IF subunit proteins has been well established as a cellular mechanism leading to the disassembly of IFs. Recently, we found that the collapse of keratin networks induced in keratinocytes by the protein phosphatase inhibitor okadaic acid proceeds considerably faster in plectin-deficient compared with wild-type cells (5) and that the same applies to the vimentin IF network of fibroblasts.4 Several groups have reported inhibition of protein phosphatases by endogenously produced reactive oxygen species, and it has been shown that exposure of purified protein phosphatases to NO donors leads to reversible enzyme inhibition (50–52). Furthermore, NO activates serine/threonine kinases, including cGMP-activated protein kinase, as well as various tyrosine kinases (53, 54). Thus, the NO-triggered accelerated collapse of the vimentin network in plectin-deficient endothelial cells, similar to okadaic acid-treated cells, could be caused by protein hyperphosphorylation effected by NO-activated phosphorylation. The mechanism by which plectin counteracts/antagonizes hyperphosphorylation of vimentin networks in wild-type endothelial cells remains to be shown. However, one can expect that, similar to other cell types, plectin influences the cytoarchitecture of the IF network and thereby exerts regulatory functions over certain signaling cascades (3, 5). The collapse of the vimentin filament network probably compromises the normal ability of endothelial cells to respond to an appropriate stimulus, causing vasodilation in smooth muscle. As such a dysfunction is thought to be a key event in the development of atherosclerosis and has also been shown to be of prognostic significance in predicting vascular events, including stroke and heart attacks (55), it will be of interest to investigate whether plectin integrity plays a role in these diseases. In conjunction with plectin knock-out mouse models, the immortalized plectin-deficient endothelial cell line described here should be useful in delineating such relations as well as providing new perspectives on cytolinker protein functions.
In conclusion, we have shown that cysteines proximal to the IF-binding site of plectin in R5 have the capability to form disulfide bridges with consequences on the conformational and biochemical properties of the repeat domain. The redox state of these cysteines was found to have an influence on vimentin binding affinity, and one of the cysteines was identified as a specific target for nitrosylation. The observation that plectin-deficient endothelial cells were more sensitive to NO donor-induced IF network collapse compared with wild-type cells suggests an antagonistic role of plectin in oxidative stress-mediated alterations of the cytoskeleton and a possible role of a plectin cysteine as a regulatory switch.
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FOOTNOTES |
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The on-line version of this article (available at http://www.jbc.org) contains supplemental "Experimental Procedures," Equations 1 and 2, Refs. 1–6, and Figs. S1 and S2.
1 Present address: College of Pharmacy, Salahaddin University, Erbil-Kurdistan Region, Federal Republic of Iraq.
2 To whom correspondence should be addressed: Dept. of Molecular Cell Biology, Max F. Perutz Laboratories, University of Vienna, Campus Vienna Biocenter, Dr. Bohr-Gasse 9, A-1030 Vienna, Austria. Tel.: 43-1-4277-52852; Fax: 43-1-4277-52854; E-mail: gerhard.wiche{at}univie.ac.at.
3 The abbreviations used are: IFs, intermediate filaments; R, repeat; DTT, dithiothreitol; BSA, bovine serum albumin; SNAP, S-nitroso-N-acetylpenicillamine; NOS, nitric-oxide synthase; PMA, phorbol 12-myristate 13-acetate.
4 R. Spurny, G. Rezniczek, M. Gregor, and G. Wiche, unpublished data.
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ACKNOWLEDGMENTS |
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TOPABSTRACTINTRODUCTIONEXPERIMENTAL PROCEDURESRESULTSDISCUSSIONREFERENCES |
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) R5 in 10 mM sodium phosphate buffer (pH 7.5) and 20 mM NaF at room temperature; middle panel, wild-type R5 at pH 7.5 (•), pH 9.0 (
), and pH 11.0 (
); lower panel, cysteine-free R5 at pH 7.5 (
), and pH 11.0 (
). B, urea-induced unfolding profiles monitored by far-UV CD and fluorescence spectroscopy. The fraction of unfolded protein observed by far-UV CD and fluorescence spectroscopy is plotted against the concentration of urea. Symbols represent the spectroscopic signals obtained, and solid lines represent the profiles obtained on the basis of Equations 1 and 2 as described under supplemental "Experimental Procedures." Closed symbols indicate normalized changes in ellipticity at 222 nm (•) and fluorescence emission at 340 nm after excitation at 290 nm (
) were coated onto microtiter plates at concentrations of 100 nM and overlaid with Eu3+-labeled recombinant vimentin at concentrations of 0.05–2 µM.Eu3+-Labeled vimentin bound to the different versions of R5 was measured as described under "Results." Scatchard plots of the binding data are shown.
