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Vol. 20, Issue 1, 400-409, January 1, 2009
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*Muscle Development Unit, Children's Medical Research Institute, Westmead, NSW, Australia; University of Western Sydney, Parramatta, NSW, Australia; ||Faculty of Medicine, University of Sydney, Sydney, NSW, Australia; ¶Department of Physiology, University of Melbourne, Parkville, VIC, Australia; #Institute for Molecular Biosciences, University of Queensland and Centre for Microscopy and Microanalysis, Brisbane, QLD, Australia; @Oncology Research Unit, The Children's Hospital at Westmead, Westmead, NSW, Australia; **Department of Pharmacology, School of Medical Sciences, University of New South Wales, Sydney, NSW, Australia; and Department of Anatomy, School of Medical Sciences, University of New South Wales, Sydney, NSW, Australia
Submitted June 18, 2008;
Revised October 17, 2008;
Accepted October 31, 2008
Monitoring Editor: Thomas D. Pollard
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ABSTRACT |
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INTRODUCTION |
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TOPABSTRACTINTRODUCTIONMATERIALS AND METHODSRESULTSDISCUSSIONCONCLUSIONSREFERENCES |
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-helical groove of the actin filament in a head-to-tail orientation and are important components of the actin microfilament cytoskeleton (Perry, 2001
). The Tm isoforms function in a variety of different cellular pathways in a range of cell and tissue types and are developmentally regulated (Gunning et al., 2008). In skeletal muscle, Tm isoforms function as a component of the contractile apparatus, regulating the actin–myosin interaction to facilitate sarcomeric contraction. Only three striated muscle Tm isoforms are required to fulfil this function; whereas the majority of the Tm isoforms are used by the actin cytoskeleton and contribute to the diversity of actin filament function (Gunning et al., 2008).
Tm isoforms are functionally distinct and sort to specific compartments within the cell (Gunning et al., 2008). Studies show that Tm isoforms protect actin filaments in an isoform-specific manner from the severing action of gelsolin (Ishikawa et al., 1989) and depolymerization by ADF/cofilin (Bernstein and Bamburg, 1982; Ono and Ono, 2002; Bryce et al., 2003). Tm isoforms can differentially regulate myosin enzymology and mechanochemistry (Fanning et al., 1994) and also the sorting of myosin motors (Tang and Ostap, 2001; Bryce et al., 2003). This compartmentalization is best described in neuronal cells (reviewed in Gunning et al., 2008). During neuronal development, specific Tm isoforms sort to the axonal shaft and growth cone. On differentiation, additional Tm isoforms are expressed and Tms relocalize to form distinct compartments in the axon, soma, dendrite, and presynaptic terminals (Had et al., 1993; Weinberger et al., 1993; Hannan et al., 1995, 1998; Schevzov et al., 1997). This compartmentalization is not restricted to neuronal cells. During the G1 phase of the cell cycle in fibroblasts, some Tm isoforms locate to stress fibers, whereas Tm5NM2 remains associated with the Golgi (Percival et al., 2004) and additional isoforms containing the 
-TM gene 9a exon localize to a perinuclear compartment (Schevzov et al., 2005a). Similar phenomena have been described in epithelial cells, in which Tm5a and 5b localize to the apical surface of cultured cells, Tm2 and Tm3 are present at the basolateral membrane and isoforms generated from the -TM gene are found in the central cytoplasm (Dalby-Payne et al., 2003). Sorting of functionally distinct Tm isoforms provides a mechanism for the spatial regulation of different actin filament populations.
Cytoskeletal Tm isoforms also have been detected in striated muscle. We demonstrated previously that a cytoskeletal Tm from the -TM gene, Tm5NM1, localizes to the sarcolemma and to a region adjacent to the Z-line in skeletal muscle fibers (Kee et al., 2004). In addition, we have shown that Tm4, the single product of the 
-TM gene, is also found in this region and in longitudinal filaments in regenerating or repairing muscle (Vlahovich et al., 2008). Both Tm5NM1 and Tm4 colocalize with a -actin cytoskeleton at these sites, suggesting a role for these filament systems in stabilization of the muscle fibers and linking myofibrillar networks to the membrane systems. In this study, we show that Tm5NM1 and Tm4 define separate filament systems and that Tm4 associates with the terminal sarcoplasmic reticulum and other tubulovesicular structures in the I-band and other regions of the muscle. Ablation of Tm5NM1 expression in the skeletal muscles of a -TM exon 9d knockout (KO) mouse indicates that Tm5NM1 plays a role in maintenance of normal excitation–contraction coupling.
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MATERIALS AND METHODS |
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TOPABSTRACTINTRODUCTIONMATERIALS AND METHODSRESULTSDISCUSSIONCONCLUSIONSREFERENCES |
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: 9d (sheep polyclonal antibody) recognizes the 9d exon from the -TM gene corresponding to Tm5NM1 in skeletal muscle and WD4/9d (rabbit polyclonal antibody) recognizes Tm4. Other primary antibodies used were: dihydropyridine receptor (mouse monoclonal, MAB47; Millipore Bioscience Research Reagents, Temecula, CA) and calsequestrin (mouse monoclonal, clone 51; BD Biosciences, Franklin Lakes, NJ). Secondary antibodies used were 488-conjugated goat anti-rabbit, 594-conjugated goat anti-mouse, and 594-conjugated donkey anti-sheep (Alexa Fluor; Invitrogen, Carlsbad, CA). Goat anti-rabbit and goat anti-mouse horseradish peroxidase (HRP)-conjugated antibodies (Bio-Rad, Hercules, CA) were used for Western blot analysis.
Mice
All animal experiments were performed in accordance with institutional and National Health and Medical Research Council guidelines. Mice were generated as described in Schevzov et al. (2008). A knockout construct was designed to specifically delete exon 9d-containing isoforms from the -TM gene. Targeted 129X1/SvJ ES cells were used to generate heterozygous mice (129X1/SvJ background; Tm5/9dneo mice). The same KO construct was electroporated into Bruce 4 C57Bl/6 ES cells. Heterozygous mice (C57Bl/6 background) were bred with mice carrying a cytomegalovirus-Cre recombinase transgene (C57Bl/6 background) (Schwenk et al., 1995) and then bred onto the C57Bl/6JArc background (Tm5/9d/89 mice) for >10 generations. All data presented is from mice of the Tm5/9dneo line and corresponding wild-type strain (129X1SvJ) unless otherwise indicated. All studies were performed on 2- to 3-mo-old male mice, and control wild-type (WT) mice were aged matched to within 1 wk of the KO mice.
Immunohistochemistry
Mouse muscles were fixed in 4% paraformaldehyde (PFA) (hindlimb muscles were stretched and held during fixation) and infused with 1.8 M sucrose/20% polyvinylpyrrolidone as described by Vlahovich et al. (2008). Semithin (0.5- to 0.8-µm) sections were cut at –60°C using an Ultracut UCT ultramicrotome (Leica, Wetzler, Germany) equipped with an EM FCS cryochamber (Leica). Sections were blocked and incubated with primary (9d, 1:50; -actinin, 1:500) and secondary antibodies (goat anti-mouse, 1:2000; donkey anti-sheep 1:1000) and viewed using either a confocal laser scanning microscope (oil immersion 63x objective, model TCS SP2; Leica) or standard fluorescent microscopy (BX51 microscope; Olympus, Tokyo, Japan).
Immunogold Labeling
Extensor digitorum longus (EDL) muscles were fixed in 4% PFA/0.1% glutaraldehyde/phosphate buffer, pH 7.2, for 1 h. Ultrathin cryosections were prepared according to Griffiths et al. (1984).
Immunogold labeling was carried out on an EM IGL Automated Immunogold Labeling System (Leica). Grids were blocked for 15 min in 50 mM glycine, 30 min in protein block, and incubated in primary antibodies (Tm4, 1:75; calsequestrin, 1:2000) diluted in immunoincubation buffer (0.25% bovine serum albumin [BSA]/12.5 mM sodium azide/phosphate-buffered saline [PBS], pH 7.4) overnight at 4°C. Grids were washed in immunoincubation buffer, incubated in gold-conjugated secondary antibodies (protein A gold 5 nm or goat anti-rabbit 10 nm gold [1:20], goat anti-mouse 20 nm gold [1:20] diluted in immunoincubation buffer; British BioCell International, Cardiff, United Kingdom) for 2 h at room temperature, washed in immunoincubation buffer, and fixed in 2% glutaraldehyde/PBS for 5 min. Grids were washed in PBS and water, and then they were embedded for 15 min in 1% methylcellulose:3% aqueous uranyl acetate diluted 9:1. Immunolabeled tissue was examined on a Philips CM10 transmission electron microscope. Quantitation of Tm4 and calsequestrin immunogold label was performed on three different grids with 10 random fields (each field containing at least 1 typical triad structure) counted per grid. The number of gold particles on each structure was counted as a percentage of the total number of gold particles in the whole field, and an average of the 10 fields/grid was calculated. Data are represented as mean ± SEM of the counts from the three grids/mouse line.
Isolation of Membrane Fractions
Membrane fractions from skeletal muscle were isolated using the method described by Saito et al. (1984). Membrane fractions at the interfaces between the discontinuous sucrose gradients (45, 38, 32, 27% sucrose) were collected, centrifuged for 2 h at 20,000 x gmax and the pellets were processed for cryo-electron microscopy (EM) for immunogold labeling.
Muscle Fiber Isolation and Analysis for Immunohistochemistry
Isolated muscle fibers from flexor digitorum brevis (FDB) muscle were isolated and cultured as described previously (Hernandez-Deviez et al., 2006). Primary (9d, 1:50; Tm4, 1:200; dihydropyridine receptor [DHPR], 1:200) and secondary (goat anti-mouse, 1:2000; goat anti-rabbit, 1:1000; donkey anti-sheep, 1:1000) antibody incubations were carried out at 37°C for 60 min or overnight at 4°C. Fibers were mounted and viewed using an LSM 510 META confocal microscope system (Carl Zeiss, Sydney, NSW, Australia). Single optical sections were captured using a Plan apochromatic 63x 1.4 numerical aperture oil immersion objective.
Detection and Quantitation of T-Tubule Dysmorphology
EDL muscles were fixed overnight in Karnovsky's fixative, processed in a LYNX tissue processor (Electron Microscopy Sciences, Hatfield, PA), and embedded in TAAB low viscosity resin. Ultrathin sections were stained with uranyl acetate and lead citrate and viewed in a Phillips CM10 transmission electron microscope. The T-tubule dysmorphology was calculated from a total of 1000 junctional membrane structures from muscles of at least three different mice of each genotype (Komazaki et al., 2003).
Western Blot Analysis
Protein extracts from mouse muscle were prepared and 12.5% SDS-polyacrylamide gel electrophoresis (PAGE) gels run as described by Kee at al. (2004). Coomassie-stained gels were used to verify equal protein loading. Protein was transferred onto polyvinylidene difluoride membranes (Millipore, Sydney, NSW, Australia), blocked in skim milk and incubated with primary (CG3, 1:1000; Tm4, 1:500; 9d, 1:500) and secondary antibodies (HRP-labeled secondary 1:10,000) according to Kee et al. (2004). Protein detection was performed using the SuperSignal West Pico chemiluminescence kit (Pierce Chemical, Rockford, IL).
Myosin Heavy Chain (MyHC) Isoform Gel Electrophoresis
MyHC isoform composition of EDL muscle was determined as described by Nair-Shalliker et al. (2004).
In Vivo Strength Tests
Whole animal strength and fatigability were measured according to the test procedure outlined in Joya et al. (2004). In brief, this test required the mice to pull themselves on top of a suspended rod with muscle weakness based on the mean percentage of passes over 15 trials in 3 min. Forearm strength was assessed using a dynamometer constructed according to the design of Smith et al. (1995). The mouse is suspended by the tail and the mouse grasps a horizontal bar. The bar is steadily moved away from the animal with increasing force and the force with which the animal releases the bar and the time taken for this to occur is recorded. This is repeated 10 times/mouse for four consecutive days and the last 2 day's readings are averaged.
Isolated Muscle Contractile Measurements
EDL muscles were carefully removed tendon-to-tendon from anesthetized (100 mg/kg body weight ketamine/10 mg/kg body weight xylazine), 10- to 12-wk-old mice for in vitro contractile measurements according to the method of Gregorevic et al. (2004). Optimal muscle length was determined with digital calipers during a series of isometric twitch contractions. Maximum isometric tetanic force was determined from the plateau of the frequency-force curve (1–200 Hz) and expressed as force/cross-sectional area (CSA). CSA was determined as described previously (Lynch et al., 2001).
Single Fiber Contractile Measurements
Single mechanically skinned fibers were prepared as described by Stephenson and Williams (1981). Mechanically skinned fibers were attached at one end to a piezoresistive force transducer (AE801; SensoNor, Horten, Norway), and the other end was fixed to a micromanipulator. Determination of the sarcoplasmic reticulum (SR) properties has been described previously (van der Poel and Stephenson, 2007). Ca2+ release from the SR was estimated from the relative areas under the caffeine-induced force response (CIFR) (van der Poel and Stephenson, 2007). Because the area under the CIFR is proportional to the SR loading times, the relative area under the CIFR was used to estimate the amount of Ca2+ in the SR (van der Poel and Stephenson, 2007). The percentage of Ca2+ lost from the SR due to the passive leak over a 60-s period was also assessed (van der Poel and Stephenson, 2007). After measurement of SR function the properties of the contractile apparatus were examined. Fibers were placed in a maximum Ca2+-activating solution (pCa 
4.5) to obtain maximum Ca2+-activated force. Fibers were then exposed to activating solutions of progressively higher [Ca2+], and the force response generated at each pCa was expressed as a percentage of the interpolated values for maximum Ca2+-activated force (van der Poel and Stephenson, 2002). Data points were fitted with a Hill equation producing two parameters: the pCa50 (i.e., pCa that produces half-maximum force) and the nH (i.e., the Hill coefficient, indicative of the steepness of the force–pCa relationship).
Depolarization-induced contractile responses (DICR) were also performed to examine T-tubule function (Plant and Lynch, 2002). Mechanically skinned muscle fibers were polarized with potassium hexamethylenediamine-tetraacetic acid (HDTA). The T-tubular membrane system was then depolarized with Na-HDTA, causing a transient DICR. Muscle fibers were then repolarized with K-HDTA before eliciting another Na-HDTA depolarization. This protocol was repeated to produce DICRs until the peak amplitude of the DICR had reached <50% of the initial value.
Statistical Analysis
Statistical significance was tested at p < 0.05 levels using Student's t test.
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RESULTS |
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TOPABSTRACTINTRODUCTIONMATERIALS AND METHODSRESULTSDISCUSSIONCONCLUSIONSREFERENCES |
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; Vlahovich et al., 2008). To determine whether Tm5NM1 and Tm4 are present in the same filament or define distinct filamentous populations and whether they are associated with organelles in the Z-line region, colocalization studies were performed. Costaining of stretched soleus muscle with the Tm-specific antibodies shows that Tm4 and Tm5NM1 occupy similar Z-line adjacent positions (Figure 1, A–C). However, the staining of the two Tms is not identical; Tm4 has some staining over the Z-line, whereas Tm5NM1 seems to localize exclusively to the Z-line adjacent region.
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To gain further insight into Tm4 localization, immunogold labeling and EM was performed on frozen sections of mouse EDL muscle (Figure 2). Immunolabeling of EDL sections showed that Tm4 localizes mainly to the sarcoplasmic reticulum and tubulovesicular structures around the triad region of the I-band (Figure 2, A, D, and G; black arrows). To confirm the identity of the SR localization, muscle sections were labeled for calsequestrin, a marker of the terminal cisternae of the SR (Figure 2, B, E, and H). Colabeling of the sections showed that Tm4 and calsequestrin colocalize to the same membrane systems (Figure 2, C, F, and I). To determine the exact localization of Tm4, quantitation of gold particles was performed on WT muscle (Table 1). The quantitative data demonstrates that calsequestrin and Tm4 localize to the same structures with approximately 30% of calsequestrin and 22% of Tm4 found on the terminal SR membranes, and approximately 70% of calsequestrin and 42% of Tm4 on tubulovesicular structures or sarcoplasmic reticulum in the I-band region (Table 1). No Tm4 labeling was detected on the T-tubules. These results indicate that the Tm4-defined cytoskeleton is in close association with the SR membrane system. Some Tm4 label was also found on nonmembranous areas of the A- and I-bands (Table 1). Some of this label could be nonspecific but some of the A-band label may be associated with longitudinal Tm4 filaments that we have described previously (Vlahovich et al., 2008). A primary antibody omission negative control for each antibody demonstrated no label for the calsequestrin control and a mean of 0.7 gold particles per field for the Tm4 control (data not shown).
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9d. Unfortunately, no specific labeling was observed perhaps due to the low abundance of Tm5NM1 in muscle (Kee et al., 2004). To overcome this problem, immuno-EM was performed on muscle from a Tm5NM1 transgenic mouse line (Tm5/52) that expresses the human protein at >5 times the endogenous protein (Schevzov et al., 2005a). Unfortunately, again no specific labeling was observed.
Tm4 association to SR membranes was also examined by muscle cell membrane fractionation. Membrane fractions were generated by ultracentrifugation of homogenates of pooled hindlimb muscle through a discontinuous sucrose gradient. The pellet of the SR fraction (fraction 4) was processed for immuno-EM. This fraction has been shown previously to be enriched for the terminal SR membranes (Saito et al., 1984). Immuno-EM staining of Tm4 and calsequestrin revealed the presence of Tm4 associated with calsequestrin containing vesicles (Figure 2, J–L) confirming that Tm4 is associated with the terminal SR.
Other Tm Isoforms from the -TM Gene and Tm4 Do Not Compensate for the Absence of Tm5NM1 in KO Mice
To examine the role of Tm5NM1 in skeletal muscle fibers, we analyzed mice that are null for Tm5NM1 and Tm5NM2 (Schevzov et al., 2008). Because Tm5NM2 is not expressed in skeletal muscle (Kee et al., 2004; Percival et al., 2004), these mice allow us to study the role of Tm5NM1, exclusively, in skeletal muscle. Using the 9d antibody that recognizes Tm5NM1 and Tm5NM2, Western blotting was performed on EDL and FDB muscles to confirm the absence of Tm5NM1 protein in KO muscles (Figure 3A). Using an antibody (CG3) that recognizes the 1b exon of the -TM gene, present in all cytoskeletal isoforms from this gene, we found that other -TM gene isoforms are not up-regulated to compensate for the lack of Tm5NM1 in skeletal muscle (Figure 3A). The typical localization of Tm5NM1 at the Z-line adjacent region of EDL muscle (Figure 3B) is lost in skeletal muscle from KO mice (Figure 3C). However, the localization of Tm4 to the terminal SR (black arrows in Figure 3D) and other membrane and nonmembranous structures remained unchanged in the absence of Tm5NM1 (Figure 3 and Table 1).
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). Triads were examined for alterations to T-tubule structure (Figure 4). The level of T-tubule dysmorphology was increased in Tm5NM1 KO muscle (Figure 4, B and C) in comparison with WT muscles (Figure 4A). In electron micrographs, the T-tubule shows up as a membrane structure orientated in parallel to the Z-line (Figure 4A, T-tubules labeled by an asterisk, SR by black arrows, and Z-line by white arrow) and flanked by the terminal SR. The T-tubules were considered abnormal if they were orientated diagonally (Figure 4B, asterisk) or were bent or crooked (Figure 4C, asterisk). Quantitation of T-tubule dysmorphology was performed on a total of 1000 junctional structures from WT and KO muscle per mouse (Figure 4D). We found a small but significant increase (3.2 times) in morphologically abnormal T-tubules in the KO muscle compared with WT skeletal muscle.
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30 Hz, p < 0.05), but not higher frequencies (Figure 5E). At physiological stimulation frequencies (75–100 Hz for the EDL) force output from the KO muscle was not different from WT muscle. This is consistent with the lack of improved muscle strength in this mouse (whole body muscle strength and fatigability test and forearm grip strength; Supplemental Figure 2). The time to reach peak twitch contraction was also more rapid in the KO muscle (Figure 5C), but relaxation time was similar to WT muscle (Figure 5D). Lack of an effect on contraction relaxation time is an indication that Ca2+ reuptake from the SR is not altered in the KO mice. Taken together, the data are consistent with either an increase in Ca2+ release from the SR with each single activation or alteration to T-tubule function in the KO mice such that each electrical stimulus leads to greater T-tubule activation and increased twitch force.
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DISCUSSION |
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TOPABSTRACTINTRODUCTIONMATERIALS AND METHODSRESULTSDISCUSSIONCONCLUSIONSREFERENCES |
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). However, due to the differences in the Tm5NM1 and Tm4 staining in the immunofluorescent images (Figure 1), it is most likely that these filaments are composed of homodimers of each Tm isoform.
The association of Tm4 with elements of the SR suggests that Tm4-defined actin filaments may have a role in the function of the SR system. Components of the spectrin-ankyrin-actin cytoskeleton have been localized to the SR in cardiac cells (Bennett and Baines, 2001). In erythrocytes, a nonmuscle tropomyosin is a critical element of the spectrin–actin cytoskeleton as without it the integrity of the erythrocyte plasma membrane is severely compromised (An et al., 2007). In a similar manner, Tm4 may stabilize an actin–spectrin cytoskeleton in the SR and help maintain the integrity of the SR membrane.
Tm5NM1 Impacts on the Structure of the T-Tubules
The accurate formation and alignment of the T-tubules and SR systems as well as preservation of the structure of these elements is critical to muscle function. The development and maturation of the SR and T-tubules takes place over several weeks and occurs in three discrete steps: 1) the independent differentiation of the membrane compartments of the T-tubules and SR, 2) the formation of the SR/T-tubule junctional triad, and 3) the alignment of the triads with the myofibrils and the transverse orientation of the T-tubules (Flucher et al., 1992; Takekura et al., 2001). Many molecules have been implicated in the formation of the triad junction and the physical coupling of the SR with the T-tubule membrane (e.g., amphiphysin 2, junctophilins, and mitsugumin 29) (Takeshima et al., 2000; Ito et al., 2001; Komazaki et al., 2001, 2002; Razzaq et al., 2001; Lee et al., 2002). The junctophilins in particular have been shown to be critical for the proper alignment of the T-tubule/SR membranes (Takeshima et al., 2000; Komazaki et al., 2002). This is thought to be mediated through interactions with proteins associated with the junctional SR (ryanodine receptor and triadin), suggesting that there is a protein network that organizes the localization of junctional proteins. Three independent cytoskeletal systems also exist at the T-tubule/SR membranes: the vinculin/talin/integrin complex, the dystrophin-glycoprotein complex and the spectrin-based membrane skeleton (Hoffman et al., 1987; Knudson et al., 1988; Kostin et al., 1998) and their absence leads to altered T-tubule morphology (Oguchi et al., 1982). In this study, the absence of Tm5NM1 led to a small increase in abnormally shaped T-tubules. This raises the possibility that the actin filament system has a role in maintenance of T-tubule membrane structure. The importance of cytoskeletal Tms in maintaining the structure and function of membrane structures has been recently shown in osteoclasts where knockdown of Tm4 resulted in changes in podosome shape and diminished bone resorptive capacity (McMichael and Lee, 2008).
Tm5NM1 Regulates T-Tubule Function
The T-tubule system is responsible for the propagation of the action potential from the sarcolemma of the muscle fiber to the contractile units. The T-tubules connect with the sarcoplasmic reticulum to form the triad and initiate the release of calcium for sarcomeric contraction. Altered excitation-contraction (E-C) coupling is a consistent feature of mice null for proteins located at the triad junction. This includes not only proteins involved directly in the process of E-C coupling (ryanodine receptor and dihydropyridine receptor) but also proteins involved in the formation of the triad junctional membrane complex (Takeshima et al., 1998, 2000; Razzaq et al., 2001). Tm5NM1 KO mice exhibit an alteration in contractile function consistent with changes to E-C coupling (increased twitch force, increase in the excitability as measured by repeated depolarization-induced force responses). It is unclear whether this is due to changes in the expression of components of the E-C coupling apparatus or some direct regulatory effect of the actin cytoskeleton on the process of E-C coupling.
We have shown that the increase in DICRs in Tm5NM1 KO muscle is not due to changes in SR function or contractile properties (Table 1) and the small increase in dysmorphic T-tubules in the KO (4%) does not account for this dysfunction. As endogenous Ca2+ in the SR (Table 1) and SR morphology were unaltered in the KO muscle it is unlikely that an increase in SR volume is responsible for the contractile phenotype. It would seem therefore that events preceding Ca2+ release from the SR are responsible for the altered T-system responses and presumably the increased twitch force. This could include 1) quicker repolarization of the T-tubule membranes in KO fibers, perhaps due to altered Na+/K+ pump function; 2) increased sensitivity of the DHPR (L-type Ca2+ channel) to change in voltage; or 3) greater ability of KO fibers to transfer the electrical signal to the ryanodine receptor triggering greater Ca2+ release.
We have shown that Tm5NM1 and Tm4 define independent -actin filament populations within the myofiber (Kee at al., 2004; Vlahovich et al., 2008). -Actin has also been shown to be part of the juxtamembrane complex, the costamere (Craig and Pardo, 1983; Rybakova et al., 2000), that is thought to provide a link between the dystroglycan complex and the sarcomeric Z-line (Rybakova et al., 2000). Surprisingly, ablation of this actin isoform in KO mouse muscle did not result in dystrophy, but a progressive breakdown of muscle (necrosis and regeneration) with features similar to centronuclear myopathy (Sonnemann et al., 2006). These mice are weak, have decreased twitch force and decreased action-potential evoked Ca2+ release. The authors attributed these contractile defects to increased mechanical compliance caused by altered connectivity between muscle fibers and/or myofibrils at the myotendinous junction. However, the data from the Tm5NM1 KO mouse in the present study suggests that the contractile defects in -actin KO muscle could be due to at least in part alterations to T-tubule/SR function.
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CONCLUSIONS |
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TOPABSTRACTINTRODUCTIONMATERIALS AND METHODSRESULTSDISCUSSIONCONCLUSIONSREFERENCES |
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). This study shows that skeletal muscle contains two cytoskeletal Tm isoforms, Tm5NM1 and Tm4, that compartmentalize to define independent actin filaments in association with internal membrane systems in skeletal muscle. Loss of Tm5NM1 in skeletal muscle resulted in contractile defects consistent with altered T-tubule function. In osteoclasts, knockdown of Tm4 results in changes in podosome shape and diminished bone resorptive capacity indicating the importance of this Tm in maintaining the structure and function of membrane structures (McMichael and Lee, 2008). Together, these studies suggest specific, independent roles for the Tm isoforms in association with different membrane structures.
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ACKNOWLEDGMENTS |
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Footnotes |
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These authors contributed equally to this work. 
Present address: The Burnham Institute for Medical Research, 10901 North Torrey Pines Rd., La Jolla, CA 92037.
Address correspondence to: Edna C. Hardeman (e.hardeman{at}unsw.edu.au).
Abbreviations used: DHPR, dihydropyridine receptor; DICR, depolarization-induced contractile response; EDL, extensor digitorum longus; EM, electron microscopy; FDB, flexor digitorum brevis; KO, knockout; MyHC, myosin heavy chain; SR, sarcoplasmic reticulum; Tm, tropomyosin; WT, wild type.
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