Altered nuclear dynamics in MDX myofibers

Microtubules provide means for nuclear movement but show altered organization in the muscular dystrophy mouse model (MDX) (dystrophin-null) muscle. Here, MDX myofibers show increased nuclear movement, altered transcriptional activity, and altered linkers of nucleoskeleton and cytoskeleton complex expression compared with healthy myofibers. Microtubule architecture was incorporated in finite element modeling of passive stretch, revealing a role of fiber malformation, commonly found in MDX muscle. The results suggest that alterations in microtubule architecture in MDX muscle affect nuclear movement, which is essential for muscle function.

Abstract

Duchenne muscular dystrophy (DMD) is a genetic disorder in which the absence of dystrophin leads to progressive muscle degeneration and weakness. Although the genetic basis is known, the pathophysiology of dystrophic skeletal muscle remains unclear. We examined nuclear movement in wild-type (WT) and muscular dystrophy mouse model for DMD (MDX) (dystrophin-null) mouse myofibers. We also examined expression of proteins in the linkers of nucleoskeleton and cytoskeleton (LINC) complex, as well as nuclear transcriptional activity via histone H3 acetylation and polyadenylate-binding nuclear protein-1. Because movement of nuclei is not only LINC dependent but also microtubule dependent, we analyzed microtubule density and organization in WT and MDX myofibers, including the application of a unique 3D tool to assess microtubule core structure. Nuclei in MDX myofibers were more mobile than in WT myofibers for both distance traveled and velocity. MDX muscle shows reduced expression and labeling intensity of nesprin-1, a LINC protein that attaches the nucleus to the microtubule and actin cytoskeleton. MDX nuclei also showed altered transcriptional activity. Previous studies established that microtubule structure at the cortex is disrupted in MDX myofibers; our analyses extend these findings by showing that microtubule structure in the core is also disrupted. In addition, we studied malformed MDX myofibers to better understand the role of altered myofiber morphology vs. microtubule architecture in the underlying susceptibility to injury seen in dystrophic muscles. We incorporated morphological and microtubule architectural concepts into a simplified finite element mathematical model of myofiber mechanics, which suggests a greater contribution of myofiber morphology than microtubule structure to muscle biomechanical performance.

NEW & NOTEWORTHY Microtubules provide the means for nuclear movement but show altered organization in the muscular dystrophy mouse model (MDX) (dystrophin-null) muscle. Here, MDX myofibers show increased nuclear movement, altered transcriptional activity, and altered linkers of nucleoskeleton and cytoskeleton complex expression compared with healthy myofibers. Microtubule architecture was incorporated in finite element modeling of passive stretch, revealing a role of fiber malformation, commonly found in MDX muscle. The results suggest that alterations in microtubule architecture in MDX muscle affect nuclear movement, which is essential for muscle function.

duchenne muscular dystrophy (DMD) is a progressive muscle-wasting disease caused by the absence of dystrophin, a 427-kDa protein located at the sarcolemmal membrane. DMD affects ~1 in 3,500 boys and results in progressive muscle degeneration and premature death. Although the exact role of dystrophin remains unclear, its absence in DMD results in a myriad of deleterious cellular consequences, which are associated with progressive muscle weakness and degeneration. The muscular dystrophy mouse model (MDX) mouse, an animal model of DMD, lacks dystrophin and is considered the most suitable murine model for DMD (69).

Although the genetic basis for DMD has been determined (30), the mechanisms responsible for the progressive muscle damage and weakness remain unclear (41, 66). The primary mechanism thought to promote the progressive nature of the disease is the mechanical weakness in the sarcolemma or the cytoskeletal-sarcolemmal interface attributable to the loss of dystrophin (33). However, this concept is further complicated by secondary factors, such as changes in dystrophin-associated proteins, calcium concentration, nitric oxide, inflammation (53), and reactive oxygen species (19, 31, 71). Such changes fit within what has been called a signaling hypothesis (51). Another hypothesis, known as the split-fiber hypothesis, has also been put forth (26). This is the result of a prevalence of malformed (branched) myofibers in dystrophic muscle, which lead to increased susceptibility to injury. In this study, we propose an additional factor that could contribute to the progressive muscle weakness, changes in nuclear movement (i.e., distance traveled and velocity of movement).

Proper nuclear movement is essential for muscle function and effective fiber repair following injury, especially in diseases like muscular dystrophy, in which mechanically induced muscle damage occurs frequently (3, 10, 17, 42). In mature myofibers, microtubules provide a means for intracellular transport, such as the movement of secretory vesicles, organelles, and nuclei (56). In fact, when specific microtubule-associated proteins are absent from muscle because of a conditional knockout, newborn mice die with severe dystrophy of muscles and myofibers showing a strong phenotype of nondispersed nuclei (67, 72). Because DMD and MDX muscle shows alterations in microtubule structure (5, 35, 55), extensive regeneration (6, 43), and nuclear mislocalization (17, 65), we investigated the movement of nuclei in MDX myofibers.

Because nuclear mobility is controlled, not only by microtubules, but also by the linkers of nucleoskeleton and cytoskeleton (LINC) complex that connects the cytoplasmic cytoskeleton with the nuclear interior (44), we also examined its components. Proteins of the LINC complex, such as nuclear envelope spectrin repeat protein (nesprin), are essential for the transmission of force to the nucleus. Myofibers are constantly subjected to mechanical forces; thus unstable regulation of the signaling pathway through the nucleus in response to mechanical stretch could be involved in the muscle pathogenesis.

In this study, we used a multidisciplinary approach to address the hypothesis that MDX myofibers would demonstrate altered nuclear movement, changes in gene expression of LINC complex proteins, and changes in global transcription compared with wild-type (WT) myofibers. We measured accumulated distance traveled and velocity of visible nuclei in WT and MDX myofibers with time-lapse imaging. We also examined nuclear histone H3 acetylation and polyadenylate-binding nuclear protein-1 (PABPN1), both indirect global measures of transcription, in WT and MDX myofibers. Disruption of the microtubule structure at the cortex of MDX myofibers has been previously reported; we extend these studies to include analysis of the core, as well as the cortex and core architecture of malformed-MDX myofibers, a common finding in dystrophic muscles, and incorporated the changes we found in cell shape and microtubule structure into a finite element model to determine potential mechanical weakness to passive stretch.

MATERIALS AND METHODS

Animals.

We used age-matched, sex-matched, control (WT) and MDX (lacking dystrophin) mice from the C57BL/10ScSnJ strain (The Jackson Laboratory, Bar Harbor, ME). A total of 10 mice were used (male, 3–4 mo old; n = 5 per group), and WT, MDX, and malformed-MDX myofibers (>50 per group) were imaged and used for analysis. All experimental procedures were approved by the University of Maryland Institutional Animal Care and Use Committee.

All experiments were performed in a blind fashion. Specifically, the procedures in image analysis (such as background removal, threshold for binarization, intensity measurement, etc.) were identical for each fiber/nucleus, precluding methodological bias by the experimenter. Each fiber/nucleus was processed in the same manner using identical algorithms (unbiased function of ImageJ to remove background, unbiased binarization threshold markers, unbiased tools to measure connectivity and number of branches, equal area in quantified region of interest, etc.) to obtain quantified measures from image analysis. Fibers were labeled in parallel. Laser power, pinhole diameter, brightness, and other confocal microscopy parameters were fixed and not altered between genotypes throughout experiments.

Myofiber isolation.

After mice were euthanized, the flexor digitorum brevis (FDB) muscles were harvested bilaterally. To obtain single myofibers by enzymatic dissociation, muscles were placed into DMEM with 1% BSA, 50 μg/ml gentamicin, and 4 mg/ml type I collagenase (C0130; Sigma, St. Louis, MO) for 1–3 h at 37°C as previously described (21, 40). Myofibers were plated on extracellular matrix (ECM; E1270; Sigma)-coated imaging dishes (P34G-1.0, 0–14-C; MatTek, Ashland, MA) before fixation.

Nuclear movement analysis.

Long-term cell incubation and time-lapse imaging were carried out using the automated Viva-View FL Incubator Microscope (Olympus, Center Valley, PA) as previously described (29) using the above-described method for myofiber isolation. Time-lapse imaging was conducted up to 10 h at 60-min acquisition intervals using a ×20 objective magnification. Time-lapse imaging was also conducted as described above with incubation of 1 h following the addition of 10 μM colchicine to some MDX myofibers. The collected time-lapse videos of WT and MDX (>90 nuclei examined for each type) were analyzed in ImageJ using Manual Tracking and Chemotaxis Tool plugins. Accumulated distance traveled and average velocity of all visible nuclei in each fiber were calculated. Data for histograms of accumulated distance and velocity of nuclei were generated in MATLAB.

Quantitative RT-PCR.

For real-time (RT)-PCR detection of LINC complex, microtubule motors and linker-related transcripts, three WT and three MDX quadriceps muscles were snap frozen and then homogenized in Trizol reagent (Invitrogen, Carlsbad, CA). Total RNA was extracted according to manufacturer’s instructions and reverse transcribed. Quantitative RT-PCR was performed as described previously, with an ABI 7300 Sequence Detection System (Applied Biosystems, Foster City, CA) using SYBR green (45). Relative expression was determined by comparison to housekeeping genes GAPDH, HPRT, and RPL13, using the geNorm software (v3.5, Ghent University Hospital, Ghent, Belgium). Transcripts for microtubule motors dynein cytoplasmic 1 heavy chain 1 (Dync1h1), kinesin-1 heavy chain (Kif5b), microtubule-associated protein 7 (ensconsin, Map7), outer nuclear envelope proteins, nesprin-1 (Syne1), nesprin-2 (Syne2), nesprin-3 (Syne3), inner nuclear envelope proteins, emerin (Emd), Sad1 and Unc84 domain-containing protein (SUN)-1 (Sun1), SUN-2 (Sun2), and lamin A/C (Lmna) were assessed. The primer sets used for PCR amplification are: Dync1h1 F: AGGAAACGCCTACAAGTGAGTCA, R: GAGGAACAACACCCCAACCTT; Kif5b F: AGCGGCCTCGCCAACT, R: ACTGGCTGGTTGTTCTGAACAA; Map7 F: CCAGAAATTCCTTTGAAACCAATT, R: GCACACCATCCACCTGAGGTA; Syne1 F: CCTTGTACCCATGTCAGAGAAAGACT, R: GGTATATCTGAGCATCGGATGGA; Syne2 F: GCAGATTCACGAACGACTGACA, R: GTTGCCCTCGTGCACCAT; Syne3 F: CGTGTAGACCGGTTACAAACTCAA, R: CTACCACGCTGTCAGAGAGTGACT; Emd F: TTACCCCCCAACTCGTCATC, R: CCACGGCGGCTGAATC; Sun1 F: TGCGGTTGTCCATGAAGATC, R: ATGTTACCAGTGGGTGATAGTGTCTTT; Sun2 F: ATGTGCACCCAGGCAACTG, R: TGCTCTAAGGTAACGGCTGTAGGT; Lmna F: TGTGGAAGGCGCAGAACAC, R: TGCGCATGGCCACTTCTT; Gapdh F: CGTGTTCCTACCCCCAATGT, R: TGTCATCATACTTGGCAGGTTTCT; Hprt F: AGCAGTACAGCCCCAAAATGG, R: AACAAAGTCTGGCCTGTATCCAA; Rpl13 F: CGAAACAAGTCCACGGAGTCA, R: GAGCTTGGAGCGGTACTCCTT.

Fluorescent antibody labeling and imaging.

Myofibers were fixed with 4% paraformaldehyde, permeabilized with 0.5% Triton X-100-PBS, blocked in 3% BSA-PBS, and then labeled overnight with an antibody to α-tubulin conjugated to Alexa Fluor 488 (anti-mouse; Invitrogen, 3 µg/ml). Isolated myofibers were washed and mounted in Vectashield (Vector Laboratories, Burlingame, CA). In some cases, 4’,6-diamidino-2-phenylindole (DAPI) was used to label nuclei. Myofibers were also incubated overnight with primary antibody against α-actinin (1:500; 87811, Sigma), β-spectrin (1:200; 58851; Santa Cruz Biotechnology, Santa Cruz, CA), acetyl-histone H3 (1:5,000; 9677; Cell Signaling Technology, Beverly, MA), nesprin (1:200; MAS-18077; Thermo Fisher Scientific, Rockford, IL), or PABPN1 (1:250; ab75855; Abcam, Cambridge, MA) and labeled with secondary antibody Alexa Flour 488 (α-actinin, acetyl-histone H3, nesprin, PABPN1) or Alexa Flour 594 (β-spectrin) (secondary antibodies, ThermoFisher Scientific, 1:100). Digital images of the microtubule and nesprin labeling were obtained with a Zeiss 510 confocal laser-scanning microscope (×63 objective) with pinhole set at 1.0 Airy unit (z-section thickness = 1 µm, 7.1680 pixels/µm for microtubules; z-section thickness = 1 µm, 14.34 pixels/µm for nesprin). Microtubules generally are up to 10 µm long although they can be up to 100 µm in axons (47, 59). Images of PABPN1 and acetyl-histone H3 in nuclei with DAPI were also obtained with a Zeiss 510 confocal laser-scanning microscope (×40 objective). WT, MDX, and malformed MDX fibers were imaged, and Z-stacked images were analyzed using ImageJ software (NIH, Bethesda, MD).

Examination of ultrastructure with electron microscopy.

Single myofibers were isolated from FDB muscles as above, plated on an ECM-coated coverslip, fixed in a 0.1 M phosphate buffer (pH 7.2) with 2% paraformaldehyde and 2.5% glutaraldehyde at 25°C for 1 h, and stored at 4°C before processing. Fixed cultures were washed and postfixed with 1% osmium tetroxide in 0.1 M phosphate buffer for 1 h and washed again.

Coverslips were dehydrated in a graded series of ethanols, chemically dried with hexamethyldisilazane (HMDS) (Electron Microscopy Sciences, Fort Washington, PA), mounted on scanning electron microscopy (SEM) pin mounts, and sputter-coated with platinum/palladium for SEM. SE images were acquired in a Quanta200 variable pressure SE.

Computing algorithms and image processing.

For nesprin, PABPN1, and acetyl-histone H3 labeling intensity and microtubule density quantifications, maximum-intensity flat z-plane projections from individual frames each spaced 1 μm apart, spanning either the nucleus, multiple nuclei, or 10 consecutive representative frames, respectively, in WT and MDX fibers, were used. Background was subtracted from the projection image and binarized with ImageJ unbiased binary function. Mean intensity of the binary image was measured in the entire nucleus in the binary projection image for nesprin, PABPN1, and acetyl-histone H3 labeling intensity. For microtubule density quantification, mean intensity of the binary image was measured in a uniform region of interest in the binary projection image (35).

Directionality of subsarcolemmal and core microtubule network in WT, MDX, and malformed MDX myofibers was analyzed using the Texture Detection Technique (TeDT) described by Liu and Ralston (39). Gray-level cooccurrence matrices were generated with sampling of 45 angle values from 0–180° using steps of 4° and pixel distances from 5–60 pixels using 11 steps with separation from 1–10 pixels. The pixel distance of 5 pixels is expected to be sufficient to analyze the microtubule lengths, given the typical size of microtubules (up to 10 µm) and the image scale (7.1680 pixels/µm). The joint probability of occurrence was calculated for each matrix. The texture correlation value for each direction was obtained by averaging the joint probability of occurrence for each angle. The relative texture correlation value was obtained by normalizing for each direction.

Whereas the subsarcolemma structure of the microtubule network in WT and MDX myofibers has been characterized and quantified, malformed MDX myofibers have not been studied. Furthermore, a comparison of the core and subsarcolemmal microtubule network has not been performed. The core microtubule network is a 3D network, whereas most quantification techniques such as the TeDT and density measurements are for 2D images. The Euler-Poincare characteristic, or Euler number, has been used to quantify structural changes in other tissues such as bone trabeculae and alveolar capillaries (23, 68). It determines the number of components in a complex structural network while including the connectivity of the network. Here we use the Euler number to quantify structural changes in the core domain microtubule network in myofibers. When viewing the fluorescently labeled core domain, the microtubule network in a myofiber exhibits connected longitudinal segments (connected microtubule networks), cavities (holes), and contiguous microtubule segments. The Euler characteristic for polyhedral surfaces is classically defined by the following:

$$\chi =\text{Vertices}-\text{Edges}+\text{Faces}$$ (1)

For the core domain microtubule network in a myofiber, the Euler number of the microtubule network was determined by the following:

$$\chi =\mathrm{C}\mathrm{o}\mathrm{n}\mathrm{n}\mathrm{e}\mathrm{c}\mathrm{t}\mathrm{e}\mathrm{d}\hspace{0.17em}\mathrm{M}\mathrm{i}\mathrm{c}\mathrm{r}\mathrm{o}\mathrm{t}\mathrm{u}\mathrm{b}\mathrm{u}\mathrm{l}\mathrm{e}\hspace{0.17em}\mathrm{N}\mathrm{e}\mathrm{t}\mathrm{w}\mathrm{o}\mathrm{r}\mathrm{k}\mathrm{s}-\text{Contiguous Microtubule Segments}+\text{Holes}$$ (2)

It is also known that the microtubules networks are connected. In Eq. 2, this simplifies the connected microtubule networks term to 1. There are no holes in the myofibers, as the microtubule network in the core domain is entirely filled with myofibrils. This eliminates the term holes to 0. Thus, Eq. 2 can be simplified to the following:

$$\chi =1-\mathrm{C}\mathrm{o}\mathrm{n}\mathrm{t}\mathrm{i}\mathrm{g}\mathrm{u}\mathrm{o}\mathrm{u}\mathrm{s}\hspace{0.17em}\mathrm{M}\mathrm{i}\mathrm{c}\mathrm{r}\mathrm{o}\mathrm{t}\mathrm{u}\mathrm{b}\mathrm{u}\mathrm{l}\mathrm{e}\hspace{0.17em}\mathrm{S}\mathrm{e}\mathrm{g}\mathrm{m}\mathrm{e}\mathrm{n}\mathrm{t}\mathrm{s}$$ (3)

χ is calculated for each 3D reconstruction voxel, through iteration of the image stack, by calculating the χ for each voxel. It is summed to provide the χ for the microtubule network.

To eliminate the edge problem of microtubule network, χ for faces, edges, and corners are deducted from χ, and the number of contiguous microtubule segments is calculated by Eq. 2.

For quantification of the contiguous microtubule segments, 10 consecutive slices spaced 1 μm apart of each myofiber core were selected. After background was subtracted, binary images were created using ImageJ through implementation of Huang’s fuzzy thresholding method. Purify function in the BoneJ plugin of ImageJ was performed such that Eq. 3 was satisfied. The Euler Number was then calculated using the connectivity function. A 3D skeleton was also developed using the binary images. Branching of the microtubule network was quantified using the 3D skeleton.

Finite element analysis.

Equivalent total strain [i.e., sum of equivalent elastic strain (linear component of strain) and equivalent plastic strain (nonlinear component of strain)] in normal and malformed MDX myofibers was calculated in ANSYS. A myofiber section was modeled with length of 500 µm, width of 50 µm, and thickness of 1 µm, with rounded edges. The passive mechanical properties of the myofibrils, titin, and other cytoskeletal components were approximated in the myofiber section in bulk as a hyperelastic material using a two-parameter Mooney-Rivlin strain energy function with C₀₁ = 3,000 Pa, C₁₀ = 1,500 Pa, and D₁ = 0.02, which were adapted from previous literature on skeletal muscle modeling (62). Microtubules were modeled as an isotropic linear elastic material with thickness of 25 nm, width of 4 µm, and varying lengths ranging from 7–500 µm and Young’s Modulus of 500 MPa and Poisson’s ratio of 0.3 (46). Microtubules were overlaid on top of the myofiber section (Fig. 5A). A force of 0.1 nN in the x direction and 1 pN in the y direction was applied to one end of the myofiber and fixed at the other end to simulate passive stretch on a resting myofiber. This force was applied to a 1-µm section of the myofiber, which is typically ~40–50 µm in thickness. This simulated passive tensile and shear force induces positive strain in the myofibers, which was determined using the model. The contact between the microtubules and the myofibers section was modeled as bonded. A 3D analysis was performed on models of a normal morphology myofiber without microtubules, a normal morphology myofiber with longitudinal microtubules, a normal morphology myofiber with denser longitudinal microtubules, a normal morphology myofiber with longitudinal and transverse microtubules, a malformed morphology myofiber without microtubules, a malformed morphology myofiber with denser longitudinal microtubules, and a malformed morphology myofiber with transverse and denser longitudinal microtubules. The organization of the microtubules used in the model was developed independently of the microtubule image analysis, using simplified and generalized approximations of microtubule organizations developed from this study and previous work (5, 35, 53). The maximum equivalent total strain was also averaged at three nodes near the bifurcation for the malformed myofibers to eliminate singularities.

Fig. 5.

Nuclei are mislocalized in malformed MDX myofibers. MDX muscle contains a high percentage of malformed myofibers, which are typically absent in healthy muscle. A: scanning electron microscopy image shows a typical MDX myofiber (top) and a malformed MDX myofiber (bottom). A variety of malformations are present in MDX muscles, but the majority are bifurcated at 1 end as shown here. Magnification ×600. Scale bar = 100 µm. B: montage of images with examples of mislocalized nuclei in malformed MDX myofibers. Nuclei are stained with either DAPI (blue nuclei) or propridium iodide (red nuclei). A healthy (WT) myofiber contains hundreds of nuclei, which are positioned peripherally to maximize the distance between adjacent nuclei. Note here that, in myofibers from malformed MDX muscle, the nuclei are not distributed equally. Scale bar = 20 µm.

Statistical analysis.

Statistical analyses were performed using JMP Version 12 (Cary, NC) and SigmaStat 3.5 (San Rafael, CA). Data are presented as means ± SD unless otherwise noted. Statistical significance was assessed using a one-way ANOVA with P < 0.05. Assumptions of normality and equal variance were confirmed. If the assumptions of normality or equal variance were not met, an ANOVA on Ranks was performed. A Tukey’s post hoc analysis was performed to determine where significant differences had occurred.

RESULTS

Nuclear movement.

In general, the nuclei in MDX myofibers were more mobile than in WT myofibers (Fig. 1A). Both the distance traveled and velocity (Fig. 1B) were greater in nuclei of MDX (19.49 ± 0.051 μm; 0.035 ± 9.50x10⁻⁵ μm/min, respectively) compared with WT myofibers (14.18 ± 0.048 μm; 0.027 ± 9.82x10⁻⁵ μm/min, respectively), as indicated by the clear rightward shift in the distributions of each parameter. Qualitatively, nuclei in MDX myofibers exhibited a loss in transverse motion (see arrows in 1A). The addition of colchicine, a microtubule depolymerizing agent, significantly reduces nuclear movement in healthy muscle (15, 70). Here, using colchicine in MDX, we found a significant reduction in nuclear distance traveled and velocity (11.34 ± 0.064 μm; 0.021 ± 0.001 μm/min, respectively) compared with WT and MDX myofibers without colchicine treatment.

Fig. 1.

Nuclear movement. A: representative images from paused time-lapse experiments in wild-type (WT) and muscular dystrophy mouse model (MDX) myofibers. Traces of nuclei movement from >90 nuclei in 7 myofibers per genotype and treatments over 10 h are superimposed onto the images. Scale bar = 25 µm. B: frequency distributions of accumulated distance traveled and velocities of nuclei in WT and MDX myofibers. Inset: means ± SE in a bar graph form for WT, MDX, and MDX treated with 10 µM colchicine (col). Note the increased distance and velocity observed in MDX myofibers (arrows) compared with WT myofibers. Interestingly, in the MDX myofibers, there is a corresponding decrease in the transverse movement of nuclei. Colchicine, a microtubule-depolymerizing agent, has previously been shown to reduce nuclear movement in healthy skeletal muscle. Here, treatment of MDX myofibers with colchicine resulted in reduced nuclear distance traveled and velocity. *P < 0.05 compared with WT; #P < 0.05 compared with MDX.

LINC complex.

Nuclear movement and stability can be affected by disruptions in the protein network that tethers the nucleus to the cytoskeleton (15, 70). We assessed transcripts for LINC proteins (Fig. 2A) such as outer nuclear envelope proteins nesprin-1 and nesprin-2. MDX had significantly lower expression of transcripts for nesprin-1 and nesprin-2 (Fig. 2B). The finding of reduced nesprin-1 intensity in immunolabeled images with confocal imaging further suggests reduced nesprin protein abundance and disruption of the LINC complex in MDX muscle (Fig. 2C). We also found significant reduction in expression of the genes encoding the inner nuclear envelope proteins SUN-1/2 in MDX muscle compared with WT muscle, which is required for proper nuclear anchorage in skeletal muscle through nesprin-1 localization and maintenance of the perinuclear space (37) (see Table 1).

Fig. 2.

Expression of nuclear envelope proteins. A: bottom schematic shows a myofiber with the dystrophin glycoprotein complex (DGC) at the sarcolemma, connecting the extracellular matrix to the underlying myofiber cytoskeleton (microtubules not shown). Dystrophin, thought to be important in linking the intracellular cytoskeleton to the extracellular matrix, is indicated in yellow. Inset: proteins that comprise the linker of nucleoskeleton and cytoskeleton (LINC) complex. Although lamin serves to stabilize the nuclear envelope, nesprins serve as a link between the nucleus and the myofiber cytoskeleton. At the NH₂ termini, nesprins-1 and -2 (yellow) have a pair of calponin homology domains that mediate binding to actin. As indicated by the schematic, nesprins interact with microtubule motors (dynein and kinesin), as well as microtubules and the actin cytoskeleton. Nesprin-3 contains a plectin-binding site on its NH₂ terminus, which allows binding to plectin, a cytoskeletal link in the cytoplasm that forms connections to actin, microtubule-associated proteins (MAPs), and intermediate filaments. INM, inner nuclear membrane; ONM, outer nuclear membrane. B: relative expression of transcripts for nuclear envelope proteins nesprin-1 and -2. Values are normalized to housekeeping genes and displayed with WT expression levels normalized to a value of 1. Data are shown as means ± SD for quadriceps muscle from 3 animals per genotype, *P < 0.05. C: representative immunolabeling of nuclei with nesprin-1 (green) and DAPI staining (blue) from WT and MDX myofibers. Confocal image shows loss in signal intensity. Bar graph shows means ± SE of nesprin-1 labeling intensity from nuclei in 4–7 myofibers per genotype, which revealed that MDX nuclei have significantly lower nesprin-1 labeling intensity compared with WT nuclei; *P < 0.05. Scale bar = 5 µm.

Table 1.

Expression of nuclear envelope proteins

Genes	WT	MDX	P Value
Dync1h1	1.00 ± 0.23	1.16 ± 0.22	0.530
Kif5b	1.00 ± 0.11	0.55 ± 0.12	0.018
Map7	1.00 ± 0.08	0.34 ± 0.04	<0.001
Syne1	1.00 ± 0.12	0.40 ± 0.02	0.002
Syne2	1.00 ± 0.11	0.38 ± 0.05	0.002
Syne3	1.00 ± 0.3	0.57 ± 0.19	0.164
Sun1	1.00 ± 0.08	0.69 ± 0.10	0.027
Sun2	1.00 ± 0.14	0.60 ± 0.01	0.017
Emd	1.00 ± 0.16	0.56 ± 0.04	0.022
Lmna	1.00 ± 0.11	1.63 ± 0.26	0.036

Values are shown as mean relative expression ± SD of transcripts in quadriceps for microtubule motors, associated proteins, and nuclear envelope proteins in 3 wild-type (WT) and 3 muscular dystrophy mouse model (MDX) animals. Note the reduced expression of transcripts for many of the nuclear envelope proteins and increased expression of transcript for lamin A/C in intact MDX muscles compared with WT. Values are normalized to housekeeping genes and displayed with WT expression levels normalized to a value of 1.

Transcriptional activity of WT and MDX nuclei.

Abnormal mobility, shape, or position of nuclei could indicate compromised global gene transcription (36), and loss of nesprin has been shown to affect nuclear chromatin (13). Because acetylation of histone H3 and PABPN1 positively correlate with active global gene transcription (2, 27), transcriptional activity of myofibers was assessed by analysis of PABPN1 and acetyl-histone H3 labeling intensity (20). We used DAPI as a marker for nuclear DNA and compared nuclear acetylation and polyadenylation between WT and MDX myofibers. We found homogeneous immunostaining of acetyl-histone H3 and PABPN1 in nuclei of WT myofibers, indicating coordinated transcriptional activity (Fig. 3). By contrast, the nuclei in MDX myofibers showed heterogeneous histone H3 acetylation and PABPN1 and even some nuclei without staining (arrows, Fig. 3). MDX nuclei, on average, had significantly reduced intensity for both PABPN1 and acetyl-histone H3 (reduction of 18.8% and 18.5%, respectively) compared with WT nuclei. These experiments are a general indicator of overall transcription and not of individual genes but nonetheless support the notion that changes in nuclear function could be related to altered nuclear mobility.

Fig. 3.

Assay of transcriptional activity in WT and MDX nuclei. Representative images of WT and MDX myofibers immunolabeled for polyadenylate-binding nuclear protein-1 (PABPN1) (green), acetyl-histone H3 (AH3) (red), and stained with 4’,6-diamidino-2-phenylindole (DAPI) (blue). Nuclei in WT myofibers have near homogeneous PABPN1 and AH3 immunolabeling, indicating similar levels of transcriptional activity. By contrast, nuclei in MDX myofibers clearly have varying PABPN1 and AH3 labeling, indicating heterogeneous transcriptional activity (arrows). Bar graph shows the mean ± SE labeling intensity of PABPN1 and AH3 in WT and MDX nuclei (n = 3–6 myofibers per genotype); *P < 0.05. Scale bar = 20 µm.

Microtubule organization of MDX myofibers.

Microtubules are considered the “railroad tracks” for cellular organelle transport, including the nucleus. In muscle fibers, microtubules have been described as having two domains (39) including the following: an organized superficial domain of cortical microtubules at the periphery of the myofiber, encircling the fiber core (cortex, beneath the sarcolemma but above the underlying myofibrils), and a deeper, less dense microtubule domain, where microtubules run parallel to the longitudinal axis of the myofiber and are intermingled between the myofibrils (core, Fig. 4A). Interestingly, microtubules bind to dystrophin, and the normal latticed network of microtubules seen in the cortex of WT myofibers is disrupted in the cortex of MDX myofibers (Fig. 4B) (5, 55). This disorganized microtubule lattice appears to further promote susceptibility to contraction-induced injury in dystrophin-deficient skeletal muscle (5, 35). Here, we used a new analytical tool to examine, for the first time, the 3D organization of microtubules in the core of WT and MDX myofibers.

Fig. 4.

Microtubule network in WT and MDX myofibers. A: cartoon demonstrating the 2 domains of the microtubule network in a myofiber: a dense, latticed network in the subsarcolemmal domain (cortex) of the myofibers, and a loosely packed microtubule network in the core that is mostly comprised of longitudinal microtubule segments with intermittent transverse microtubule segments. B: microtubules in flexor digitorum brevis fibers were immunolabeled with Alexa-488 (green)-conjugated primary antibody against α-tubulin. Top: latticed microtubule network in the cortex domain of WT myofibers, which is disorganized in MDX myofibers. Scale bar = 5 µm. Bottom: core domain microtubule network. This particular myofiber nicely demonstrates the loss of transverse microtubule segments in the MDX myofibers. Scale bar = 10 µm. The amount of contiguous microtubule segments (C) and the number of branches (D) were analyzed in the core of WT, MDX, and malformed MDX myofibers. Measurement of the Euler number in 3D image volumes revealed differences in MDX myofibers in microtubule organization, such as a significant reduction in contiguous microtubule segments in the core domains of MDX myofibers compared with WT (C). Decreased branching of the microtubule network was also observed in MDX myofibers compared with WT (D) (not shown: contiguous microtubule segments and number of branches in malformed MDX myofibers, which were also significantly decreased compared with WT, were not different than in MDX myofibers with normal morphology). Data are shown as means ± SD from 7 myofibers per genotype; *P < 0.05, 1-way ANOVA.

A subtle decrease in transverse microtubule segments in MDX myofibers (i.e., connectivity) in both the cortical and core domains is observed with immunofluorescent labeling (Fig. 4B). When quantified, measurement of the Euler number in 3D image volumes revealed differences in MDX myofibers in microtubule organization, such as a significant reduction in contiguous microtubule segments in the core domains of MDX myofibers compared with WT (Fig. 4C). Decreased branching of the microtubule network was also observed in MDX myofibers compared with WT (Fig. 4D). On the basis of quantitative immunofluorescence analysis, we also found increased density in the microtubule network and reduced transverse directionality of cortical microtubules in MDX myofibers compared with WT myofibers, consistent with previous reports (5, 35, 39, 55), but no differences in the core microtubule transverse directionality between WT and MDX myofibers (not shown).

Myofiber morphology.

Healthy, mature skeletal myofibers do not branch (Fig. 5A, top myofiber), but there is an age-dependent increase in the number of malformed myofibers (40, 52) that could account for the age-dependent increase in muscle damage and weakness seen in the murine MDX model of dystrophic muscle (12). The number of branched myofibers can be as high as 90% in MDX muscle (12, 52), and malformed myofibers have also been reported in boys with DMD through serial sectioning of muscle biopsies (6, 26). Malformed myofibers are typically simple with only one branch (Fig. 5A, bottom myofiber) but can also be complex, and many different configurations have been described (28, 40). Figure 5B shows an assortment of malformed MDX myofibers, which typically show mislocalized nuclei, as seen here. Indeed, it is in malformed MDX fibers in which malpositioned nuclei are most prevalent (16, 40). In terms of the core microtubule architecture, contiguous microtubule segments and branching were significantly reduced in malformed MDX myofibers compared with WT myofibers although they were not significantly different compared with MDX myofibers (not shown). Similarly, we found increased microtubule density and reduced cortical microtubule transverse directionality in malformed MDX myofibers compared with WT myofibers although not significantly different compared with MDX myofibers. We found no differences in core microtubule transverse directionality between malformed MDX, MDX, and WT myofibers. Accordingly, we wanted to determine the relative contributions of altered microtubule architecture and altered myofiber shape to the defective biomechanical properties that result in susceptibility to damage.

Biomechanical modeling (myofiber shape vs. microtubule cytoskeleton).

Experimentally measuring the strain distributions in malformed myofibers under load is fraught with challenges (such as effects of gripping ends of myofibers on strain distributions at the bifurcation, geometrical heterogeneity of malformations, and isolating the effects of differences in microtubule/cytoskeletal organization on strain distributions). To overcome these challenges and to examine the relative contributions of fiber morphology and microtubule architecture to muscle function, finite element analysis was performed on models of normal and malformed myofibers (Fig. 6, A and B), with varying amounts and different orientations of networked microtubules. In this model, we simulated passive tensile and shear forces to induce positive strain in the myofibers. In the absence of microtubules (i.e., only considering myofiber geometry and bulk fiber material properties), we found increased equivalent total strain (~80% higher) and nonuniform strain distributions in the malformed morphological myofiber (Fig. 6D, solid bar) compared with the normal morphological myofiber (Fig. 6C, solid bar). This increased strain and nonuniform strain distribution could indicate increased susceptibility to damage in pathological myofibers, as reported by others, on the basis of geometry alone, especially in the region of the bifurcation. Inclusion of the microtubule network models in both myofiber geometries reduced maximum strain, indicating that microtubules contribute significantly to myofiber stiffness. The removal of transverse microtubule segments and the addition of increased microtubule density (Fig. 6C, open bar), despite having normal morphology, resulted in a <13% increase in maximum strain compared with the model of normal myofibers with a typical latticed microtubule network (Fig. 6C, shaded bar). However, the malformed myofiber model was more sensitive to changes in microtubule organization. Our simulation of the observed pathology in branched fibers (see Fig. 5) (i.e., with increased microtubule density and reduced transverse microtubule segments, Fig. 6D, open bar) were found to have increased maximum strain (71% higher) and nonuniform strain distribution at the bifurcation compared with normal myofiber shape and microtubule organization (Fig. 6C, shaded bar). Although the addition of transverse microtubule segments partially rescued (i.e., reduced) maximum strain in the malformed myofibers (Fig. 6D, shaded bar), our model indicated that the malformed myofibers remained mechanically compromised primarily attributable to their geometric differences.

Fig. 6.

Mathematical modeling of myofiber mechanical properties. A: examples of overlay of microtubule network on myofiber sections of 1-µm thickness. Modeling with and without overlaid longitudinal or latticed microtubule network with morphologies of normal and malformed MDX myofibers was performed. Passive stretch of 0.1 nN in the longitudinal direction (x-axis) and 1 pN in the transverse direction (y-axis) was simulated on the section of the resting myofibers, with network of myofibrils as the underlying hyperelastic material and the overlaid microtubule structure as a linear-elastic material. The underlying hyperelastic material (myofiber section) has length of 500 µm, width of 50 µm, and thickness of 1 µm. The overlaid microtubule network (linear-elastic material) had varying lengths ranging from 7-500 µm, thickness of 25 nm, and width of 4 µm. B: color map shows equivalent total strains calculated for myofibers with normal myofibers (top, latticed network, simulating WT) and malformed MDX myofibers (bottom, absent transverse segments). C: for normal morphology, analysis included modeling without any microtubules (-MT), with dense longitudinal (+ MT, Long), and with longitudinal and transverse microtubules (MT, Lattice). D: for malformed morphology, equivalent total strains were also calculated for the same conditions. The presence of microtubule transverse components decreases maximum strain in WT fibers. Nonuniform strain distributions can be observed in malformed fibers, whereas uniform strain distribution is observed in fibers with normal morphology, which could indicate increased susceptibility to stretch-induced damage in malformed myofibers. Bar graphs represent equivalent total strains of the myofiber to equal forces without microtubules (solid bar) and with longitudinal microtubules (open bar) and with latticed microtubules (shaded bar). Note that restoring the transverse microtubules rescues the strain response in this model.

DISCUSSION

Nuclei in mature skeletal myofibers are relatively anchored, but increased nuclear movement is associated with reorganization of cytoskeleton (10, 17). Because MDX muscle has altered microtubule structure (5, 39, 55), we compared nuclear movement in WT and MDX muscles. We found increased nuclear movement (velocity and distance traveled) in MDX myofibers compared with WT myofibers. We further found decreased gene expression and labeling intensity of nesprin-1, a key LINC complex protein in nuclear mechanotransduction, along with altered global transcriptional activity in the nucleus of MDX myofibers. As nuclear mobility is microtubule dependent, we analyzed microtubule structure at the core and cortex, and altered microtubule structure is found at both the core and the cortex for MDX myofibers. In addition, finite element modeling of passive stretch of myofibers using morphological and microtubule architecture data (including malformed MDX myofibers) revealed a greater role of myofiber morphology, such that malformed MDX myofibers exhibited mechanical weakness with increased strain and nonuniform strain distribution at the bifurcation.

Much of the previous work on mechanical influences on muscle gene expression (i.e., mechanotransduction) has focused on structures like the dystrophin glycoprotein complex (see dystrophin glycoprotein complex in Fig. 2) and stretch-sensitive channels at the cell surface, which transduce mechanical forces into biochemical signaling cascades that affect nuclear gene expression (14). In addition to this conventional, indirect signaling, recent data indicate that the nucleus can respond directly, and even more quickly, to mechanical stress (44). The nuclear envelope shows plasticity in regard to its environment, such that stiffness can change with changes in the extracellular environment (64). In fact, when certain proteins in the LINC complex are absent, nuclear anchorage and nuclear-cytoskeletal strain transmission are also decreased, resulting in altered expression of known mechanosensitive genes (4, 13). The nucleus and the LINC complex contribute to the organization of the microtubule skeleton (63), and recent work suggests that the nucleus through the LINC complex is critical for proper assembly and stability of the sarcomere (3). It is unlikely that microtubules alone account for changes in nuclear movement, as defects in nuclear positioning occur even in the absence of effects on microtubule organization (42). Such reports add further complications to our findings on increased nuclear movement in MDX myofibers.

Compared with WT myofibers, we found increased nuclear mobility in MDX myofibers, both in terms of distance traveled and velocity of movement within the myofiber. We also found changes in expression of the LINC complex, including nesprins. Nesprins are a key component of the LINC complex, which binds to the cytoskeleton, microtubule-based motors, and plectin (22). Nesprin-1 allows for nucleus stiffening and subsequent nuclear mechanical response, when subjected to force, and is crucial for proper positioning and anchorage of nuclei in skeletal muscle (13, 22, 72). We show for the first time that expression levels of transcripts for nesprin-1/2 and SUN 1/2 were reduced and also that labeling intensity of nesprin-1 in MDX myofibers was reduced compared with WT myofibers. Although quantitative global protein content for nesprin-1 in muscle was not determined in this study, the immunofluorescent analysis of nesprin-1 showed a reduction of labeling specific to the nucleus. SUN-1/2 is required for proper nuclear anchorage in skeletal muscle through nesprin-1 localization and maintenance of the perinuclear space (37). Interestingly, nuclear clustering at the synaptic region is linked to muscle-specific kinase (MuSK) patterning (10), and we have previously shown that MuSK is reduced in MDX muscles (54). Defects in nesprins, emerin, and lamin A/C have also been shown by previous studies to contribute to the pathology in Emery-Dreifuss muscular dystrophy (73). The changes we report in gene expression of nuclear e nvelope proteins suggest that MDX muscles could potentially have impaired nuclear mechanotransduction. Such changes can further alter gene expression, transcriptional activity, and downstream effects of protein expression. One limitation in using indirect measures of global markers of transcription is their lack of transcriptional specificity to the key proteins involved in nuclear mobility. Nevertheless, these experiments support the notion that MDX myofibers have changes in nuclear function that could be related to altered nuclear mobility (13, 20, 36). We did not examine changes in nuclear shape and volume, but such alterations could also change nuclear protein concentration, and gene regulation and transcription (36).

The branching seen in malformed MDX myofibers presumably occurs in myofibers undergoing regeneration (52), a process in which nuclear movement may be essential for subsequent fiber repair and muscle function. Generation of nascent skeletal muscle is not restricted to development but also occurs in normal adult muscle after injury (such as eccentric contraction-induced injury, cardiotoxin-induced injury, etc.) or in muscle disease. For example, nuclei are relatively more mobile in developing muscle (myotubes ~0.20–0.88 µm/min) compared with nuclei in mature muscle (0.03–0.14 µm/min) (10). The gross internal structure of malformed MDX myofibers at first appears relatively normal under light microscopy (40). However, closer inspection has shown some irregularities in the normal register of myofibrils (9, 18, 21, 58). These structural irregularities extend deep into the MDX myofibers (18), but the prevalence and consequence of these structural abnormalities are unclear. One possible explanation for this altered sarcomere structure is altered nuclear dynamics, particularly in muscle regenerating after injury, as recent work shows that the nuclei are, not only important as mechanosensors, but also crucial for sarcomere assembly and stability (3).

The neuromuscular junction (NMJ) has clustered nuclei with NMJ-specific mRNAs and low mobility (15). To support the large cytoplasm in the rest of skeletal myofiber, extrasynaptic nuclei are positioned to maximize the distance between adjacent nuclei (8). This spatial distribution of nuclei minimizes transport distances, such that each nucleus regulates the gene products in a fixed volume of a muscle fiber, a concept known as the myonuclear domain (25, 38, 48). The number of peripherally located nuclei is lower in MDX compared with WT myofibers, resulting in a much larger (~23%) myonuclear domain (65). Despite the altered positioning of nuclei in MDX myofibers (especially with malformations), little is known regarding the role of nuclear mobility in diseased muscle (17). Disruption of the microtubule network will certainly impact spatial distribution of nuclei (17), but it can also affect nuclear mobility, which can greatly influence the maintenance of myonuclear domains as well as the transcriptional profile of individual nuclei, resulting in dramatic effects on muscle structure and function (10, 17, 42).

Microtubules are highly dynamic polymers of tubulin, constantly undergoing assembly and disassembly. Regulation of the microtubule network is extremely important for cell development, and, in addition to acting as compression-bearing struts that contribute to cell shape (1, 32), microtubules provide tracks for directing organelle transport (11, 24, 61, 70). Microtubules are capable of growing and shrinking to generate force, and the associated motor proteins carry organelles and other cellular components along the microtubule network (15, 24). This combination of roles makes microtubules important for organizing and moving intracellular constituents and nuclei. Movements and rotations of nuclei in healthy skeletal muscle are halted when microtubules are depolymerized with colchicine (15) or nocodazole (70), confirming a role for microtubules in nuclear movement. Indeed, in our hands, the addition of colchicine to MDX myofibers resulted in significantly reduced nuclear distance traveled and velocity. Microtubules are dramatically reorganized during normal muscle development (10, 11, 17). They are also noticeably reorganized under pathological conditions, such as denervation (57), unloading (60), and in muscle diseases (35, 49, 55).

The organization of microtubules within the cell is cell-type specific. In healthy mature muscle, they form a highly structured grid pattern, or lattice-type network, arranged primarily along the long axis of the myofibers. The normal structure of microtubules is disrupted in MDX myofibers, which lack dystrophin (5, 49, 55). We (40) and others (7, 52) have shown that malformed MDX myofibers are a unique subset of MDX myofibers. In addition to the age-dependent increase in the number of malformed myofibers, we have previously reported that malformed MDX myofibers are functionally different from WT and normal morphology MDX myofibers in terms of excitation-contraction coupling, action potential properties, and sarcolemmal mechanics (21, 28, 40). Here, we studied the microtubule network of malformed MDX myofibers and found no significant changes in microtubule density or cortical and core domain microtubule transverse directionality compared with normal morphology MDX myofibers. In the core of myofibers, the number of contiguous microtubule segments was reduced in malformed MDX myofibers compared with WT myofibers but not significantly reduced compared with MDX myofibers with normal morphology.

Although direct measurements of contractile and passive mechanical properties of myofibers can be made, there are limitations to experimentally measuring stresses and strains in abnormally shaped myofibers, attributable to the effects of gripping ends of myofibers on strain distributions at bifurcations, geometrical heterogeneity of malformed myofibers, changes in microtubule organization, etc. Thus we developed a detailed finite element myofiber model, in which passive stretch was simulated on a resting myofiber. The components of the model were adjusted to conform to the differences in microtubule densities and transverse directionality that we observed in normal, MDX, and malformed MDX fibers. The model reveals that the variation in architecture and geometry of malformed myofibers is predicted to cause large strains and substantial nonuniform strain distributions within a myofiber. This suggests that, even with the same application of force, malformed MDX myofibers are more likely to be susceptible to damage than normally shaped myofibers. The increase in microtubule density seen in MDX and malformed MDX myofibers appears to be a compensatory but insufficient mechanism to reduce increased susceptibility to damage of the malformed MDX myofibers, which may play a role in the progressive nature of DMD.

Drugs that affect microtubule stability are presently under investigation as potential therapies for disease and injury (5, 35, 46, 55); however, the exact role of the microtubule network in muscle is still being elucidated. In addition to its role as a mechanosensor, the microtubule network affects the position of organelles, such as nucleus, Golgi complex, and mitochondria, and rate and direction of their transport. Interestingly, altered distribution of mitochondria in MDX myofibers has been reported (50), with a paucity of subsarcolemmal mitochondria. The role of microtubule-associated proteins, LINC proteins, and posttranslational modifications (34) in vesicle trafficking is unclear. We show here that nuclear movement is altered in MDX muscles, with changes in gene expression of LINC complex proteins and associated microtubule proteins and motors. The mechanistic underpinnings for nuclear mechanotransduction must be further elucidated with disease and injury and can be targeted for downstream effects in gene expression and cytoskeletal organization.

GRANTS

This work was supported by grants from the National Institutes of Health, including training grant T32 AR-007592 (S. Iyer) and research grants R37-AR055099 (M. Schneider), R01-AR059179, and R21-AR067872-01 (R. Lovering).

DISCLOSURES

No conflicts of interest, financial or otherwise, are declared by the authors.

AUTHOR CONTRIBUTIONS

S.R.I., S.B.S., A.P.V., R.M.L. conceived and designed the study; S.R.I., E.O.H.-O., S.B.S., S.S.B., and J.P.S. performed experiments; S.R.I., S.B.S., M.F.S., E.O.H., J.P.S., and S.S.B. analyzed data; S.R.I., S.B.S., M.F.S., E.O.H.-O., J.P.S., S.S.B., and R.M.L. interpreted results of experiments; S.R.I., J.P.S., and R.M.L. prepared figures; S.R.I., S.B.S., and R.M.L. drafted manuscript; S.R.I., S.B.S., A.P.V., M.F.S., E.O.H.-O., J.P.S., S.S.B., and R.M.L. edited and revised manuscript; S.R.I., S.B.S., A.P.V., M.F.S., E.O.H.-O., J.P.S., S.S.B., and R.M.L. approved final version of manuscript.