# Myofibrillar malformations that arise in mdx muscle fibers are driven by detyrosinated microtubules

Anicca Harriot, Tessa Altair-Morris, Camilo Venegas, Jacob Kallenbach, Kaylie Pinto, Humberto Joca, Marie-Jose Moutin, Guoli Shi, Jeanine Ursitti, Anna Grosberg, et al.

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Anicca Harriot, Tessa Altair-Morris, Camilo Venegas, Jacob Kallenbach, Kaylie Pinto, et al. Myofibrillar malformations that arise in mdx muscle fibers are driven by detyrosinated microtubules. 2023. [hal-04284322](https://hal.science/hal-04284322v1)

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1 **Myofibrillar malformations that arise in mdx muscle fibers are driven by detyrosinated**
2 **microtubules**

3 *Anicca Harriot*¹, *Tessa Altair-Morris*², *Camilo Venegas*³, *Jacob Kallenbach*⁴, *Kaylie Pinto*³,

4 *Humberto C. Joca*¹, *Marie-Jose Moutin*⁵,⁶, *Guoli Shi*⁴, *Jeanine Ursitti*⁴, *Anna Grosberg*²,⁷,

5 *Christopher W. Ward*¹,⁴*

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7 ¹Department of Biochemistry & Molecular Biology, University of Maryland School of Medicine,
Baltimore, MD 21201

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2 Edwards Lifesciences Foundation Cardiovascular Innovation & Research Center and Center for
Complex Biological Systems and The NSF-Simons Center for Multiscale Cell Fate Research,
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University of California, Irvine, Irvine CA 92697

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13 ³Department of Molecular Medicine, University of Maryland School of Medicine, Baltimore, MD
21201

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⁴Department of Orthopedics, University of Maryland School of Medicine, Baltimore, MD 21201

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⁵Grenoble Institut des Neurosciences (GIN), Université Grenoble Alpes, F-38000 Grenoble,
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France

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⁶Inserm, U1216, F-38000 Grenoble, France

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⁷Department of Biomedical Engineering and Department of Chemical & Biomolecular
Engineering and Sue and Bill Gross Stem Cell Research Center

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*Corresponding author:

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C.W. Ward Department of Orthopedics, University of Maryland School of Medicine, Baltimore,
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MD 21201

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Email: [ward@som.umaryland.edu](mailto:ward@som.umaryland.edu)

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**Abstract**

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In Duchenne muscular dystrophy (DMD), alterations in the myofibrillar structure of skeletal
muscle fibers that impair contractile function and increase injury susceptibility arise as a
consequence of dystrophic pathology. In murine DMD (mdx), myofibrillar alterations are abundant
in advanced pathology (>4 months), an age where we formerly established the densification of
microtubules (MTs) post-translationally modified by detyrosination (deTyr-MTs) as a negative
disease modifier. Given the essential role of MTs in myofibrillar growth, maintenance, and repair,
we examined the increased abundance of deTyr-MTs as a potential mechanism for these
myofibrillar alterations. Here we find increased levels of deTyr-MTs as an early event in dystrophic
pathology (4 weeks) with no evidence of myofibrillar alterations. At 16 weeks, we show the level
of deTyr-MTs is significantly increased and co-localized to areas of myofibrillar malformation.
Profiling the enzyme complexes responsible for deTyr-tubulin, we identify vasohibin 2 (VASH2)
and small vasohibin binding protein (SVBP) significantly elevated in the mdx muscle at 4 wks. We
then use the genetic increase in VASH2/SVBP expression in 4 wk wild-type mice and find
densified deTyr-MTs that co-segregate with myofibrillar malformations similar to those in the 16
wk mdx. Given that no changes were identified in fibers expressing EGFP as a control, we conclude
that disease dependent densification of deTyr-MTs underscores the altered myofibrillar structure
in dystrophic skeletal muscle fibers.

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**Keywords: Dystrophy, skeletal muscle, myoarchitecture, microtubule network**
## 1. Introduction

Skeletal muscle fibers exhibit highly ordered myofibrillar structure which is essential for efficient force generation within the muscle fiber. Myofibrils (1µm diameter) are composed of individual contractile units (i.e., sarcomeres, ~2 µm in length) arranged in series to span the length of the muscle fiber (500µm to a few cm). The number of parallel packed myofibrils within the muscle fiber governs the contractile force. Until recently, myofibrils were thought to be independent units co-registered through protein links between their Z-line sarcomere boundaries. However, new evidence of sarcomeres branching between registered myofibrils¹ has redefined these structures as a continuous myofibrillar matrix that facilitates the highly coordinated, unilateral contraction of the muscle fiber.

In contrast to the registered myofibrillar matrix seen in healthy skeletal muscle, are myopathies such as Duchenne muscular dystrophy (DMD) where myofibrils become misaligned and torturous resulting in misorientation of force vectors and dyssynchronous activation of sarcomeres²⁻⁵. These changes result in significant reductions in isometric force and velocity of contraction as well as increased shear-stress that predisposes damage at these locations ⁶⁻⁸. While misalignment of the sarcomeres is now established as pathognomonic in DMD, the mechanisms that predispose their occurrence are unknown.

The cytoskeleton is a dynamic structural and signaling scaffold of microtubule (MT), actin and intermediate filaments (IF) that is essential for the intracellular trafficking, maintenance of cellular architecture, and positioning of organelles in all cells. Microtubules garnered early attention in DMD muscle as a cytoskeletal element that was disorganized early in disease and
became densified with disease progression⁹⁻¹¹. The discovery of dystrophin as an MT binding partner has resolved the mechanisms that initiate the disorganized MT structure in DMD¹¹ and inspired many groups to determine how these MT alterations may impact dystrophic pathology.

The structure and function of MTs are regulated by post-translational modifications (PTM) to their tubulin monomers. Detyrosination (deTyr), the reversible enzymatic removal of α-tubulin's COOH-terminal tyrosine, promotes the interaction of MTs with binding partners¹²⁻¹⁵. Work by our group has identified that MTs enriched in deTyr-tubulin (deTyr-MTs) regulate the stiffness of the muscle fiber cytoskeleton and thus the activation of NADPH Oxidase 2 (Nox2) dependent reactive oxygen species (ROS) and calcium (Ca²⁺) signals by mechanotransduction¹⁶,¹⁷. In the murine model of DMD (i.e., *mdx*), the densification of deTyr enriched MTs is a consequence of disease pathology that increases the passive mechanics of muscle fibers. Together with the increased expression of Nox2 proteins, these changes drive the excess mechanotransduction elicited Nox2-ROS and Ca²⁺ signals linked to dystrophic pathology¹⁶,¹⁸,¹⁷.

Our lab's previous work on dysregulated MT mechanotransduction in murine DMD (*mdx*) focused on murine models between 3-9 months, when pathology is entrenched yet progression is evident¹⁶,¹⁷. Within this timeframe of disease our group and others have previously identified and profiled the increased occurrence of muscle fibers with gross structural malformations (i.e., splitting, branching)²,³,⁷,¹⁹,²⁰ that increase the susceptibility to contractile damage in DMD. Here we were intrigued by work suggesting that these gross alterations in muscle fiber structure arose from structural changes in the myofibrils³. Informed by our observation of the MT densification often occurring in discrete areas in *mdx* muscle fibers¹⁶ ¹⁷, and evidence that microtubules are
essential for myofibrillar growth, maintenance, and repair<sup>21-24</sup>, we hypothesized a link between the
disease altered MTs and the occurrence of myofibrillar malformations in DMD.

In the present study, we focused early in disease pathology to bias our capture of the
mechanisms that underlie the development of myofibrillar malformations. In muscle fibers from
young mice (4 wks) we identify a small but significant increase in deTyr-MTs in *mdx*, yet no
evidence of myofibrillar malformation above WT. Profiling muscle fibers at 16 wks, we find the
level of deTyr-tubulin increases disproportionally in the *mdx* where it occurs largely in bundles of
MTs that co-localize with areas of myofibrillar malformation.

Microtubules participate in the highly orchestrated growth, maintenance, and repair of
myofibrillar structure<sup>24</sup>. In fact, an increased level of deTyr enriched MTs is an early and critical
event in sarcomerogenesis<sup>25,26</sup>. Profiling the enzyme complexes responsible for deTyr-tubulin we
found vasohibin 2 (VASH2) significantly elevated in the *mdx* muscle at 4 and 16 wks. To determine
the consequences of elevated VASH2 activity on myofibrillar structure we overexpressed VASH2
in muscles of 4wk wild-type mice. We show that VASH2 overexpression modeled the densification
of deTyr-MTs seen in the 16 wk *mdx*. Furthermore, we demonstrate that these deTyr-MTs co-
segregate with myofibrillar malformations comparable to those found in 16wk *mdx*. We conclude
that disease altered microtubules are an early event in dystrophic pathology that predisposes the
altered myofibrillar structure in dystrophic skeletal muscle fibers.

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# 2. Methods

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## 2.1 Animal Use

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All mice were obtained from Jackson Laboratories (Bar Harbor, MA) and studied with procedures approved by the University of Maryland, Baltimore Institutional Animal Care and Use Committee (IACUC).

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### 2.1.1 Electroporation

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Anesthetized mice (2% isoflurane) were injected with 25 µL of 1 mg/mL of hyaluronidase (Sigma-Aldrich) subcutaneously into the sterilized food pad of both hindlimbs. After 1 h, one footpad was injected with plasmid cDNA (20 µL at 1 µg/µL) containing a bicistronic construct of VASH2-GFP/SVBP (kind gift from Marie Jo Moutin²⁷) with the contralateral footpad receiving a plasmid cDNA containing EGFP as a control (1 µg/µL). The plasmid cDNA was then delivered to the flexor digitorum brevis (FDB) muscle by electroporation through sterile electrodes placed subcutaneously at the proximal and distal ends of the FDB. The pulse protocol consisted of 30 pulses of 150 V for 20 ms duration at a frequency of 1 Hz. Mice were humanely euthanized and FDBs harvested after 5 days.

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### 2.1.2 Muscle Fiber Preparation

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FDB muscles were harvested bilaterally in sterile mouse ringer and maintained overnight in DMEM (supplier) supplemented with collagenase A (Roche, 0.2 mg/ml) and 1% Penicillin-Streptomycin in a CO₂ incubator (37°C, 5% CO₂). Following gentle trituration to yield single FDB fibers, the cells were washed once in DMEM supplemented with 10% fetal bovine serum then washed twice in physiological Ringer solution with 1mM EGTA (pH, 7.4). Fibers were then either

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maintained in Ringer solution at room-temperature for live cell imaging or fixed in 4% PFA with 133
5mM EGTA for 20 minutes at room temperature, washed twice in PBS then stored in PBS with 134
0.4% sodium azide until used. 135

### 2.1.3 *In-vivo* contractile function

Contractile performance and injury susceptibility were tested in vivo as described 137 previously¹⁶,¹⁷. Anesthetized mice (2-3% isoflurane) were placed in supine position on the 138 temperature maintained (Deltaphase Isothermal Pad, Braintree Scientific) platform of an Aurora 139 3100 with the knee stabilized and foot affixed on the footplate of the torque transducer. The plantar 140 flexor muscle group (gastrocnemius, soleus) was activated by percutaneous stimulation. The force 141 frequency relationship was evaluated with 500msec trains of square pulses (0.1 ms) between 1 and 142 150Hz. The susceptibility to contraction force-loss was evaluated with 25 eccentric (i.e., 143 lengthening) contractions. 144

### 2.1.4 Mechanical properties

FDBs maintained in Ringer were placed on a glass-bottom dish coated with ECM (E6909; 146 Sigma-Aldrich). The near membrane mechanical properties of the FDB were quantified with a 147 Chiaro nano-indenter (Optics11) using a cantilever (0.044 N/m stiffness) with round probe (3-µm 148 radius). Indentation (1µM) profiles at speeds from 0.5 to 25 µm/s were analyzed with a Hertzian 149 contact model to calculate the Young's modulus (i.e., stiffness) of the FDB. 150

## 2.2 Western Blotting

Western blots were conducted as previously described¹⁷. Briefly, 20µg of clarified muscle was 152 processed via SDS-PAGE, transferred to nitrocellulose membranes, and washed with 5% milk 153
solution in PBS before blocking for 1h in the same solution. The membrane was probed overnight for β-tubulin (T4026, Sigma-Aldrich), deTyr-tubulin (31-1335-00, RevMAb Biosciences), acetylated tubulin (T7451, clone 6-11B-1; Sigma-Aldrich), and gp91phox (Abcam; ab129068). Next membranes were washed twice in 5% milk solution, incubated with the appropriate secondary antibody (1:10,000) at room temperature for 1h, and washed with 0.5% Tween solution in PBS twice for 10min. Blots were imaged and analyzed using the LICOR Odyssey CL-x system.

## 2.3 RT-qPCR

Gastrocnemius muscles were collected from *mdx* and WT mice and snap-frozen in isopentane cooled on dry ice. Tissues were later powdered and homogenized in TRI-reagent (Zymo Research). Phase separation was performed using 0.2mL of chloroform per 1mL of TRI-reagent, with samples shaken vigorously for 2min then centrifuged at 12,000 x g for 10min at 4°C. To precipitate RNA from the aqueous phase, 0.5mL isopropyl alcohol per 1 ml TRI-reagent used for lysis was added and incubated at room temperature for 10 minutes before centrifuging for 10 minutes at the aforementioned settings. The resulting RNA pellet was washed with 75% ethanol, centrifuged for 5 minutes at 7,500 x g at 4°C, then dissolved in 30µL DNase-RNase free water at which point RNA concentration was measured using a spectrophotometer. 2.5 µg samples of RNA were reverse transcribed using the SuperScript IV First-Strand Synthesis System (Invitrogen), following manufacturer protocol. Gene expression was measured via RT-qPCR using PowerUp SYBR Green Master Mix (Applied Biosystems) on the QuantStudio3 Real-time PCR System (Applied Biosystems by Thermo Fisher Scientific) and QuantStudio Design and Analysis software v1.5.2 (Applied Biosystems). Primers are listed in Supplementary Table 1.
## 2.4 Immunofluorescence and automated imaging

Fixed FDB fibers were blocked for 2h at room temperature in Superblock™ Blocking Buffer in PBS (Thermo Scientific) with 0.04% saponin. Fibers were incubated in an Eppendorf tube with primary antibodies to detect microtubule structure (beta-tubulin; T4026, Sigma-Aldrich) and the population of microtubule tubulin modified by detyrosination (deTyr-tubulin; 31-1335-00, clone RM444, RevMAb Biosciences USA, Inc.). Sarcomeric actin was decorated with phalloidin conjugated to Alexa Fluor 633 (A22284, Invitrogen) to visualize myofibrillar structure. Primary Antibodies and phalloidin were used overnight at 4°C. The following day FDB fibers were incubated with the appropriate secondary antibodies (diluted in PBS containing 0.04% saponin and 0.1% sodium azide) for 2 h at room temperature, washed three times in PBS, then mounted onto slides with ProLong Gold + Dapi mountant (Invitrogen).

Fixed FDB fibers were imaged on an inverted Nikon C2+ confocal fluorescence system using an automated protocol developed in NIS Elements AR JOBS. Single fibers were identified by their actin labeling (i.e., phalloidin 633) from a full-slide tile scanned image (10x air obj.). Fibers without evidence of bends or hypercontraction under visual inspection (30-50 fibers per slide) were logged as regions-of-interest (ROI). Each identified fiber ROI was imaged using an automated routine that identified the muscle fiber surface and collected a full thickness z-stack (0.5 µm steps; 4 frame average) at 40x (1.4 N.A. Plan Apo air obj.) and 1.3 Airy units which yielded 0.31 µm/pixel resolution.

### 2.3.1 Automated microtubule structural analysis

The properties of the microtubule network were quantified in NIS Elements AR General
Analysis 3. Briefly, an inverse binary mask of the phalloidin label (Cy5 channel) identified areas of myofibrillar structure (black) and areas of myofibrillar gaps (white) linked to the regions of altered continuity. Within each z-stack image the density of deTyr-tubulin (binarized deTyr- tubulin) was determined within the areas of myofibrillar structure and continuity gaps and normalized to the measured area. The area around the nuclei was masked to exclude any microtubule alterations around the nuclei as a confounding factor. For global density measures, the total deTyr-tubulin stain for each z-slice was normalized to the fiber area (as determined by the phalloidin label).

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## 3. Results

### 3.1 Young *mdx* mice exhibit functional deficits and microtubule alterations

Our lab's previous work on dysregulated MT mechanotransduction and gross structural alterations in *mdx* was in mice 3-9 months of age when pathology is well-established and still progressing. In this study, we examined wild-type (C57BL.10/J) and *mdx* (C57BL.10/J *mdx*) mice at 4 and 16 wks of age to elucidate the mechanisms that underlie the development of myofibrillar malformations. Our initial experiments sought to establish the functional status of the muscle at these ages. Evaluating *in vivo* plantar flexor function, we confirmed deficits in maximal isometric force in the *mdx* at both 4 and 16 wks (Fig 1A). Measuring the weight of the gastrocnemius muscle we identified no differences between genotypes at 4 wks yet a significant increase in the mass of the *mdx* gastrocnemius at 16 wks, a finding consistent with the pseudohypertrophy reported at this age (Fig 1B). Calculating the specific force (i.e., force normalized to mass) revealed no difference between genotypes at 4 wks yet a significant drop in the specific force of the *mdx* was observed at
16 wks (Fig 1C). Finally, evaluating isometric force loss following 20 *in vivo* eccentric contractions we again found no significant deficit between genotypes at 4 wks; this however, progressed to a significant decrease in *mdx* at 16 wks (Fig 1D). Taken together, we identified an acceleration in functional deficits after 4 wks of age in the *mdx*.

Our group, and others<sup>28-31</sup>, have implicated disease dependent MT alterations as negative disease modifiers in adult *mdx* mice with advanced pathology. Western blot profiling of gastrocnemius muscle from 4 wk *mdx* mice finds no significant alteration in the expression of tubulin protein, its modification by acetylation (acetyl), nor the abundance of gp91phox, the catalytic subunit of the reactive oxygen species (ROS) producing enzyme, Nox2. While not statistically significant, detyrosination (deTyr) shows a marked increase at this age (Fig 2). As disease progresses from 4 to 16 wks we identify an increased abundance of gp91phox, increased expression of tubulin protein as well as its modification by detyrosination (deTyr) and acetylation. Our published works have linked these tubulin changes to the increased myofiber passive mechanics (i.e., stiffness) and together with gp91phox, to the excess MT dependent mechanotransduction activation of Nox2-ROS and Ca<sup>2+</sup> signals that drive disease pathology in DMD.

## 3.2 *mdx* Mice Exhibit Malformed Myofibrillar Structure

Alterations in myofibrillar structure are a consequence of deficient myofibrillar repair following acute muscle damage<sup>32</sup> or disease pathology<sup>2,3,19</sup> that predisposes the occurrence of gross malformations in muscle structure (i.e., split and branched fibers)<sup>2</sup>. Our group previously reported a low percentage of grossly malformed (i.e., bifurcated, split, etc.) skeletal muscle fibers in 6-9 wk
238 *mdx*, with less than 10% abnormal fibers found in the flexor digitorum brevis (FDB)³³,³⁴. These
239 observations were made by visual inspection in brightfield where we found > 90% of the FDB
240 muscle fibers having no detectable abnormalities. In the current study, we show that when labeled
241 for myofibrillar structure (i.e., phalloidin labeled actin), and imaged with confocal microscopy, a
242 more significant number of muscle fibers with abnormalities in myofibrillar structure becomes
243 apparent (Fig 3).

244 Using an automated confocal strategy, we imaged single isolated FDB myofibers from WT
245 and *mdx* at 4 wks and 16 wks. Our qualitative visual inspection identified four distinct
246 morphologies of myofibrils within the muscle fiber: (1) canonical aligned striations; (2) evidence
247 of “braided” myofibril structure with misregistration; (3) misalignment characterized by
248 myofibrils wrapping around the peripheral myofibrils; (4) fibers exhibiting gross malformations
249 (i.e., branches, splits) as previously described³³ (Fig 3A-C). In the 4 wk *mdx*, myofibers with
250 evidence of braided myofibrils make up 22% of the total FDB population. By 16 wks, only 38%
251 of myofibers in the *mdx* FDB exhibit canonical aligned striations; 39% of fibers have braided
252 myofibrils, 14% have myofibrillar wrapping, and 9% of muscle fibers are branched (Fig 3C).
253 Although muscle fibers with altered myofibrillar morphologies were identified in the wild-type,
254 these comprised less than 11% of the total FDB population.

255 While fibers with canonical aligned striations were in the majority in both genotypes, a
256 significant number of otherwise normal *mdx* fibers presented with separations between myofibrils,
257 marked by bundles of microtubules (Fig 3C). In fact, these myofibrillar separations were evident
258 in a majority of 16 wk *mdx* muscle fibers and were observed in some wild-type FDBs albeit at a
259 markedly reduced occurrence. These initial qualitative observations provided the basis for our
260 adopting quantitative methods. Informed by our past observations of the MT densification often
261 occurring in discrete areas in mdx muscle fibers (Khairallah [Fig 2], Kerr [Fig 1]), and new
262 qualitative evidence that this MT densification is coincident with altered myofibrillar structure, we
263 posited a link between the disease altered MTs and the occurrence of myofibrillar malformations
264 in DMD.
265

3.3 The Continuity of Z-line Striations as a Metric of Myofibrillar Structure
265

We next quantified the continuity of myofibrillar Z-line striations as a metric of myofibrillar
266 structure. The quantitative assessment of Z-line striations in each image was performed using a
267 custom MATLAB routine established by Morris et al.³⁵ for profiling myofibrillar structure in
268 developing cardiomyocytes and skeletal myotubes and adapted here for mature skeletal muscle
269 fibers.
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In brief, multi-channel Nikon confocal fluorescence images were converted into RGB TIFFs
271 and the Cy5 channel containing phalloidin-633 for actin was output to a new image stack. To
272 decrease processing time, each image was cropped to the fiber of interest and rotated to align the
273 myofiber long-axis to the x-axis of the frame. The maximal myofiber boundary was identified with
274 an Otsu's threshold³⁶ of the maximum intensity projection (max-IP) of the z-stack (Fig 4A). In the
275 resulting binary image, the myofiber area was calculated and orientation axis determined using the
276 least mean square orientation estimation algorithm. Subsequently, each z-slice was binarized based
277 on Otsu's threshold and compared to the binarized maximum intensity projection to determine the
278 ratio of "true" pixels in the z-slice of interest compared to the max-IP (Fig 4B-D). Z-slice ratios
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280 above a user defined threshold (nthresh = 0.6) were selected for analysis (Fig 4E).

281 For each analyzed z-stack image, the minor axis length (i.e., width) of the myofiber was 282 determined for every 20 pixels along the long axis of the muscle fiber (Fig 5A). Subsequently the 283 phalloidin labeled Z-line structure was detected via the ZlineDetection algorithm developed by 284 Morris et al.³⁷. The continuity of each Z-line was determined by its length divided by the nearest 285 minor axis length and plotted with color code (Fig 5B) with a continuous Z-line spanning the 286 muscle fiber perpendicular axis yielding a measure of 1 (red), with interruptions in the continuity 287 yielding lower values and cooler colors. The output for each fiber includes a mean striation 288 continuity score for each z-slice as well as a boxplot for the entire z-stack (Fig 5C). The median 289 striation length for the entire muscle fiber z-stack is also reported as the continuity score.

290 **3.4 Z-line continuity decreases with dystrophic progression and predicts altered myofibrillar structure.**

291 We show that striation continuity effectively identifies the minor interruptions in the 292 myofibrillar structure seen in WT muscle fibers as well as the more significant disruptions in the 293 *mdx* (Fig 6A-F). We demonstrate continuity scores < 0.4 in fibers displaying areas of myofibrillar 294 braiding (Fig 6B,D), while scores decrease to < 0.2 in fibers with significant myofibrillar wrapping 295 (Fig 6C,F).

296 Using the automated imaging strategy and ZlineDetection to quantitate Z-line continuity, we 297 screened FDB myofibers from WT and *mdx* mice at 4 wks and 16 wks of age. At 4 wks we find 298 no significant difference in the mean striation continuity of the entire fiber between *mdx* and WT 299 (Fig 6G), the 4-wk *mdx* do achieve lower minimum continuity scores than the 4-wk wild-type. 300
Because severely misaligned myofibrils are a rare event at 4 wks, the reduced continuity score is manifest from microtubule bundles between myofibrils causing separations as previously described (Fig 3A).

By 16 wks, the differences between morphology in *mdx* and wild-type become more pronounced. There is an increase in the average continuity score in wild-type, while the continuity of the *mdx* decreases as altered morphologies emerge at a greater frequency (Fig 6G). This increase in the proportion of separations between myofibrils in the mdx between 4 and 16 wks suggests a progression of myofibrillar alterations with disease progression.

### 3.5 Increased tubulin detyrosination occurs commensurate with myofibrillar malformations

Examining FDB myofibers labeled for deTyr-tubulin and actin we find a significant increase in the density of deTyr-MTs in the *mdx* at 4 wks that progresses at 16 wks (Fig 7A-B). Revisiting our previous observation of bundled microtubules coinciding with myofibrillar separations, we quantified the density of deTyr-tubulin within the regions of myofibrillar separation versus the density in areas with otherwise normal myofibrillar connectivity (Fig 7C-E). At 4 wks we find no difference in the density of deTyr-MTs between these areas in WT, but in the *mdx* we find a significant increase in deTyr-MTs only in the regions of myofibrillar separation.

In 16 wk WT muscle fibers we again find no difference in the density of deTyr-MTs within the regions of myofibrillar separation versus areas with otherwise normal myofibrillar connectivity. In the 16 wk *mdx* we again find a significant elevation in deTyr-MTs in areas of myofibrillar separation but now find these changes in areas of otherwise normal myofibrillar connectivity as well. (Fig 7E). Taken together, these results suggest that deTyr-MTs become abundant first
between myofibrils then progress more globally throughout the myofibrillar structure as disease progresses.

**3.6 Overexpression of VASH2-SVBP models the increased tubulin detyrosination, cytoskeletal stiffness, and myofibrillar malformations established as pathognomonic in mdx.**

Detyrosination is the reversible enzymatic cleavage of the COOH-terminal tyrosine from α-tubulin by vasohibin 1 (VASH1) or vasohibin 2 (VASH2) and their partner the small vasohibin binding protein (SVBP)¹³,³⁸. Examining the transcripts of these proteins at 4 wks (Fig 8A-D) finds no significant change in VASH1 yet a significant increase in VASH2 and SVBP in the *mdx*. At 16 wks the VASH1 remained unchanged, while VASH2 remained elevated in the *mdx*, and SVBP now exhibited no difference between genotypes (Fig 8F-I).

Microtubules are essential for myofibrillar growth, maintenance and repair²¹⁻²⁴. Given that bundles of deTyr modified MTs are associated with myofibrillar separations in 4 wk *mdx* muscle fibers that then progress to more significant changes at 16 wks, we sought to determine if an experimental increase in deTyr-MTs in 4 wk WT fibers was sufficient to recapitulate the changes seen in the 16 wk *mdx*. To this end we used electroporation to introduce VASH2-GFP/SVBP cDNA, or eGFP cDNA as a control, into the 4 wk old mouse FDB and examined the muscle fiber properties 5-7 days later (Fig 9A-C).

In WT FDB fibers transduced with VASH2-GFP/SVBP we find an increased abundance of deTyr-enriched MT bundles when compared to the eGFP controls (Fig 9A-C). In *mdx* muscle fibers we previously linked the elevated levels of deTyr-MTs to an increase in cytoskeletal mechanics (i.e., stiffness)¹⁷,³⁹. Using nanoindentation to measure the viscoelastic properties we
343 show a significant increase in passive stiffness in VASH2-GFP/SVBP over expressing FDB muscle
344 fibers compared to the eGFP control (Fig 9D).

345 Our evidence suggests a link between the increased abundance of deTyr-enriched MTs and the
346 altered myofibrillar structure in the 16 wk *mdx*. Consistent with this finding in *mdx* was a
347 significant reduction in striation continuity in WT fibers overexpressing VASH2-GFP/SVBP
348 determined by Z-line detection reporting (Fig 10C). Further evidence supporting a link between
349 deTyr-MTs and myofibrillar malformations came from visual inspection that revealed a 21.3%
350 occurrence of braided myofibrils in the VASH2-GFP/SVBP over-expressing myofibers with no
351 evidence for this occurrence in controls. Concordant with our finding deTyr-enriched MTs co-
352 registered with myofibrillar malformations in the *mdx* (Fig7), in VASH2-GFP/SVBP over-
353 expressing myofibers we found bundled deTyr-MTs abundant only in areas with altered
354 myofibrillar structure (Fig 10E).

355 Taken together, our results demonstrate that VASH2-GFP/SVBP overexpression in WT
356 muscle is sufficient to model the altered myofibrillar structure that arises in the 16 wk *mdx*. We
357 posit that disease altered microtubules are an early event in dystrophic pathology that predispose
358 the altered myofibrillar structure in dystrophinopathies.

359

360 **Discussion**

361 Altered myofibrillar structure is a consequence of dystrophic pathology in humans<sup>40,41</sup> and
362 rodents <sup>3,4,19,42</sup> that decreases force production and increases susceptibility to contraction injury.
363 In fact, evidence from optical predictions<sup>8</sup> and mathematical models<sup>6</sup> suggests myofibril
misalignment and myofibrillar stiffness are the dominant factors contributing to decreased isometric force and contraction velocity<sup>6</sup> in dystrophinopathies. While the consequences of altered myofibrillar structure are well defined, the mechanisms that underlie their occurrence have been elusive.

Microtubules play a critical role in the myofibrillar growth, maintenance, and repair of striated muscle through the patterned recruitment of myosin, actin, mRNAs, and ribosomes for the assembly of sarcomeres<sup>24</sup>. Consistent with tubulin PTM's as regulators of MT function, MTs enriched in deTyr-tubulin have been implicated in the regulation of mechanotransduction dependent ROS and $Ca^{2+}$ signals, in the directional transport of cargo (i.e., lysosomes), and in the highly orchestrated myofibrillar assembly during myogenesis<sup>25,26</sup>.

Here we show that myofibrillar malformations are not inherent to dystrophin's absence, rather they arise in the *mdx* between 4 and 16 weeks of age coincident with the densification of deTyr-enriched MTs in these malformed areas. Transcriptional evidence of increased VASH2 and SVBP in the 4 wk *mdx* suggested deTyr-enriched MTs may be responsible for the myofibrillar alterations arising by 16 wks. Evidence supporting a causative link was established by showing that VASH2-GFP/SVBP overexpression in WT muscle fibers was sufficient to recapitulate the densification of deTyr-enriched MTs and the altered myofibrillar structure seen in the 16 wk *mdx*. These results suggest that the enhanced expression/activation of VASH2/SVBP in dystrophic muscle fibers is an early event in dystrophic pathology that drives the densification of deTyr-MTs to disrupt myofibrillar maintenance and/or repair.

This report extends our previous discovery implicating deTyr enriched MTs as an early event
385 in DMD pathology that drives the excess mechanotransduction elicited Nox2-ROS and Ca²⁺

386 signals linked to dystrophic progression¹⁶,¹⁸,¹⁷. Together these results give strong support for

387 disease altered MTs as negative disease modifiers early in DMD pathology. Given the

388 transcriptional and proteomic evidence for these alterations in DMD patients¹⁶,⁴³, targeted

389 therapeutics to reduce deTyr-MTs may be a viable option to slow dystrophic progression. As

390 pharmacologic approaches are developed, future studies to genetically reduce VASH1, VASH2,

391 or SVBP in mdx fibers will further advance our mechanistic understanding.

392 While here we focused on DMD, it is notable that myofibrillar malformations were modeled

393 in WT muscle fibers by VASH2/SVBP overexpression. This result demonstrates that dystrophin's

394 absence is not obligate in this process, nor is the dysregulated signaling linked to dystrophic

395 pathology. It is then tempting to speculate that the occurrence of myofibrillar malformations seen

396 in aging muscle⁴⁴,⁴⁵, in disparate genetic diseases⁴⁶, and in conditions of supraphysiologic muscle

397 growth⁴⁷,⁴⁸ may be driven by this same axis. In this regard, future work profiling these

398 conditions, and mechanisms that increase VASH/SVBP expression and activity, will likely yield

399 insights of broad importance.

400
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<figure><img src="image_2.png" /><figcaption>Fig 1. Force production decreases with age in *mdx* while injury susceptibility increases.</figcaption></figure>

(A) Isometric force produced at 150Hz in wild-type (white) and *mdx* (red) gastrocnemius muscles at 4 and 16 wks, respectively. (B) Weights of the gastrocnemius muscles from WT and *mdx* mice used to normalize force production to measure (C) specific force production in age-matched WT and *mdx* mice at 150 Hz. (D) Percent force production in gastrocnemius muscles from 4 and 16 wk WT and *mdx* mice after 25 eccentric contractions. *Mdx* mice experience increased force loss when compared to wild-type. Values are means ± SEM. All analysis are one-way ANOVA with Šídák's multiple comparisons test (* P < 0.05; ** P < 0.01; ***P<0.001; ****P < 0.0001)

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<figure><img src="image_3.png" /></figure>

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<figure><img src="image_5.png" /></figure>

<figure><img src="image_6.png" /></figure>

<figure><img src="image_7.png" /><figcaption>**Fig 2.** Detyrosination increases with dystrophic disease progression. (A) Western blots of the gastrocnemius muscle from wild-type and *mdx* mice at 4 wks (n=4) and 16 wks (n=3) (B) increased tubulin abundance and post-translational modification by (C) detyrosination and (D) acetylation. (E) We also see a significant increase in the NOX2 subunit *GP91<sub>phox</sub>*. Values are means ± SEM. All analysis are one-way ANOVA with Šídák's multiple comparisons test (* P*<*0.05*; ** P*<*0.01*)</figcaption></figure>
<figure><img src="image_8.png" /></figure>

<figure><img src="image_9.png" /></figure>

<figure><img src="image_10.png" /></figure>

<figure><img src="image_11.png" /></figure>

<figure><img src="image_12.png" /></figure>

<figure><img src="image_13.png" /><figcaption>**Fig 3.** Qualitative survey of fiber morphology (A) The percent distribution of each morphology was determined based on a survey of approx. of 150 fibers (n = 5, ~30 fibers/animal) from each condition. (B) Representative images showing aligned striations at 4 and 16 wks with canonical rectilinear MT structure. MT bundling is apparent in *mdx* as early as 4wks. Detyrosinated tubulin appears increased in 16 wk *mdx*. (C) Fibers from 16 wk *mdx* representative of altered morphologies, showing areas of MT bundling appearing coincident with alterations in striation continuity, in fibers with myofibrillar malformation as well as in fibers with otherwise aligned striations.</figcaption></figure>

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**Fig 4. Automated image selection for myofiber analysis (A)** Each z-slice is binarized and compared with the binarized maximum intensity projection (max-IP). (**B**) Within the current z-slice pixels with sufficient intensity for analysis are logged as true. The total number of true pixels in the current z-slice is compared as a ratio between true pixels in the max-IP. (**C**) A representative selection shows the fiber area of 4th z-slice (outlined in orange) was less than 60% of the max-IP area whereas (D) the fiber area of 17th z-slice exceeded the 60% threshold. (E) A representative plot of the z-slices to be included in analysis based on the user defined threshold (nthresh = 0.6).
A

<figure><img src="image_14.png" /></figure>

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<figure><img src="image_15.png" /></figure>

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<figure><img src="image_16.png" /></figure>

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<caption>**Fig 5.** Striation continuity quantification. (A) Representative z-slice of phalloidin channel of WT FDB shown in Fig. 2. Insets (right) showing Z-line structure. (B) Automated Z-line detection output from ZlineDetection. Insets (right) showing the heterogeneity of Z-line continuity in WT FDB at minor separations between myofibrils and the nucleus. (C) Quantification of the striation length as compared to myofiber width within each z-slice (left) and the corresponding box plot for the entire z-stack (right).</caption>
<figure><img src="image_17.png" /></figure>

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<caption>**Fig 6.** Median Striation Continuity scores decrease with altered morphology. Striation detection for (A) the full 16wk wild-type fiber depicted in Figure 1, (B) the 16wk mdx fiber with braided myofibrils, and (C) the 16wk mdx fiber with wrapped myofibrils. Their respective continuity scores per z-slice are illustrated with min, max, median scores displayed in boxplots (E-F). (G) Average striation continuity scores for each fiber within each experimental group (n=50-125 fibers per condition). One-way ANOVA revealed significant difference between groups with Šídák's multiple comparisons test elucidating a significant difference between 16wk WT and MDX (*F*(3, 333) = 38.36, *p* < 0.0001).</caption>
<figure><img src="image_24.png" /><figcaption>**Fig 7.** Detyrosination is enriched in dystrophic fibers and at the site of myofibrillar malformation. (A-B) We show the average deTyr density normalized to fiber size via the actin stain in fibers from wild-type and *mdx* mice 16 wks (n = 5 mice). Values are means ± SEM. Statistical significance determined using t-tests. (C) Representative images showing co-localization of detyrosinated tubulin with myofibrillar break sites. (D-E) Quantification of deTyr-tubulin density in regions with continuous striations across myofibrils vs. within break sites. Analysis was completed using one-way ANOVA with Šídák's multiple comparisons test (\*\*P < 0.01; \*\*\*P < 0.001; \*\*\*\*P < 0.0001).</figcaption></figure>

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<figure><img src="image_25.png" /><figcaption>**Fig 8.** Expression of vasohibins and SVBP are increased in dystrophy. (A-D) Relative expression of VASH1, VASH2, and SVBP at 4 wks reveals (B) VASH1 expression is not significantly increased at 4 wks whereas (C) VASH2 expression is increased approximately 18-fold on average in the *mdx* and (D) SVBP is increased approximately 4-fold. (E) Relative gene expression of both tubulin carboxypeptidases, and especially VASH1, appears increased in wild-type 16 wk compared to 4 wk and *mdx* expression relative to wild-type remains increased at 16 wks. (F) VASH1 shows a trend toward increased expression in *mdx*, only (G) VASH2 expression remains significantly increased, while its binding partner (H) SVBP is not significantly increased in *mdx* at 16 wks. All analysis was performed using one-way ANOVA with Šídák's multiple comparisons test (*P* < 0.05; **P**<0.001; ***P* < 0.0001).</figcaption></figure>

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<figure><img src="image_29.png" /><figcaption>Fig 9. VASH2 overexpression increases the density of deTyr-MTs and the passive stiffness of the muscle fiber (A) Representative image muscle fiber expressing EGFP(+) as a control finds deTyr-tubulin enriched MTs dispersed throughout the fiber. (B) VASH2+GFP expressing muscle fiber exhibits a denser and more bundled network of deTyr-tubulin enriched MTs. (C) Quantification of the density of deTyr-MTs finds a significant elevation with VASH2 overexpression (n=3 mice per condition; 10-15 fibers per mouse). (D) Nano-indentation (5µm/sec) measure of viscoelastic resistance finds increased stiffness in VASH2-GFP muscle fibers. (t-test, * P < 0.05; ****P < 0.0001).</figcaption></figure>
A +EGFP Control

DeTyr+ DAPI+
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<figure><img src="image_30.png" /></figure>

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+Phalloidin
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F VASH2 +Phalloidin+DAPI

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<figure><img src="image_40.png" /><figcaption>**Fig 10.** Detyrosinated microtubules are enriched at areas of Z-line disruption</figcaption></figure>

(A-B) Representative images of EGFP and VASH-SVBP electroporated fibers respectively, with insets showing the disparate morphology. A closer look at the inset (B) reveals extensive veneering of striations, perhaps indicative of the onset of myofibrillar “braiding”. (C)

Quantification of striation continuity using Z-line detection. There was a statistically significant difference between groups as determined by t-test (D) Representative image of EGFP(+)VASH2 myofiber showing myofibrillar break sites identified as regions-of-interest (ROI) within NIS-Elements. (E) The density of the deTyr-MTs within each ROI was calculated and normalized to the respective area. There was a statistically significant difference between groups as determined by one-way ANOVA with Šídák's multiple comparisons test (F(3,92) = 25.12). (F)

Representative image of VASH2-SVBP electroporated FDB fiber with evidence of altered myofibrillar directionality (i.e., braiding). (*P* < 0.05; ***P* < 0.001; ****P* < 0.0001).

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<table><thead><tr><td>Gene</td><td>Forward</td><td>Reverse</td></tr></thead><tbody><tr><td>VASH1</td><td>GGGAAGACTTCCGCAAGGA</td><td>CCTGTCCCCTTGCCAATCT</td></tr><tr><td>VASH2</td><td>CATCAGGAAGGAGCTGGAGAAG</td><td>TGGGCACTTGCAGGTTTCA</td></tr><tr><td>SVBP</td><td>TTCAGAAGCCAAACCATGGAT</td><td>CTCCACTCTGAAGGCTGGTTCTT</td></tr></tbody></table>

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<caption>Supplementary table 1. Primer sequences for RT-qPCR</caption>