Mef2c-Dependent Downregulation of Myocilin Mediates Cancer-Induced Muscle Wasting And

Author Manuscript Published OnlineFirst on March 4, 2020; DOI: 10.1158/0008-5472.CAN-19-1558 Author manuscripts have been peer reviewed and accepted for publication but have not yet been edited.

MEF2c-dependent downregulation of Myocilin mediates cancer-induced muscle wasting and

associates with cachexia in cancer patients

Sarah M. Judge1*, Michael R. Deyhle1, Daria Neyroud1, Rachel L. Nosacka1, Andrew C.

1, Miles E. Cameron1,2, Ravneet S. Vohra3, Ashley J. Smuder4, Brandon M. Roberts1,

Chandler S. Callaway1, Patrick W. Underwood2, Stephen M. Chrzanowski3, Abhinandan Batra3,

Meghan E. Murphy1, Jonathan D. Heaven1, Glenn A. Walter3, Jose G. Trevino2 and Andrew R.

Judge1*

1Department of Physical Therapy; 2Department of Surgery and 3Department of Physiology,

College of Medicine; University of Florida Health Science Center, Gainesville FL 32610, USA.

4Department of Health and Human Performance, University of Florida, Gainesville FL 32610,

USA.

*Correspondence to: Andrew R. Judge, Ph.D. or Sarah M. Judge, Ph.D. Department of Physical Therapy Department of Physical Therapy University of Florida University of Florida Gainesville, FL 32610 Gainesville, FL 32610 Ph: 1 (352) 273-9220 Ph: 1 (352) 273-9146 Email: [email protected] Email: [email protected]

Running title: Loss of Myoc via MEF2c mediates cancer cachexia

Conflict of interest statement: The authors declare no potential conflicts of interest.

Downloaded from cancerres.aacrjournals.org on October 2, 2021. © 2020 American Association for Cancer Research. Author Manuscript Published OnlineFirst on March 4, 2020; DOI: 10.1158/0008-5472.CAN-19-1558 Author manuscripts have been peer reviewed and accepted for publication but have not yet been edited.

Abstract

Skeletal muscle wasting is a devastating consequence of cancer that contributes to increased complications and poor survival, but is not well understood at the molecular level. Herein we investigated the role of Myocilin (Myoc), a skeletal muscle hypertrophy-promoting protein that we showed is downregulated in multiple mouse models of cancer cachexia. Loss of Myoc alone was sufficient to induce phenotypes identified in mouse models of cancer cachexia, including muscle fiber atrophy, sarcolemmal fragility and impaired muscle regeneration. By 18 months of age, mice deficient in Myoc showed significant skeletal muscle remodeling, characterized by increased fat and collagen deposition compared to wild type mice-thus also supporting Myoc as a regulator of muscle quality. In cancer cachexia models, maintaining skeletal muscle expression of

Myoc significantly attenuated muscle loss, while mice lacking Myoc showed enhanced muscle wasting. Further, we identified the Myocyte enhancer factor 2 C (MEF2C) transcription factor as a key upstream activator of Myoc whose gain-of-function significantly deterred cancer-induced muscle wasting and dysfunction in a pre-clinical model of pancreatic ductal adenocarcinoma

(PDAC). Lastly, compared to non-cancer control patients MYOC was significantly reduced in skeletal muscle of PDAC patients defined as cachectic and correlated with MEF2c. These data therefore identify disruptions in MEF2c-dependent transcription of Myoc as a novel mechanism of cancer-associated muscle wasting that is similarly disrupted in muscle of cachectic cancer patients.

Statement of Significance

This work identifies a novel transcriptional mechanism that mediates skeletal muscle wasting in murine models of cancer cachexia that is disrupted in skeletal muscle of cancer patients exhibiting cachexia.

INTRODUCTION

Cancer-associated cachexia is a multifactorial syndrome characterized by the involuntary loss of

body and skeletal muscle mass (with or without fat loss), which cannot be completely reversed

through nutritional support (1). Skeletal muscle loss reduces tolerance to cancer treatments,

increases complications following cancer surgery and is strongly predictive of reduced survival

(2). However, due to the inherent complexity of cancer cachexia, effective therapeutics to preserve

muscle mass in cancer patients are not currently available.

The etiology of cancer-associated muscle wasting involve impaired protein homeostasis

and negative energy balance that are driven by a number of factors, including increased

metabolism, reduced appetite leading to reduced caloric intake, and chronic systemic inflammation

induced by both host- and tumor-derived factors (1,3,4). Inflammatory cytokines may play a role

in the wasting process through several mechanisms including interference of centrally-mediated

pathways involving the hypothalamus which regulate satiety and hunger, and through direct

stimulation of pathways which lead to an imbalance in muscle protein breakdown and synthesis

(5). In addition to inflammatory cytokines, other circulating factors dysregulated in tumor-bearing

hosts may also contribute to muscle wasting, including TGF- family members, Angiotensin II, parathyroid hormone-related protein, heat shock proteins and tumor-derived exosomes (1). For a more detailed review of the systemic mechanisms involved in cancer-associated cachexia and muscle loss derived from experimental models and patient samples, readers are referred to the work by Baracos et al. (1).

The intracellular mechanisms whereby muscles atrophy in response to cancer have largely been derived from animal models and, similar to other models of muscle wasting, support increased muscle protein breakdown coupled with reductions in protein synthesis as key mechanisms of wasting (6,7). Despite this, less convincing data exists to support an imbalance in

these pathways in muscle of cancer patients with cachexia. However, mechanistic studies in mouse

models of cancer cachexia indicate that muscle membrane damage (8) coupled with impaired muscle regeneration and repair (9) may also play causative roles in cancer associated muscle loss. In support of these findings, disruptions to membrane architecture (8) and muscle damage

(10) have been observed in muscle biopsies from cachectic cancer patients. Yet, the mechanisms that induce these phenotypes are not known.

Previous studies profiling the transcriptional networks changed in mouse skeletal muscle in response to tumor burden have provided key insight into pathways that may be involved in the muscle wasting phenotype (11), including signaling through the atrophy-associated Forkhead Box

O (FoxO) transcription factors (12). Indeed, multiple atrophy associated genes involved in muscle protein breakdown are upregulated through the FoxO transcription factors (12,13), which mediate muscle atrophy in mice bearing Lewis Lung Carcinoma (14) and C26 tumors (12). In addition to upregulating genes involved in muscle proteolysis, we also identified FoxO factors as key upstream factors necessary for the cancer-induced repression of many downstream target genes

(12). While the significance of these repressed targets are not yet clear, these findings may be particularly relevant to human cancer cachexia, since muscle biopsies from cachectic cancer patients display increased levels of FoxO1 (10,15), and significant gene repression (16).

One FoxO target gene repressed in skeletal muscle in response to C26 tumor burden that could play a role in the atrophy phenotype is Myocilin (Myoc). Although few studies have examined the role of Myoc in skeletal muscle, Myoc has been identified as a pro-hypertrophic protein that binds and stabilizes the dystrophin glycoprotein complex (DGC) (17), which is well- established to stabilize the muscle fiber membrane. Myoc is most well-known for its role in the pathogenesis of glaucoma, where genetic mutations in MYOC lead to mis-folding and aggregation

of MYOC protein within the trabecular meshwork, resulting in high intraocular pressure (18).

-syntrophin at the sarcolemma (17). In transgenic mice overexpressing Myoc, skeletal muscles show a ~40% increase in skeletal muscle mass and suppression of the FoxO signaling axis (17). These mice also exhibit increased protein levels of dystrophin and enhanced binding of 1-syntrophin to dystrophin implicating Myoc as a key regulator of DGC stability and, by association, sarcolemma stability.

Although muscles from mice lacking Myoc have not been fully characterized, these mice show

-syntrophin to dystrophin (17), suggesting that in conditions of Myoc deficiency, the sarcolemma may be less stable.

In the current study we determined whether Myoc downregulation is a common response in multiple models of cancer cachexia, and further investigated the biological significance of Myoc deficiency on skeletal muscle in isolation and in the context of cancer cachexia. Through these studies we demonstrate that Myoc is downregulated in multiple models of cancer cachexia and that MYOC gain-of-function deters muscle wasting in the C26 model of cachexia, and in a murine model of PDAC-associated cachexia (19), involving orthotopic implantation of pancreatic tumor cells isolated from a syngeneic C57BL/6 KRASG12D P53R172H +/+ (KPC) mouse. We further identify Myocyte enhancer factor 2C (MEF2c) as a key upstream transcription factor necessary for maintaining Myoc expression in skeletal muscle, whose gain-of-function prevents Myoc downregulation and deters cancer-induced muscle wasting and dysfunction. Lastly, using patient biopsies we demonstrate that MYOC mRNA is reduced in skeletal muscle of cachectic PDAC patients, and correlates with the levels of MEF2c, thus indicating that these findings may be relevant to muscle wasting in cancer patients.

MATERIALS AND METHODS

Mice and mouse models of cancer cachexia

The University of Florida Institutional Animal Care and Use Committee approved all animal

procedures described herein. Mice were housed in a temperature-controlled facility with a 12 hour

light/dark cycle, with standard diet and water provided ad libitum. Myocilin null (Myoc-/-) mouse

breeding pairs were received from Dr. S Tomarev from the National Eye Institute, and have been

described previously (20).

Murine Colon-26 (C26) cells (NCI-DTP Cat# Colon 26, RRID:CVCL_0240) were

purchased from the National Cancer Institute Cell Repository (Bethesda, MD). Culturing and

subcutaneous inoculation of C26 cells into CD2F1 mice to induce cachexia was performed as

described previously (14). Panc02 (RRID:CVCL_D627) murine pancreatic adenocarcinoma cells

and Panc02-H7 cells (RRID:CVCL_D628), which are a highly metastatic variant of Panc02 (21), were received from Dr. Min Li at The University of Oklahoma. For the orthotopic KPC (LSL-

KrasG12D/+;LSL-Trp53R172H/+;Pdx-1-Cre) model of cancer cachexia (19), we acquired KPC

FC1245 tumor cells derived from a KPC mouse backcrossed to the C57BL/6 genetic background from Dr. David Tuveson (Cold Spring Harbor Laboratory, Cold Spring Harbor, New York). The authenticity of the KPC FC1245 cell line has been described previously (22). Cell lines were verified to be mycoplasma free prior to receipt or by IDEXX BioAnalytics using PCR evaluation.

Tumor cells were maintained in either RPMI (Panc02/Panc02-H7) or DMEM (KPC) supplemented with 10% FBS, 1% penicillin, 1% streptomycin in a humidified chamber at 37 C with 5% CO2.

Panc02 and Panc02-H7 cells (1x106) or KPC cells (2.5x105) were orthotopically injected into the pancreas of 10-12 week-old male and female C57BL/6 wild type (WT) mice (The Jackson

Laboratory, Bar Harbor, ME) and Myoc-/- mice as described previously (23). For orthotopic studies, Sham mice underwent a comparable Sham surgery, in which the pancreas was surgically

exposed and injected with sterile saline. Mice were euthanized when tumor-bearing groups reached

IACUC-mandated experimental endpoint criteria based on deterioration of body condition score,

which is reflective of estimated tumor-free body weight loss of 15% and/or signs of pain or distress, including hunched posture and failure to groom. Cell lines were verified to be mycoplasma free prior to receipt or by IDEXX BioAnalytics using PCR evaluation within the last

6 months. All cell lines were utilized prior to passage 13.

Additional pre-clinical models of cancer cachexia, including those which utilize human

L3.6 pancreas-liver (L3.6pl) cells (RRID:CVCL_0384) and the PDAC patient-derived xenograft

(PDX) model, in which tibialis anterior (TA) muscles were utilized for qRT-PCR analyses of

Myoc and Mef2c, were described previously (23-25). Briefly, human L3.6pl pancreatic cancer cells were injected orthotopically into the pancreas or subcutaneously into the flank of immunocompromised NOD.Cg-Prkdcscid IL2rgtm1Wjl/SzJ (NSG) mice. PDAC tumor fragments were either sutured to the pancreas or subcutaneously implanted into the flank of NSG mice. Sham mice for the orthotopic L3.6pl and PDX studies underwent a comparable surgery, in which the pancreas was surgically exposed and either injected with sterile saline (L3.6pl studies) or a suture placed loosely around the pancreas and then removed (PDX studies). The L3.6pl cell line was authenticated within 6 months of receipt by short tandem repeat (STR) analysis.

Patient samples

Attainment of biospecimens from non-cancer control subjects undergoing benign abdominal surgery and from PDAC patients undergoing tumor resection surgery was compliant with an approved Institutional Review Board protocol, with written informed consent received from all

patients. Detailed demographic information on the patients included in this study were previously

published (10), and can be found in Supplemental Tables S1 and S2.

Body Composition Analysis

Pre-operative computed tomography (CT) scans were identified. An axial image at the level of the

third lumbar (L3) vertebra was accessed for each patient; the lower aspect of the L3 transverse

processes was used to guide image acquisition. Cross-sectional area at the L3 vertebra correlates

to total body composition (26,27). Manual image segmentation was performed using

that correspond to the radiodensities of skeletal muscle and adipose. Cross-sectional skeletal muscle area (cm2) was calculated by the summative area of -30 and 0 Hounsfield Units (HU), 1 and 25 HU, and 26 and 150 HU in order to further characterize myosteatosis (28). Intermuscular adipose cross-sectional area was measured between -150 and -50 HU. Skeletal muscle area was normalized to height (m2) yielding a lumbar skeletal muscle index (SMI, cm2/m2). Mean muscle attenuation (MA) is reported in HU for the entire muscle area at the L3 vertebrae.

Downhill running and assessment of muscle damage

Mice were subjected to downhill treadmill running at a decline of 14 at a speed of 8 to 10 m/min, for 45 min as described previously (29). To assess muscle damage, Magnetic Resonance Imaging

(MRI) was performed on hindlimb muscles in a 4.7T, horizontal bore magnet (Agilent; VMJ version 3.1) immediately following completion of the running protocol. This protocol is detailed in Supplemental Methods, and has been described previously (30,31). Six hours following treadmill running, mice received an intraperitoneal (IP) injection of 2.5mg EBD)

per 25g BW dissolved in 100µL sterile saline, and the next day muscles were harvested for

cryosectioning and histological analyses of EBD uptake and morphology.

Cardiotoxin injury

To study muscle regenerative capacity and subsequent regrowth, TA muscles of 10-12 week-old

male and female WT and Myoc-/- mice were injected with 75 µL of 10 µM cardiotoxin

(Calbiochem) and harvested 2, 4 or 6 weeks post-injury.

AAV vectors and delivery, Plasmid injections

AAV9-tMCK-MYOC-polyA-CMV-GFP-polyA and AAV9-tMCK-MEF2c-polyA-CMV-GFP-

polyA vectors containing codon-optimized mouse transgenes were custom created by SignaGen

Laboratories (Rockville, MD). A truncated muscle creatine kinase (tMCK) promoter was selected

to drive expression of MYOC and MEF2c in adult muscle fibers only. AAV9-CMV-GFP was used

as control. Two-three weeks prior to beginning tumor studies, AAV9 vectors (1x10E11 vg)

dissolved in 20 µL Lact through the skin and into the mid- belly of TA muscles, through both heads of the gastrocnemius muscles to optimally target the soleus, and/or into the intrapleural cavity to target the diaphragm, as described previously (12).

The constitutively active FoxO1 and.FoxO3a plasmids, also known as triple phosphorylation mutants (TM), due to mutations within the three Akt phosphorylation sites which renders them unable to be inactivated by Akt, have been described previously (12). The plasmid encoding for dominant negative (d.n.) MEF2c (amino acids 1 to 105) contains the DNA binding domain, but lacks the transcriptional activation domain, and was kindly provided by Eric Olson

(32). The 3XMEF2-luc plasmid was obtained from Addgene (plasmid #32967), and was deposited

by Ron Prywes. Expression plasmids or empty vector (EV) plus or minus the MEF2 luciferase

reporter dissolved in 50 µL 1x PBS were injected and electroporated into rat soleus muscles,

harvested 4 days later, and processed for either qRT-PCR or measurement of luciferase activity

(12). The Myoc promoter reporter was custom-created by Switch Gear Genomics (An Active Motif

Company, Carlsbad, CA) and comprises of a 521 base pair (bp) fragment of the mouse Myocilin

promoter region, including 432 bp upstream, and 89 bp downstream, of the transcription start site

(TSS) (C57BL/6J chromosome 1, GRCm38.p4 C57BL/6J, 162638718 to 162639238). The Myoc

promoter fragment was cloned into the pLightSwitch reporter vector immediately upstream of the

coding sequence for an optimized Renilla luminescent reporter gene (WT Myoc promoter

reporter). To create the mutated (m)MEF2 Myoc promoter reporter, the conserved MEF2 binding

motif located 169 bp upstream of the TSS (TTCAAAATAGC), was mutated to TTCACGCTAGC, to prevent MEF2 binding. The WT or mMEF2 Myoc promoter reporter plasmids were injected and electroporated into TA muscles of mice (10 µg/25 µL 1xPBS) and muscles harvested 4 days later.

Muscles were homogenized in 1:10 weight/volume passive lysis buffer (Promega) and luciferase activity measured guidelines (SwitchGear Genomics) and a GloMax Single Tube Luminometer (Promega).

Histochemistry and CSA analyses

Processing of skeletal muscle tissue for histological analysis, including H&E staining, was performed as previously described (33). Oil Red O and Trichrome staining was performed by the

Molecular Pathology Core at the College of Medicine, University of Florida, on fresh frozen sections (10 µm) provided by our lab. Determination of transfection efficiency (based on GFP fluorescence) and EBD uptake (using Texas-Red filter) were performed on muscle cross-sections

immediately following cryosectioning using direct fluorescent microscopy. Sections were

visualized and imaged using a Leica DM5000B microscope attached to a digital camera (Leica

Microsystems, Bannockburn IL), with representative images spanning each region of the muscle

captured for analyses. A Leica Application suite, version 3.5.0 software or Image J were used to

trace and measure cross-sectional area (CSA) of muscle fibers, while Image J was used to quantify

extracellular tissue surrounding myofibers in H&E-stained images, both of which have been

described previously (33).

Western Blotting

Homogenization of mouse skeletal muscle tissue, isolation of myofibrillar fractions, western

blotting and quantification using the Odyssey Infrared System (LICOR) were performed as

described previously (34,35). Primary antibodies directions: [anti-Myocilin (MABN866, Sigma Aldrich); anti-alpha-tubulin (T6199, Sigma-

Aldrich); anti-Spectrin- -062, Thermo Scientific)].

RNA isolation, qRT-PCR and human microarray data

RNA isolation, cDNA synthesis and qRT-PCR were performed as described previously (34) using a 7300 Real-time PCR system and TaqMan gene expression primer sets from Applied Biosystems

(Austin, TX), listed in Supplemental Methods. Microarray data from rectus abdominis muscle biopsies from non-cancer control patients and PDAC patients (Series GSE130563) was described previously (10). Additional details can be found in Supplemental Methods. Correlative relationships between specific transcripts of interest and clinical variables in these patients are presented in Supplementary Tables S3 and S4.

Muscle Function

The methods and solutions used for studies of mouse diaphragm muscle isometric function have been described by us previously (35). Mice were anesthetized using 5% isoflurane. Upon reaching a surgical plane of anesthesia the diaphragm was excised and immediately placed in a dissecting chamber containing a Krebs-Hensleit solution equilibrated with 95% O2-5%CO2 gas. A muscle strip, including the tendinous attachment at the central tendon and rib cage was dissected from the mid-costal region. The strip was suspended vertically between two lightweight plexiglass clamps with one end connected to an isometric force transducer (Dual-mode lever system, Aurora

Scientific, Aurora, ON, Canada) within a jacketed tissue bath. The force output was recorded via a computerized data acquisition system (LabView 8.6, National Instruments, Austin, TX). After a

15-min equilibration period, in vitro diaphragm contractile measurements were made. The muscle strip was stimulated along its entire length with platinum wire electrodes (S48 stimulator, Grass

Instruments) to determine the optimum contractile length (Lo). Lo was determined by systematically adjusting the length of the muscle and evoking single twitches. To measure the force frequency response each strip was stimulated supramaximally with 120-V pulses at 15, 30,

60, 100 and 160 Hz while at Lo. The duration of each train was 500 ms to achieve a force plateau.

Contractions were separated by a two-minute recovery period. Diaphragm force production was normalized to specific optimal force. The total muscle cross-sectional area, measured at a right angle to the long axis, was calculated by the following algorithm: total muscle cross-sectional area

(mm2) = [muscle mass/(fiber length x 1.056)], where 1.056 is the density of muscle (in g/cm2).

Fiber length was expressed in centimeters measure at Lo.

Statistical Analyses.

GraphPad Prism statistical software, version 8.0 were used for data analysis. All variables are

presented as mean ± standard error, unless otherwise stated. Mann-Whitney test was used to

compare two groups and Kruskal-Wallis to compare groups of three or more, with comparisons post-hoc test, when appropriate. Two-way ANOVA was used to determine main effects and interactions of two factors, followed by multiple comparison test to determine differences between individual groups when necessary. In the absence of significant interaction

(36), Sidak multiple comparisons post hoc test was used to determine differences within groups for factors showing significant main effects. Univariate and multivariate correlations were performed using Spearman rank-order correlation coefficient. A P-value of <0.05 was considered statistically significant.

RESULTS

Myocilin is decreased in multiple models of cancer cachexia

To determine the time course in which tumor-burden decreases skeletal muscle expression of

Myoc, we harvested TA muscles from sham and C26 tumor-bearing mice at various time points

post-tumor cell inoculation that correspond to different stages of the muscle atrophy process,

which was assessed on days 18 and 23 via MRI imaging (Supplemental Fig. S1A-I). TA muscles

were subsequently processed for qRT-PCR or western blot analyses. The magnitude of Myoc

downregulation was greatest in muscle of tumor-bearing mice at time points prior to the

development of significant muscle atrophy , yet remained decreased at experimental endpoint (day 26) when mice display significant muscle loss (Fig. 1A-C) (12,35). We next determined if the transcriptional downregulation of Myoc in skeletal muscle is a consistent response across other mouse models of cancer cachexia. We thus measured Myoc mRNA at

IACUC-mandated experimental endpoints in skeletal muscle of three different cohorts of PDAC-

PDX mice (and their respective sham groups), in which PDAC tumor fragments from three different patients were either implanted subcutaneously into the flank (flank model) or directly sutured to the mouse pancreas (orthotopic model) and shown to induce cachexia (24,25). We also measured Myoc mRNA at experimental endpoint in skeletal muscle of sham mice and mice inoculated with human L3.6pl pancreatic cancer cells subcutaneously into the flank or orthotopically into the pancreas (23). Compared to their respective sham groups, each of the

PDAC-PDX and L3.6pl tumor-bearing cohorts showed significantly decreased skeletal muscle expression of Myoc (Fig. 1D,E).

Loss of Myocilin causes muscle fiber atrophy, sarcolemmal fragility and impaired muscle

regeneration following injury

Since the muscle phenotype of mice lacking Myoc (Myoc-/- mice) has not been fully characterized,

we first compared the gross morphology of skeletal muscles from adult Myoc-/- mice (Fig. 2A) to

WT mice. Compared to WT mice, muscle mass of the TA was marginally reduced in in Myoc-/-

mice, while gastrocnemius muscle complex (also known as the triceps surae) mass was not

significantly different (Fig. 2B). However, since muscle mass may be a reflection of not only

muscle fiber size, but the relative levels of extracellular connective tissue and water, we further

stained sections from TA muscles with H&E (Fig. 2C) and quantified muscle fiber CSA.

Compared to WT, muscles from Myoc-/- mice displayed visible increases in extracellular space,

and significantly decreased (~18%) muscle fiber CSA (Fig. 2D,E).

Since Myoc has previously been established to stabilize the DGC, which plays a critical

role in stabilizing the muscle fiber membrane, we determined whether mice lacking Myoc show

sarcolemmal fragility and increased susceptibility to damage induced by downhill running. To do

this we injected EBD which is a membrane impermeable dye into cage-restricted mice, or into mice subjected to downhill running. While only faint staining of EBD was observed in myofibers of cage-restricted Myoc-/- mice, downhill running induced significant uptake of EBD into myofibers within the diaphragm and limb muscles of Myoc-/- mice, but not WT mice (Fig. 2F,G).

Some limb muscles of Myoc-/- mice were so damaged following downhill running that ~50% of the total muscle area was positive for EBD, with the majority of myofibers in these areas already degraded and replaced by mononuclear cells. Thus, these data support the notion that Myoc deficiency induces significant sarcolemmal fragility. Importantly, since tumor-bearing mice have significantly reduced levels of Myoc, we further determined whether cachectic C26 tumor-bearing

mice also display sarcolemmal fragility. We found that diaphragm muscles of cage-restricted C26

mice showed clear evidence of myofiber uptake of EBD in the absence of downhill running, which

was exacerbated in response downhill running (Fig. 2H) indicating significant sarcolemmal fragility and damage of myofibers within the diaphragm. In contrast, hindlimb muscles of C26 mice did not show similar evidence of sarcolemmal fragility and damage, which was assessed via both MRI imaging (Supplemental Fig. S2A,B) and EBD uptake assays (Supplemental Fig.

S2C,D).

We also assessed the ability of TA muscles from Myoc-/- mice to regenerate following muscle injury, since muscle regeneration has been linked with muscle wasting in the LLC and C26 models of cancer cachexia (8,9). To do this we injected TA muscles of WT and Myoc-/- mice with cardiotoxin, and harvested muscles for histological analyses at multiple time points post-injury

(Fig.3A). Although the morphology of injured muscles from WT and Myoc-/- mice were largely comparable 2 weeks following CTX injury, at later time points Myoc-/- mice showed reduced CSA of regenerating muscle fibers (Fig.3B,C), coupled with increased extracellular connective tissue

(Fig.3D), indicating an important role for Myoc in the later stages of muscle regeneration.

Since our data indicate that Myoc-/- mice have both increased susceptibility to contraction- induced damage and impaired muscle regeneration, we hypothesized that Myoc deficiency throughout the lifespan could lead to pathological muscle remodeling. Indeed, pathological muscle remodeling is a known consequences of repeated cycles of muscle damage, coupled with impaired regeneration (37). To test this hypothesis we harvested and histologically analyzed TA and diaphragm muscles from 18 month-old WT and Myoc-/- mice using H&E. While reduced muscle quality is expected with the normal aging process, we observed a notable increase in connective tissue deposition within diaphragm muscles of Myoc-/- mice compared to WT mice (Fig. 3E and

Supplemental Fig. S3A). Through additional staining of muscle cross-sections with

Trichrome and Oil Red O, we further confirmed the increased connective tissue to be composed of both fibrotic scar tissue (Supplemental Fig. S3B) and fatty tissue deposition (Supplemental

Fig. S4). TA muscles of Myoc-/- mice also showed evidence of enhanced fibrotic scarring

(Supplemental Fig. S5A,B), but were less affected than the diaphragm which is consistent with findings in models of muscular dystrophy in which the diaphragm is more severely affected than limb muscles (38).

Loss of Myocilin mediates tumor-associated muscle wasting

In order to determine whether the downregulation of Myoc in skeletal muscle in response to tumor- burden plays a causative role in tumor-induced muscle wasting, we injected TA muscles with rAAV9-tMCK-Myoc-GFP (or rAAV9-GFP as control), in which Myoc expression is driven by a muscle-specific (muscle creatine kinase) promoter. Two weeks later, mice were subcutaneously inoculated with C26 tumor cells (or PBS in sham mice), and tissues harvested at experimental endpoint, when mice show significant body wasting (Supplemental Fig. S6A) and muscle wasting

(Fig.4A). Upregulation of MYOC significantly increased muscle mass (normalized to total body mass) in both sham and C26 tumor-bearing mice (Fig.4A), despite the same tumor-burden

(Supplemental Fig. S6B). Successful transduction of muscle with AAV vectors was confirmed via measurement of Myoc via qRT-PCR (Fig. 4B), and through visualization of GFP fluorescence in muscle-cross sections (Fig. 4C). Measurement of muscle fiber CSA showed a significant 38% decrease in muscle fiber CSA in TA muscles of tumor-bearing mice transduced with AAV9-GFP

(Fig. 4D). In contrast, C26 mice transduced with AAV9-tMCK-MYOC-GFP showed only 21% fiber atrophy (vs. Sham-AAV9-GFP), a ~45% sparing of muscle fiber CSA. Despite these

findings, no effects of MYOC rescue were observed on the activation of muscle atrophy biomarkers, Fbxo32/atrogin-1/MAFbx, Trim63/MuRF1 or Fbxo30/MUSA1 (Supplemental Fig.

S6C). Thus, the partial protection against muscle wasting by MYOC gain-of-function likely occurs independently of pathways that induce protein degradation.

We next determined whether skeletal muscles from mice lacking Myoc exhibit exacerbated muscle wasting in response to tumor burden. To test this, we orthotopically injected the pancreas of WT and Myoc-/- null mice with either Panc02 or Panc02-H7 murine pancreatic cancer cells, which are syngeneic with the C57BL/6 background of Myoc-/- mice. As shown in Kaplan Meier survival curves in Supplemental Fig. S6D, there was a trend towards decreased survival in Myoc-

/- mice vs WT mice bearing orthotopic Panc02/Panc02-H7 tumors (P = 0.0877). Indeed, 7/16 tumor-bearing Myoc-/- mice (44%), died or reached IACUC-mandated endpoint (related to deterioration of body condition score) prior to the expected endpoint of 25 days post-surgery, compared to only 2/13 tumor-bearing WT mice (15%). No significant differences were observed in primary tumor mass or in primary tumor-free body mass between WT and Myoc-/- mice

(Supplemental Fig. S6E,F) which could be related to marked metastasis observed in this model.

However, Panc02-H7 tumor burden induced significant loss of gonadal fat mass in both WT and

Myoc-/- mice, consistent with cachexia (Supplemental Fig. S6G). Moreover, despite similar tumor-burden, Myoc-/- mice showed significantly greater loss of muscle mass than WT mice (Fig.

4E). Compared to Sham mice, gastrocnemius muscle mass of Myoc-/- mice decreased by 22.9%

(P<0.001) in response to Panc02-H7 tumor burden, while WT mice showed a 10.8% decrease

(P<0.01). A similar trend was evident in the TA muscle (Fig.4F), but was not significant between genotypes. However, further measurement of muscle fiber CSA in TA muscle cross-sections stained with H&E (Fig. 4G) revealed significant main effects of both Panc02-H7 tumor-burden

and Myoc deficiency (Fig. 4H). Similar to our findings with MYOC overexpression, the absence

of Myoc did not significantly alter the activation of muscle atrophy biomarkers, despite a trend

towards increased levels of Musa1 (Supplemental Fig. S6H-J).

Myocilin is transcriptionally regulated by MEF2c

To gain further insight into the mechanisms whereby Myoc is transcriptionally downregulated in

response to tumor-burden we analyzed the human and mouse Myoc gene promoters. Despite our

previous identification of Myoc as a downstream target of FoxO, we were unable to identify a

consensus FoxO binding motif suggesting FoxO factors likely downregulate Myoc through an indirect mechanism. However, within a conserved regulatory module ~200 bp upstream of the

TSS, we identified a conserved binding motif for MEF2. Of the Mef2 isoforms expressed in skeletal muscle, Mef2c is significantly decreased at the mRNA [(11,12) and Supplementary

Fig.S7A,B] and protein level (39) in skeletal muscle of tumor-bearing mice along with several downstream targets of MEF2c (39), thus also supporting reduced MEF2c-dependent transcription.

We therefore reasoned that Myoc downregulation in response to tumor burden could be mediated, at least in part, through FoxO-dependent inhibition of MEF2. In support of this, we demonstrate here that transfection of muscle with plasmids encoding constitutively active mutants of FoxO1 or

FoxO3a, is sufficient to decrease global MEF2 transcriptional activity (Fig. 5A), and decrease the gene expression of both Myoc and Mef2c (Fig. 5B), the latter of which is also known to be regulated by MEF2 activity (40).

To determine if a reduction in MEF2c activity alone is sufficient to decrease Myoc transcription, we transfected rodent muscles with an empty vector or dominant negative (d.n.)

MEF2c expression plasmid and, 4 days later, measured Myoc mRNA. Compared to empty vector,

d.n.MEF2c repressed Myoc gene expression by ~50%, thus highlighting MEF2c as a key activator of Myoc gene transcription (Fig. 5C). In support of this, mutation of the conserved MEF2 binding site within the Myoc proximal promoter (Fig. 5D) significantly decreased its promoter activity in skeletal muscle, in vivo, as measured via transfection of muscle with luciferase reporter plasmids driven by the Myoc proximal promoter containing either the WT or mutated MEF2 (mMEF2) motif

(Fig. 5E).

MEF2c gain-of-function blocks the cancer-associated downregulation of Myoc and muscle wasting

We next determined whether reduced activity of MEF2c specifically within adult muscle fibers plays a causative role in the C26-induced downregulation of Myoc and mediates muscle loss. To do this, we injected mouse TA muscles with AAV9-GFP or AAV9-tMCK-MEF2c-GFP, and 2 weeks later inoculated mice with C26 cells. Tissues were subsequently harvested at experimental endpoint. Successful targeting of the Mef2c-expressing vector to the TA was confirmed via qRT-

PCR (Fig. 5F). We found that MEF2c gain-of-function in skeletal muscle significantly increased muscle mass across both sham and tumor-bearing groups, which maintained muscle mass in C26 mice to levels comparable to Sham (Fig. 5G). Aligned with this finding, TA muscles transduced with AAV9-tMCK-MEF2c-GFP were significantly protected against C26-induced muscle fiber atrophy (Fig. 5H,I). In support of this, qRT-PCR analyses further demonstrated that maintaining

MEF2c activity was sufficient to deter not only the C26-induced downregulation of Myoc (Fig.

5J,K) but significantly blocked the activation of muscle atrophy biomarkers involved in muscle protein breakdown, atrogin-1/MAFbx and MuRF1/Trim63 (Fig. 5L).

MEF2c gain-of-function prevents muscle wasting and dysfunction in a pre-clinical model of

PDAC-associated cachexia

While the data generated thus far using the C26 model offer novel insight into mechanisms of

cancer-induced muscle wasting, we aimed to determine whether our findings carry over to muscle

wasting associated with pancreatic ductal adenocarcinoma (PDAC), a cancer type that shows high

prevalence of cachexia (1). We therefore conducted additional studies using a murine model of

PDAC-associated cachexia, involving orthotopic implantation of pancreatic tumor cells isolated

from a syngeneic C57BL/6 KRASG12D P53R172H +/+ (KPC) mouse, that has been established to recapitulate key features of PDAC-associated cachexia (19). We found that skeletal muscle expression of Myoc was significantly reduced in KPC mice as early as 8 days post-surgery

(Fig. 6A), and remained repressed until IACUC-mandated experimental endpoint, which was reached ~15-17d post-surgery. The mRNA for Mef2c was significantly reduced by day 12 post- surgery, and remained decreased until study endpoint. To determine if MYOC or MEF2c gain-of- function would provide similar protection against PDAC-associated muscle wasting, as we found in the C26 model, we transduced TA, soleus and diaphragm muscles of mice with AAV9-GFP,

AAV9-tMCK-MYOC or AAV9-tMCK-MEF2c. Three weeks later, mice were inoculated with

KPC cells (or saline for Sham), and tissues harvested on days 13-14 post-surgery, when KPC mice display significant body and muscle wasting (Fig. 6B-D). We found that both MYOC and MEF2c gain-of-function were protective against KPC-induced loss of muscle mass in the TA (Fig. 6C) and solei (Fig. 6D). Since MEF2c acts upstream of MYOC, and is well-established to regulate additional genes critical to muscle structure and function (39,41), we further determined whether maintaining MEF2c activity would also provide protection against KPC-induced dysfunction of the diaphragm, which is implicated in cachexia-associated mortality. We found that the decrease

in muscle specific force observed in the diaphragm of KPC mice transduced with GFP was blunted in muscles transduced with MEF2c across multiple force frequencies (Fig. 6E), with maximum specific force significantly higher in diaphragm muscles of KPC mice transduced with MEF2c

(Fig. 6F).

MYOC is reduced in PDAC patients exhibiting cachexia

We next determined whether our findings are relevant to PDAC patients exhibiting cachexia. To do this we extracted the normalized levels of MYOC mRNA from a recently published microarray analyses that we performed on rectus abdominis muscle biopsies from n = 16 non-cancer control patients and n = 20 PDAC patients (10) (Supplementary Table S1). Using CT images taken at the level of the third lumbar vertebra from these same patients, we subsequently performed CT- based measurements of skeletal muscle index (SMI) and muscle attenuation (MA) using

SliceOmatic software, which are quantitative measures of skeletal muscularity and myosteatosis, respectively (28). From this cohort of patients, CT scans were available from n = 15 control patients and n = 19 PDAC patients. Using previously established sex- and BMI-specific thresholds

(42), patients were further classified as having normal muscularity or muscle depletion (low SMI), and having normal or low MA. We subsequently compared the mRNA levels of MYOC and its upstream regulators FOXO1 and MEF2c, in non-cancer control patients with normal muscularity

( n = 9) to PDAC patients defined as cachectic based on body weight loss of >8%, in combination with muscle depletion and low MA, which are cachexia thresholds previously established to associate with short survival in cancer patients (42). Using these cachexia criteria, n = 9 PDAC patients (47%) were defined as cachectic (Supplementary Table S2). Compared to non-cancer controls, cachectic PDAC patients showed significantly reduced levels of MYOC mRNA (P =

0.0207, Fig. 7A), as well as increased levels of FOXO1 (P = 0.0315, Supplemental Fig. S8A).

MEF2c mRNA was not statistically different between groups (P = 0.1903, Supplemental Fig.

S8B). We also measured MYOC protein levels, but found high variability within the control group, and no statistical difference between groups (Supplemental Fig. S9A-C). We also further examined in PDAC patients alone, whether the mRNA levels of MYOC are associated with the mRNA levels of FOXO1 and MEF2c, and with clinical parameters including sex, age, cancer- stage, BMI, % BW loss, SMI, MA and survival time post-surgery using a multivariable correlation matrix (Supplemental Table S3). A significant positive correlation was identified between skeletal muscle MYOC mRNA and survival time post-surgery (P = 0.0008, r = 0.700, Fig. 7B). In support of this, stratification of PDAC patients based on survival time of >1 year vs <1 year post- surgery revealed significantly lower levels of MYOC in patients surviving <1 year post-surgery (P

= 0.005, Fig. 7C). We also identified a positive correlation between the mRNA levels of MYOC and MEF2c (r = 0.4754, P = 0.0397), and a negative correlation between the mRNA levels of

FOXO1 and MEF2c (r = -0.4947, P = 0.0266). These data therefore highlight the clinical relevance of our findings in mice, and collectively implicate the dysregulation of a FoxO1-MEF2c-Myoc axis in skeletal muscle of cancer patients that is associated with cachexia.

DISCUSSION

In the current study we identify Myoc as a downstream target gene repressed by the FoxO transcription factors whose downregulation precedes and contributes to muscle loss in two different models of cancer cachexia, and which is mediated through reduced activity of MEF2c.

We further establish that MEF2c gain-of-function not only prevents Myoc downregulation in response to tumor burden, but protects against muscle wasting and weakness in a pre-clinical model of PDAC-associated cachexia. We also demonstrate significant clinical relevance to these findings, showing that MYOC is significantly downregulated in skeletal muscle biopsies from cachectic PDAC patients who also show increased levels of FoxO1, and correlates with the levels of MEF2c.

Through additional characterization of mice lacking Myoc we demonstrate the biological significance of Myoc in skeletal muscle. In this regard, we show that Myoc deficiency induces muscle fiber atrophy, increased susceptibility of the sarcolemma to damage and impaired muscle regeneration, which are phenotypes implicated in muscle loss associated with cancer, at least in the C26 and LLC models of cachexia (8,9). Moreover, in aged mice lacking Myoc, we also observed a significant decline in skeletal muscle quality, characterized by increased fatty deposition and fibrotic scar tissue, which are known consequences of chronic damage and impaired regeneration (37). Moreover, although abundant deposition of fat and fibrotic tissue are not phenotypes typically associated with skeletal muscle wasting in rodent models of cancer cachexia, recent work from our lab identified fibrotic remodeling as a feature of cachexia in the orthotopic

PDAC patient-derived xenograft (PDAC-PDX) model, which was particularly evident in the diaphragm (43). Moreover, low muscle attenuation (myosteatosis, i.e. fatty muscle) identified via

CT imaging is a well-established feature of cachexia in cancer patients (28) which also indicates

reduced muscle quality as a feature of human cancer cachexia. In support of this, increased fat and

fibrotic tissue have also been observed histologically in skeletal muscle biopsies from cachectic

PDAC patients (10). However, it is important to note that the reduced muscle quality observed in

muscles of aged mice lacking Myoc could be related to the absence of Myoc in not only muscle,

but other cell types, since Myoc deletion was not restricted to skeletal muscle. In contrast, in our

cancer cachexia studies, MYOC gain-of-function experiments were accomplished using AAV9

and a muscle-specific promoter to selectively upregulate MYOC within muscle fibers only.

Through the studies herein we further identify MEF2c as a key upstream transcription

factor that directly activates Myoc transcription under baseline condition, whose reduced function

mediates Myoc downregulation in response to tumor burden. In support of this, previous work has

established that the protein expression of MEF2c, and several bonafide downstream target genes

of MEF2 are also reduced in skeletal muscle of C26 tumor-bearing hosts (39) which together support reduced MEF2c transcriptional activity. We extend the significance of these findings by demonstrating that MEF2c gain-of-function not only prevents the cancer-associated downregulation of Myoc, but also significantly blocks cancer-induced muscle wasting and dysfunction. Moreover, the protection against muscle loss conferred by MEF2c was also associated with a significant inhibition in the cancer-induced activation of atrogin-1 and MuRF1, thus further linking MEF2c dysfunction with the activation of transcriptional pathways which promote muscle proteolysis.

MEF2c is a well-characterized transcription factor involved in multiple steps of skeletal muscle development, including myoblast differentiation and fusion, and various processes which regulate post-natal skeletal muscle homeostasis and plasticity (44-46). In this regard, skeletal muscle expression of MEF2c is necessary for the maintenance of post-natal muscle fiber integrity

(41), normal muscle fiber-type specification (47), as well as normal growth and glucose

metabolism (48). MEF2c also promotes muscle regeneration and re-growth following injury (49),

and when constitutively active, can promote muscle hypertrophy (50). Thus, MEF2c gain-of-

function in skeletal muscle may protect against cancer-induced muscle wasting through its

regulation of a wide range of target genes involved in these processes, which warrants further

investigation.

The mechanisms whereby MEF2c-dependent transcription is reduced in response to tumor

burden are not clear. However, our data indicate that FoxO activation may be involved, as both

FoxO1 and FoxO3a are sufficient to decrease Mef2c mRNA and global MEF2 transcriptional

activity and blocking FoxO activation prevents the cancer-induced downregulation of several

MEF2c target genes, including Myoc, and Mef2c (12). However, MEF2 activity is well-established to be regulated through neural activity (51) and calcium-dependent signaling pathways such as calcium/calmodulin-dependent kinase CaMK) signaling (52) which activate MEF2 through preventing its association with histone deacetylase (HDAC) proteins, which are potent repressors of MEF2-dependent transcription. Since HDAC inhibitors have shown significant efficacy in blocking cancer-induced muscle wasting in the C26 and LLC models (53), exploring the link between HDACs and MEF2 in the context of cancer cachexia warrants further study.

Limitations of this study include the assumptions that mechanisms previously identified using the well-characterized C26 and LLC models of cancer cachexia carry over to more clinically relevant models of cancer cachexia, such as the orthotopic KPC model used herein. In this regard, additional studies are needed to determine whether FoxO activation, DGC disruptions and impaired regeneration similarly contribute to muscle wasting in the KPC model and other models of PDAC-associated cachexia. The small sample size of PDAC patients and non-cancer controls

used for analyses herein is also a significant limitation of the study, as is the possibility of sampling

bias thus highlighting the need to validate our findings using a larger cohort of cancer patients.

In summary, our data identify MEF2c-dependent transcription of Myoc as a novel transcriptional pathway repressed by FoxO that mediates muscle loss in two different models of cancer cachexia, and which is similarly dysregulated in cachectic cancer patients. These findings thus provide novel insight into mechanisms of cancer-associated muscle wasting and dysfunction that could be further explored as therapeutic targets.

ACKNOWLEDGEMENTS

This work was supported by the National Institute of Arthritis, Musculoskeletal and Skin Diseases

(R01AR060209 to ARJ); the National Cancer Institute (R21CA194118 to ARJ); the UF Health

Cancer Center (Bridge Funding to ARJ); the UF Clinical and Translational Science Institute

(CTSI) (Pilot Award to ARJ, CTSI Pilot Award to SMJ) [the UF CTSI is supported by the National

Center For Advancing Translational Sciences of the National Institutes of Health

(UL1TR001427)]; and the V Foundation for Cancer Research (V2015-021 to JGT). D.N. is supported by a Swiss National Science Foundation grant (P4000PM_180814/1). RLN is supported by a National Institute of Child Health and Human Development Grant (T32-HD-043730), and

ACD is supported by a National Heart, Lung and Blood Institute Grant (T32HL134621).

REFERENCES

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FIGURE LEGENDS

Figure 1. Myoc is transcriptionally downregulated in skeletal muscle in multiple experimental models of cancer cachexia. A-C) Skeletal muscles harvested from C26 tumor- bearing mice were assessed for Myoc mRNA (A, TA muscles) and MYOC protein (B, gastrocnemius muscles) at various time points throughout tumor growth and the progression of muscle wasting (C, and Supplemental Fig. S1). Data are representative of n = 3-6 mice/group. D, E) The relative expression of Myoc mRNA was assessed in TA muscles in multiple experimental models of cancer cachexia, including in mice bearing human patient-derived xenograft (PDX) tumors implanted subcutaneously into the flank (PDX1-F, PDX2-F) or orthotopically into the pancreas (PDX-O) (D), and in mice inoculated with human L3.6pl tumor cells subcutaneously into the flank (L3.6pl-F) or orthotopically into the pancreas (L3.6pl-O) (E). Data are expressed as mean ± SEM, normalized to their respective sham group. Data are representative of n = 4-6 mice/group. For visualization purposes only, Sham groups for each cohort of PDX mice were combined, as were the Sham groups for the L3.6pl tumor-bearing groups. *P<0.05, **P<0.01, ***P<0.001, ****P<0.0001 vs Sham.

Figure 2. Myoc deficiency causes muscle fiber atrophy and sarcolemmal fragility. A) qRT- PCR validation of Myoc knockout, showing negligible detection of Myoc mRNA in TA muscles of Myoc-/- mice vs WT mice. B) Muscle mass of the TA and gastrocnemius muscle complex of 12 week-old WT and Myoc-/- mice. C,D) Average CSA (C) and frequency distribution (D) of TA muscle fibers in WT and Myoc-/- mice measured from H&E-stained muscle cross-sections (E). Data in A-D represent mean ± SEM mean, n = 4-6 mice/group. Scale bars = 50 µM. F) Representative cross-sections from the plantaris muscle and the diaphragm of WT and Myoc-/-

(EBD, red), a membrane-impermeable fluorescent dye that can only enter the cytosol of fibers following significant tearing of the sarcolemma. Muscle damage was further confirmed via H&E staining (representative of n = 4 mice/group). Scale bars = 400 µm (plantaris) and 100 µm (diaphragm). G) The average number of EBD+ fibers per 20x field view in diaphragm muscle cross-sections from WT and Myoc-/- mice 1 day following downhill running (n = 4 mice/group). White arrows show EBD positive muscle fibers matching corresponding hypereosinophilic

(damaged) fibers in serial sections stained with H&E. Scale bar = 100 µm. H) Representative cross-sections from the diaphragm of C26 mice subjected to normal cage activity or downhill treadmill running, showing myofiber uptake of EBD that is exacerbated in response to downhill running. Data are representative of n = 4 mice/group. *P<0.05 vs WT.

Figure 3. Myoc deficiency impairs skeletal muscle regeneration and leads to reduced muscle quality in aged mice. A) Representative H&E-stained cross-sections from TA muscles harvested from WT and Myoc-/- mice 2, 4 and 6 weeks post CTX-injury (n = 4-5 mice/group). Scale bars = 100µm. The average cross-sectional area (B) and frequency distribution (C) of regenerating myofibers with centralized nuclei was calculated in muscles of WT and Myoc-/- mice 4 weeks post CTX-injury (~2000 fibers measured/group). D) The amount of non-eosin stained area surrounding muscle fibers (extracellular tissue/space) was calculated in regenerating muscles of WT and Myoc- /- mice 6 weeks post-injury (*P<0.05; n = 5 mice/group). E-G) Representative cross-sections from diaphragm muscles of aged (18 month-old) WT and Myoc-/- mice, stained with H&E (E Trichrome (collagen stains blue) (F), and Oil Red O (lipid stains orange) (G). Scale bars = 250 µm (top panel) and 100 µm (bottom panel).

Figure 4. Loss of MYOC mediates tumor-associated skeletal muscle wasting. A-D) TA muscles from mice were transduced with AAV9-GFP or AAV9-tMCK-MYOC-GFP and 2 weeks later subcutaneously inoculated with C26 tumor cells or 1xPBS (Sham). C26-induced changes in TA muscle mass normalized to total body mass (A), and Myoc mRNA as measured via qRT-PCR (B). C) Representative cross-sections from TA muscles of C26 mice showing successful transduction of AAV vectors into myofibers, as visualized via direct GFP fluorescence. Scale bars = 50 µm. D) TA muscle fiber CSA measured at experimental endpoint (*P<0.05 vs. Sham-AAV9- GFP; N = 4 mice/group, except Sham-MYOC-GFP, N = 3 mice/group). E-H) Panc02-H7 tumor cells were orthotopically injected into the pancreas of WT and Myoc-/- mice and tissues harvested at experimental endpoint. Changes in gastrocnemius muscle complex mass (*P<0.05 vs WT Sham; ****P<0.0001 vs. Myoc-/- Sham) (E) and TA muscle mass (*P<0.05 vs WT Sham; ****P<0.0001 vs. Myoc-/- Sham) (F). Data are normalized to sex-matched WT Sham mice (n = 6-8 mice/group). Representative TA muscle cross-sections stained with H&E (G) and average TA muscle fiber CSA (H) of WT and Myoc-/- mice in response to Panc02-H7 tumor-bearing (**P<0.01 vs WT Sham;

*P<0.05 vs. Myoc-/- Sham; n = 5 mice/group). Scale bars = 50 µm. *P<0.05, **P<0.01, ***P<0.001, ****P<0.0001

Figure 5. MEF2c gain-of-function blocks Myoc downregulation and cancer-associated muscle wasting. A,B) Luciferase activity derived from a 3xMEF2-dependent luciferase reporter plasmid (A), and qRT-PCR analysis of Myoc and Mef2c mRNA (B) from rat soleus muscles transfected with expression plasmids encoding constitutively active FoxO1 or FoxO3a, which harbor mutations in the 3 inhibitory Akt phosphorylation sites, and are thus known as triple mutant (TM) forms of FoxO1 or FoxO3. C) Analysis of the mouse and human Myoc proximal promoter regions identified a conserved MEF2 binding motif located within ~200bp upstream of the transcription start site. D) qRT-PCR analysis of Myoc mRNA in rat solei injected with either empty vector (EV) or dominant negative (d.n.) MEF2c expression plasmid (*P<0.05; n = 5-6 rats/group). E) Plasmid-based luciferase activity driven by an ~0.5kb fragment of the Myoc proximal promoter (WT), compared to a mutated version (mMEF2), in which a canonical MEF2 binding motif was mutated to prevent MEF2 binding, 4 days following plasmid transfection into mouse TA muscles, in vivo (Paired t-test; ***P<0.001). F) qRT-PCR analysis of Mef2c mRNA in TA muscles of sham and C26 tumor-bearing mice transduced with AAV9-GFP or AAV9-tMCK-MEF2c-GFP confirming successful upregulation of MEF2c (n = 3 mice/group). G-I) Muscle mass normalized to total body mass (*P<0.05 vs. Sham AAV9-GFP) (G), representative muscle cross-sections (H), and muscle fiber CSA (***P<0.05 vs. Sham AAV9-GFP) (I) from TA muscles harvested at experimental endpoint from Sham and cachectic C26 tumor-bearing mice transduced with AAV9- GFP or AAV9-tMCK-MEF2c-GFP. Data are representative of n = 6-8 mice/group, except Sham AAV9-tMCK-MEF2c-GFP group, which represents N = 3 mice. J) qRT-PCR analysis of Myoc mRNA (K) and subsequent linear regression analysis with Mef2c mRNA showing a significant correlation between the mRNA levels of Mef2c and Myoc (r2=0.68, P<0.001). L) qRT-PCR analysis of atrophy-related biomarkers atrogin-1/Fbxo32, MuRF1/Trim63 and Musa1/Fbxo30 mRNA (*P<0.05 vs C26 AAV9-GFP group). Data in J-L represent N = 3 mice/group. *P<0.05, **P<0.01, ***P<0.001, ****P<0.0001.

Figure 6. MEF2c gain-of-function protects against muscle wasting and dysfunction in a murine model of PDAC-associated cachexia. A) qRT-PCR analysis of Myoc and Mef2c in TA

muscles harvested from sham mice, and mice bearing orthotopic KPC tumors on days 6, 8, 10, 12 and 14 following tumor inoculation surgeries, and at IACUC-mandated endpoint (END), which was reached ~15-17 days post-surgery. B-F) Mice received intramuscular and/or intrapleural injections of AAV9-GFP, AAV9-tMCK-MYOC or AAV9-tMCK-MEF2c vectors, and 3 weeks later received an orthotopic injection of KPC cells (or saline, Sham) into the pancreas. Tumor-free body mass (B), TA muscle mass (C), and soleus muscle mass (D) on day 13-14 post-surgery from Sham mice (transduced with AAV9-GFP) and KPC mice (transduced with AAV9-GFP, AAV9- tMCK-MYOC or AAV9-tMCK-MEF2c). Specific force-frequency relationship (E) and maximum specific force (F) in diaphragm strips from Sham mice transduced with AAV9-GFP, and KPC mice transduced with either AAV9-GFP or AAV9-tMCK-MEF2c. All data are representative of n = 3-5 mice/group, *P<0.05, ***P<0.001 (vs. Sham unless otherwise indicated).

Figure 7. MYOC is reduced in rectus abdominis muscle biopsies from PDAC patients exhibiting cachexia, and is associated with reduced survival. A) Normalized levels of MYOC mRNA in skeletal muscle biopsies from non-cancer controls with normal muscularity, compared to cachectic PDAC patients. Cachexia was defined based on body-weight loss of >8%, in combination with CT-defined measurements of muscle depletion and low muscle attenuation (MA) cachexia thresholds which have been identified previously in cancer patients to associate with short survival (42). Data are reported as mean ± SEM, (*P<0.05). One PDAC patient was identified as an outlier (ROUT Q = 1%), and removed from analyses. B) Spearman correlation was performed between the skeletal muscle levels of MYOC mRNA and survival time (days) of PDAC patients following tumor resection surgery. C) The relative mRNA levels of MYOC in PDAC patients stratified based on survival time post tumor resection surgery (>1 year vs. < 1 year) (**P<0.01).

MEF2c-dependent downregulation of Myocilin mediates cancer-induced muscle wasting and associates with cachexia in cancer patients

Sarah M Judge, Michael R Deyhle, Daria Neyroud, et al.

Cancer Res Published OnlineFirst March 4, 2020.

Updated version Access the most recent version of this article at: doi:10.1158/0008-5472.CAN-19-1558

Supplementary Access the most recent supplemental material at: Material http://cancerres.aacrjournals.org/content/suppl/2020/03/04/0008-5472.CAN-19-1558.DC1

Author Author manuscripts have been peer reviewed and accepted for publication but have not yet been Manuscript edited.

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