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Correspondence to James M. Ervasti: jervasti@umn.edu Abbreviations used in this paper: CH, calponin homology; MAP, MT-associated protein; MOM, mouse on mouse; MT, microtubule; wt, wild type.
ytolinkers are giant proteins that can stabilize cells by linking actin filaments, intermediate filaments, and microtubules (MTs) to transmembrane com- plexes. Dystrophin is functionally similar to cytolinkers, as it links the multiple components of the cellular cytoskeleton to the transmembrane dystroglycan complex. Although no direct link between dystrophin and MTs has been docu- mented, costamere-associated MTs are disrupted when dystrophin is absent. Using tissue-based cosedimenta- tion assays on mice expressing endogenous dystrophin or truncated transgene products, we find that constructs harboring spectrinlike repeat 24 through the first third of the WW domain cosediment with MTs. Purified Dp260, a truncated isoform of dystrophin, bound MTs with a K d of 0.66 µM, a stoichiometry of 1 Dp260/1.4 tubulin heterodimer at saturation, and stabilizes MTs from cold-induced depolymerization. Finally, - and -tubulin expression is increased 2.5-fold in mdx skeletal muscle without altering the tubulin–MT equilibrium. Collectively, these data suggest dystrophin directly organizes and/or stabilizes costameric MTs and classifies dystrophin as a cytolinker in skeletal muscle.
Dystrophin, the protein absent in patients with Duchenne muscular dystrophy (Hoffman et al., 1987), shows structural and functional similarities to cytolinkers, which suggests the hypoth­ esis that dystrophin performs a cytolinker role in muscle. Dystro­ phin’s large molecular mass of 427 kD, spectrinlike repeats, and ability to bind actin filaments via a tandem CH domain (Way et al., 1992) highlight three similarities with cytolinkers. Although dystrophin lacks a plakin repeat domain, dystrophin–intermediate filament interactions have been documented (Stone et al., 2005; Bhosle et al., 2006). Thus, the ability to link the actin and inter­ mediate filament cytoskeletons to the transmembrane dystro­ glycan complex (Suzuki et al., 1992) illustrates how dystrophin functions similarly to other cytolinkers. Finally, the muscle membrane fragility associated with the loss of dystrophin (Petrof et al., 1993) parallels the structural deficiencies observed in other cytolinker­deficient tissues, further demonstrating a close rela­ tionship between dystrophin and other cytolinkers. Collectively, these data support the hypothesis that dystrophin may function as a cytolinker in skeletal muscle.
Introduction The plakins are a class of giant cytolinker proteins that can link transmembrane protein complexes to the actin, intermediate fila­ ment, and microtubule (MT) cytoskeletons in various combi­ nations (Fuchs and Karakesisoglou, 2001). Plakins can bind actin filaments via tandem calponin homology (CH) domains, intermediate filaments via plakin repeat domains, and MTs through either a Gas2­related domain or a glycine/serine/ arginine domain (for review see Leung et al., 2002). The ability to cross­link multiple components of the cellular cytoskeleton allows cytolinkers to stabilize cells from mechanically induced damage. Mouse knockout studies exemplify the stabilizing effects of cytolinkers, as loss of the cytolinker plectin resulted in skin blistering and a form of muscular dystrophy (Andra et al., 1997), and ablation of BPAG­1, another cytolinker, caused skin blistering upon mechanical stimulation (Guo et al., 1995). and the restoration of costameric MT organization through virally mediated expression of a microdystrophin (Percival et al., 2007) indicates that dystrophin is necessary for proper costameric MT organization in skeletal muscle. Moreover, acute knockdown of ankyrin­B, a protein necessary for delivery of dystrophin to the sarcolemma and neuromuscular junction, caused the loss of costa­ meric MTs and aberrant MT organization in a subset of MTs underlying the neuromuscular junction (Ayalon et al., 2008).
In this study, we investigated the hypothesis that dystro­ phin directly interacts with costameric MTs. We confirmed that costameric MTs were disrupted in dystrophin­deficient skeletal muscle and showed endogenous dystrophin cosedimented with MTs in tissue homogenates. Using purified proteins, we found that the carboxyl­terminal two thirds of dystrophin bound MTs with a K d of 0.66 µM and stabilized MTs from cold­induced de­ polymerization. Finally, we documented a 2.5­fold increased expression of ­ and ­tubulin without alteration in the tubulin– MT equilibrium in mdx skeletal muscle. These results demon­ strate that dystrophin is a MT­associated protein (MAP) that stabilizes costameric MTs and functions as a costameric cyto­ linker in skeletal muscle.
these results confirm a role for dystrophin in the stabilization and MT derangement occurred before muscle cell necrosis and regeneration. proper organization of costameric MTs independent of muscle Bars: (A and B) 10 µm; (C and D) 20 µm. necrosis and regeneration. rod domain absent from Dp260. A small amount of Dp260 pel­ leted in the absence of MTs, but substantially more Dp260 shifted to the pellet fraction when MTs were present (Fig. 3 A). After sub­ tracting self­pelleting Dp260, Dp260 displayed a concentration­ dependent and saturable cosedimentation with a Dp260/­ tubulin heterodimer stoichiometry of 1:1.4 and a K d of 0.66 µM (Fig. 3 C). As predicted, DysN Term ­R 10 did not cosediment with MTs up to concentrations approaching 10 µM (Fig. 3, B and C). Next, we assessed how the presence of 1 µM Dp260 affected the tubulin–MT equilibrium in vitro. Dp260 had no significant effect on the fraction of tubulin in the MT fraction when incu­ bated at room temperature (67.3 ± 0.72% vs. 68.6 ± 1.3%). However, the presence of Dp260 significantly increased the fraction of tubulin retained in the MT pellet (33.6 ± 2.9% vs. 42.2 ± 2.0%) when MTs were induced to depolymerize by incu­ bating at 4°C. (Fig. 3, D and E). Collectively, these results dem­ onstrate that dystrophin directly binds and stabilizes MTs from cold­induced depolymerization.
Because misregulation of other MAPs can alter tubulin expression and MT stability (Harada et al., 1994; Takahashi et al., 2003), we investigated how the loss of dystrophin affects the regulation of tubulin expression and the tubulin–MT equi­ librium in skeletal muscle fibers. Tubulin levels in wt and mdx skeletal muscle extracts were examined by quantitative West­ ern blot analysis. With mAb B512, we observed no difference Next, we performed a tissue­based MT cosedimentation assay (Fig. 2 A) to determine whether dystrophin cosedimented with MTs. Under conditions that induced MT depolymeriza­ tion, virtually no muscle protein pelleted (Fig. 2 B). However, numerous proteins pelleted under MT­stabilizing conditions, and this fraction of proteins represents MTs and the MAPs of skeletal muscle (Fig. 2 B). We Western blotted each fraction obtained from the tissue cosedimentation assay performed on wt mice expressing full­length dystrophin or transgenic mdx mice expressing Dp260, microdystrophin (R4­23), or Dp71 (Fig. 2 C, right). Full­length dystrophin, Dp260, and R4­23 all pelleted with MTs, whereas Dp 71 did not (Fig. 2 C, left). By comparing the dystrophin domains present or absent in each construct (Fig. 2 C, right) along with each construct’s ability to cosediment with MTs, we suggest that spectrinlike repeat 24 through the first third of the WW domain encodes a novel MT­binding domain.
Figure 2. Dystrophin cosediments with MTs in skeletal muscle extracts. (A) Flowchart of tissue MT cosedimentation assay. (B) Coomassie blue–stained SDS-PAGE showing supernatant (S) and pellet (P) fractions in conditions that favored MT depolymerization or polymeriza- tion. The pellet fraction in the presence of MTs represents the MAP fraction of skeletal muscle. The molecular mass standards (given in kilo- daltons) are indicated on the left. (C, left) Western blot analysis of tissue cosedimenta- tion assay from skeletal muscle extracts of wt mice expressing dystrophin or mdx mice transgenically expressing Dp260, R4-R23, and Dp71. (right) Diagrammatic representation of constructs analyzed in tissue cosedimenta- tion. ABD, actin-binding domain; H, hinge region; W, WW domain; CR, cysteine-rich domain; CT, carboxy-terminal domain; MT BD, MT-binding domain. in ­tubulin expression in mdx skeletal muscle (Fig. 4, A and B). Because levels of ­ and ­tubulin are coregulated (Gonzalez­ Garay and Cabral, 1995), we investigated ­tubulin levels to determine whether ­tubulin is up­regulated in mdx skeletal muscle. ­Tubulin expression was elevated 2.5­fold in mdx skeletal muscle (Fig. 4, A and B), suggesting that expression of both ­ and ­tubulin is increased in mdx skeletal muscle.
Thus, we conclude that mAb DM1A is able to recognize a pop­ ulation of ­tubulin not detected by mAb B512. To examine MT stability in mdx skeletal muscle, we analyzed levels of tyrosinated ­tubulin, a marker of dynamic MTs (Gundersen et al., 1984, 1987), and acetylated ­tubulin, a marker of long­ lived MTs (Bulinski and Gundersen, 1991). The levels of tyro­ sinated ­tubulin were increased 2.5­fold in mdx extracts (Fig. 4, A and B), whereas the levels of acetylated ­tubulin were not (Fig. 4, A and B). The loss of dystrophin’s MT­stabilizing ability may explain why acetylated ­tubulin was not more abundant in mdx skeletal muscle extracts, but alterations in the tubulin–MT equilibrium could also explain the lack of more stable MTs. Therefore, we examined the tubulin–MT equilib­ rium in wt and mdx skeletal muscles and found that the loss of dystrophin did not affect the equilibrium (Fig. 4, C and D). Collectively, these results show that tubulins are misregulated in dystrophin­deficient skeletal muscle without affecting the tubulin–MT equilibrium. The loss of dystrophin’s MT­stabilizing ability likely explains why there are not more stabilized MTs even in the presence of more tubulin dimer in dystrophin­deficient skeletal muscle.
An indirect link between dystrophin and MTs mediated by ankyrin­B was recently shown to be important for proper trafficking of dystrophin and ­dystroglycan to the sarcolemma (Ayalon et al., 2008). However, costameric MTs are disorga­ nized in mdx skeletal muscle even in the presence of properly localized ankyrin­B (Ayalon et al., 2008). Because the MT­ and ankyrin­B–binding domains of dystrophin do not overlap (Fig. 5), our results and previous results suggest that dystrophin interacts with MTs in vivo through two distinct mechanisms. We propose that ankyrin­B delivers dystrophin to the sarcolemma depen­ dent on MTs and that dystrophin and ankyrin­B collaborate to stabilize and organize MTs in skeletal muscle.
Figure 3. Dp260 binds and stabilizes MTs in vitro. (A) Coomassie blue– stained SDS-PAGE showing the supernatant (S) and pellet (P) fractions of purified Dp260 in the presence and absence of MTs. Dp260 shifted to the pellet fraction when MTs were present, indicating that Dp260 binds MTs. (B) Coomassie blue–stained SDS-PAGE showing the supernatant and pellet fractions of DysN-R 10 in the presence and absence of MTs. The presence of MTs did not cause a shift of DysN-R 10 into the pellet fraction, indicating that DysN-R 10 does not bind MTs. (C) Concentration-dependent binding of Dp260 to taxol-stabilized MTs (2 µM tubulin) from three independent experiments. DysN-R 10 did not show significant MT-binding activity. (D) Coomassie blue–stained SDS-PAGE of the supernatant and pellet frac- tions of MTs induced to depolymerize by incubating at 4°C in the presence or absence of 1 µM Dp260. (E) Quantification of tubulin in the MT fraction when induced to depolymerize by incubating at 4°C (n = 6). The presence of 1 µM Dp260 significantly (t test; *, P ≤ 0.05) increased the amount of tubulin in the MT fraction, indicating that Dp260 stabilizes MTs. Error bars represent mean ± SEM. (A, B, and D) Molecular mass standards (given in kilodaltons) are indicated on the left. pathophysiology. For example, disorganized MTs are also asso­ ciated with Golgi mislocalization (Percival et al., 2007), which in combination, would likely lead to impaired trafficking of membrane­bound proteins and may explain the decreased levels of ­dystroglycan and the sarcoglycans at the sarcolemma of mdx skeletal muscle (Ohlendieck and Campbell, 1991). Because no MT knockout mouse has been generated, the exact function of MTs in skeletal muscle remains unknown. However, the impor­ tance of MTs in skeletal muscle biology is illustrated by the muscle weakness and increased levels of serum creatine kinase associ­ ated with colchicine toxicity in human patients (Boomershine, 2002; Caglar et al., 2003; Wilbur and Makowsky, 2004; Altman et al., 2007). Therefore, it is possible that derangement of the MT cytoskeleton contributes to some of the phenotypes associ­ ated with dystrophin deficiency.
Mice Control C57BL/6 and mdx mice were initially obtained from The Jackson Laboratory. The utrn  / and mdx/utrn  / mice were provided by D. Lowe (University of Minnesota, Minneapolis, MN). Mdx mice transgenically ex- pressing Dp260 and R4-R23 were provided by J. Chamberlain (Univer- sity of Washington, Seattle, WA), and the Dp71 line was provided by J. Rafael-Fortney (Ohio State University, Columbus, OH). All animals were housed and treated following guidelines set by the University of Minnesota Institutional Animal Care and Use Committee.
Antibodies The mAbs to -tubulin (B512), -tubulin (D66), tyrosinated tubulin (TUB 1-A2), and acetylated tubulin (6-11B-1) were purchased from Sigma-Aldrich. The mAb to -tubulin (DM1A) was purchased from Abcam. The mAb to dystrophin (Dys2) was purchased from Novacastra. The polyclonal antibody to dystrophin (Rb2) was described previously (Rybakova et al., 1996). Infrared dye–conjugated anti–mouse secondary antibodies were purchased from LI-COR Biosciences.
phenotype (Warner et al., 2002). Because Dp116 harbors only the MT­binding domain and one of two synemin­binding sequences, an inability to bind cytokeratin filaments and actin filaments likely explains why transgenic overexpression of Dp116 fails to rescue the mdx phenotype (Judge et al., 2006). Finally, the mild muscle phenotypes of  cyto ­actin (Sonnemann et al., 2006) or keratin 19 knockout mice (Stone et al., 2007) may be explained by dystro­ phin’s linkage with the remaining components of the cortical cyto­ skeleton. Collectively, these results support the hypothesis that dystrophin must bind all three components of the cellular cyto­ skeleton to function properly in skeletal muscle.
Figure 4. Mdx mice exhibit tubulin misregulation in the absence of an altered tubulin–MT equilibrium. (A) Repre- sentative Western blots of tubulin levels in skeletal muscle extracts from wt and mdx mice. (B) Quantification of tubu- lin levels from three wt and three mdx extracts. (C) Repre- sentative Western blots of tubulin (tub) and MT fractions of skeletal muscle extracts from wt and mdx mice using mAb DM1A. (D) Quantification of tubulin–MT equilibrium from five wt and four mdx tibialis anterior muscles. Loss of dystrophin does not affect the tubulin–MT equilibrium. Error bars represent mean ± SEM.
Figure 5. Diagrammatic representation of MT- and ankyrin-B–binding domains of dystrophin. Numbers indicate amino acids of full-length dys- trophin. H, hinge region; W, WW domain; CR, cysteine-rich domain; CT, carboxy-terminal domain; MT BD, MT-binding domain; Ank-B BD, ankyrin-B–binding domain.
36:824–826. Bulinski, J.C., and G.G. Gundersen. 1991. Stabilization of post­translational modifica­ tion of microtubules during cellular morphogenesis. Bioessays. 13:285–293.
MT cosedimentation analysis MT cosedimentation assay was performed as described by the manufactur- er’s instructions (Cytoskeleton, Inc). In brief, increasing amounts of purified protein were added to preformed MTs then centrifuged at 100,000 g for 30 min. The amount of free and bound protein was determined densito- metrically from Coomassie blue–stained gels of the supernatant and pel- leted fractions using imaging system software (UVP). The fraction of protein pelleting in the absence of MTs was subtracted from each data point. The resultant data from three independent experiments were fitted to a hyper- bolic binding equation using nonlinear regression analysis on Prism soft- ware (GraphPad Software, Inc.).
221,115 M  1 cm  1 for DysN-R 10 as predicted by the Expert Protein Analy- sis System proteomics server (Swiss Institute of Bioinformatics).
Immunofluorescence analysis To analyze the MT lattice in dystrophic animal models, hindlegs of wt (3 and 8 wk), mdx (3, 5, and 8 wk), utrn  / (8–10 wk), and mdx:utrn  / (3, 5, and 8 wk) mice were skinned, cut as close as possible to the body, and fixed at room temperature for 2 h with 4% para-formaldehyde in phos- phate buffer. They were stored in phosphate buffer until the extensor digito- rum longus muscle was dissected and separated with fine forceps into mostly single fibers. These were transferred to a 24-well tissue culture plate and incubated with mouse on mouse (MOM)–blocking buffer (Vector Labo- ratories) for 2 h at room temperature. Blocking buffer and every subse- quent buffer for incubation or washing contained 0.04% saponin for permeabilization and 0.05% sodium azide. Fibers were incubated over- night with mouse antitubulin (DM1A, 1:500; or B512, 1:4,000) in MOM diluent, washed three times for 20 min, and stained with 1:500 dilutions of Alexa Fluor 488 anti–mouse and Alexa Fluor 568 anti–rabbit secondary antibodies (Invitrogen) in MOM diluent for 2 h at room temperature. After three 20-min washes, one of which contained the nuclear stain Hoechst 33342 (Sigma-Aldrich) at 2 µg/ml, fibers were mounted onto a glass slide in a drop of Vectashield (Vector Laboratories). Confocal images were cap- tured with a 63× NA 1.4 oil immersion lens on a TCS SP5 confocal micro- scope (Leica) in the Light Imaging Section of the National Institute of Arthritis and Musculoskeletal and Skin Diseases. Gain and laser power set- tings were adjusted to avoid saturation and use the whole linear range of fluorescence intensity. Unless specified, the parameters were adjusted for each new fiber imaged. The raw TIF images were transferred to a com- puter (Macintosh G5; Apple), opened in Photoshop (CS2; Adobe), assem- bled into montages, and adjusted for brightness when needed. The final illustrations give a faithful representation of the collected images.
Caglar, K., Z. Odabasi, M. Safali, M. Yenicesu, and A. Vural. 2003. Colchicine­ induced myopathy with myotonia in a patient with chronic renal failure.
Interactions of intermediate filament protein synemin with dystrophin and utrophin. Biochem. Biophys. Res. Commun. 346:768–777. Boomershine, K.H. 2002. Colchicine­induced rhabdomyolysis. Ann. Pharmacother.
100,000 g for 30 min at 25°C. The soluble portion (tubulin containing) was saved for analysis, whereas the pelleted portion (MT fraction) was re- suspended in 1 mL of 1% SDS buffer then boiled for 10 min. The pellet fraction was centrifuged at 13,000 g for 10 min, and the soluble portion was saved for analysis. 25 µl of the tubulin and MT fraction was analyzed and quantified via Western blot using mAb DM1A on the infrared imaging system (Odyssey). Statistical analysis All data are presented as mean ± SEM. Comparison between groups was performed using a t test with significance defined as P ≤ 0.05.
Muscle­specific expression of insulin­like growth factor I counters mus­ cle decline in mdx mice. J. Cell Biol. 157:137–148. Bhosle, R.C., D.E. Michele, K.P. Campbell, Z. Li, and R.M. Robson. 2006.
135:1189–1200. Barton, E.R., L. Morris, A. Musaro, N. Rosenthal, and H.L. Sweeney. 2002.
1997. Targeted inactivation of plectin reveals essential function in main­ taining the integrity of skin, muscle, and heart cytoarchitecture. Genes Dev. 11:3143–3156. Ayalon, G., J.Q. Davis, P.B. Scotland, and V. Bennett. 2008. An ankyrin­based mech­ anism for functional organization of dystrophin and dystroglycan. Cell.
Altman, A., M. Szyper­Kravitz, and Y. Shoenfeld. 2007. Colchicine­induced rhabdomyolysis. Clin. Rheumatol. 26:2197–2199. Andra, K., H. Lassmann, R. Bittner, S. Shorny, R. Fassler, F. Propst, and G. Wiche.
This work was supported by the National Institutes of Health Training Program in Muscle Research (AR007612), a grant from the National Institutes of Health (AR042423), and Gregory Marzolf Muscular Dystrophy Fellow- ships to J.L. Humston and K.W. Prins. K.W. Prins is a member of the Medical Scientist Training Program at the University of Minnesota. E. Ralston, V. Tate, and A. Mehta were supported by the National Institutes of Health Intramural Research Program.
We would like to thank Dr. Kevin Sonnemann for generating the Dp260 baculovirus expression construct and Dr. Sonnemann and Thomas Cheever for their helpful discussions.
Suzuki, A., M. Yoshida, H. Yamamoto, and E. Ozawa. 1992. Glycoprotein­ binding site of dystrophin is confined to the cysteine­rich domain and the first half of the carboxy­terminal domain. FEBS Lett. 308:154–160. Takahashi, M., H. Shiraishi, Y. Ishibashi, K.L. Blade, P.J. McDermott, D.R.
Leung, C.L., K.J. Green, and R.K. Liem. 2002. Plakins: a family of versatile cytolinker proteins. Trends Cell Biol. 12:37–45. Ohlendieck, K., and K.P. Campbell. 1991. Dystrophin­associated proteins are greatly reduced in skeletal muscle from mdx mice. J. Cell Biol. 115:1685–1694. Ohlendieck, K., J.M. Ervasti, K. Matsumura, S.D. Kahl, C.J. Leveille, and K.P.
Sonnemann, K.J., D.P. Fitzsimons, J.R. Patel, Y. Liu, M.F. Schneider, R.L. Moss, and J.M. Ervasti. 2006. Cytoplasmic gamma­actin is not required for skeletal muscle development but its absence leads to a progressive myop­ athy. Dev. Cell. 11:387–397.
Utrophin binds laterally along actin filaments and can couple costameric actin with sarcolemma when overexpressed in dystrophin­deficient mus­ cle. Mol. Biol. Cell. 13:1512–1521. Rybakova, I.N., J.L. Humston, K.J. Sonnemann, and J.M. Ervasti. 2006.
Rybakova, I.N., J.R. Patel, K.E. Davies, P.D. Yurchenco, and J.M. Ervasti. 2002.
19:10694–10705. Rybakova, I.N., K.J. Amann, and J.M. Ervasti. 1996. A new model for the inter­ action of dystrophin with F­actin. J. Cell Biol. 135:661–672.
Ralston, E., Z. Lu, and T. Ploug. 1999. The organization of the Golgi complex and microtubules in skeletal muscle is fiber type­dependent. J. Neurosci.
S.C. Froehner. 2007. rAAV6­microdystrophin rescues aberrant Golgi complex organization in mdx skeletal muscles. Traffic. 8:1424–1439. Petrof, B.J., J.B. Shrager, H.H. Stedman, A.M. Kelly, and H.L. Sweeney. 1993.
Menick, D. Kuppuswamy, and G. Cooper IV. 2003. Phenotypic conse­ quences of beta1­tubulin expression and MAP4 decoration of micro­ tubules in adult cardiocytes. Am. J. Physiol. Heart Circ. Physiol. 285:H2072–H2083. Warner, L.E., C. DelloRusso, R.W. Crawford, I.N. Rybakova, J.R. Patel, J.M.
Judge, L.M., M. Haraguchiln, and J.S. Chamberlain. 2006. Dissecting the signal­ ing and mechanical functions of the dystrophin­glycoprotein complex.
Yoshitake, Y. Takei, T. Noda, and N. Hirokawa. 1994. Altered microtubule organization in small­calibre axons of mice lacking tau protein. Nature. 369:488–491. Harper, S.Q., M.A. Hauser, C. DelloRusso, D. Duan, R.W. Crawford, S.F. Phelps, H.A. Harper, A.S. Robinson, J.F. Engelhardt, S.V. Brooks, and J.S.
Gennerich, A., and R.D. Vale. 2009. Walking the walk: how kinesin and dynein coordinate their steps. Curr. Opin. Cell Biol. 21:59–67. Gonzalez­Garay, M.L., and F. Cabral. 1995. Overexpression of an epitope­ tagged beta­tubulin in Chinese hamster ovary cells causes an increase in endogenous alpha­tubulin synthesis. Cell Motil. Cytoskeleton. 31:259–272. Gundersen, G.G., M.H. Kalnoski, and J.C. Bulinski. 1984. Distinct populations of microtubules: tyrosinated and nontyrosinated alpha tubulin are distrib­ uted differently in vivo. Cell. 38:779–789. Gundersen, G.G., S. Khawaja, and J.C. Bulinski. 1987. Postpolymerization de­ tyrosination of alpha­tubulin: a mechanism for subcellular differentiation of microtubules. J. Cell Biol. 105:251–264. Guo, L., L. Degenstein, J. Dowling, Q.C. Yu, R. Wollmann, B. Perman, and E.
Fuchs, E., and I. Karakesisoglou. 2001. Bridging cytoskeletal intersections.
J. Muscle Res. Cell Motil. 20:383–393.
Expression of the N­terminal domain of dystrophin in E. coli and demon­ stration of binding to F­actin. FEBS Lett. 301:243–245. Wilbur, K., and M. Makowsky. 2004. Colchicine myotoxicity: case reports and literature review. Pharmacotherapy. 24:1784–1792. Williams, M.W., and R.J. Bloch. 1999. Differential distribution of dystrophin and beta­spectrin at the sarcolemma of fast twitch skeletal muscle fibers.
Ervasti, and J.S. Chamberlain. 2002. Expression of Dp260 in muscle teth­ ers the actin cytoskeleton to the dystrophin­glycoprotein complex and partially prevents dystrophy. Hum. Mol. Genet. 11:1095–1105. Way, M., B. Pope, R.A. Cross, J. Kendrick­Jones, and A.G. Weeds. 1992.
D.M. Toivola, J.A. Ursitti, M.B. Omary, and R.J. Bloch. 2007. Absence of keratin 19 in mice causes skeletal myopathy with mitochondrial and sarco­ lemmal reorganization. J. Cell Sci. 120:3999–4008.
Dystrophin is a microtubule-associated protei..