Effect of Exercise on Statin-Induced Skeletal Muscle Myopathy

Senior Thesis

Alay S. Parikh

The School of Molecular and Cellular Biology

University of Illinois at Urbana-Champaign

Research Advisor:

Associate Professor Marni D. Boppart, ScD.

Department of Kinesiology and Community Health

University of Illinois at Urbana-Champaign

1

Abstract

HMG-CoA reductase inhibitors, or statins, significantly decrease

hypercholesterolemia and protect against cardiovascular disease. Statins directly inhibit

the action of the HMG-CoA reductase enzyme in the cholesterol synthesis pathway. One

of the most common side effects of statin treatment is skeletal muscle myopathy, a

condition that may be exacerbated by exercise. Our recent results suggest that short-

term exercise does not aggravate statin-induced myopathy as assessed by force

production, but may initiate myofiber damage and atrophy in a manner that can result in

exacerbated myopathy with long-term statin administration. The purpose of this study was

to directly assess myofiber damage and atrophy in hypercholesterolemic mice (ApoE-/-)

treated with statins and exercise. Male, ApoE-/- mice were provided access to a running

wheel for two weeks, then continued exercise for an additional two weeks while receiving

daily injections of simvastatin or saline treatment (accustomed exercise). A second

exercise group received access to a running week for two weeks concomitant with

simvastatin or saline treatment (no prior training, novel exercise). A control group received

simvastatin or saline treatment for two weeks without access to a running wheel

(sedentary). There was no evidence of overt fiber damage as a result of statin

administration in the absence or presence of exercise. However, Type 2B fiber size was

significantly reduced in the accustomed exercise group that received statin therapy. The

results from this study suggest that statin treatment may stimulate early myofiber

degradation of Type 2 fibers when combined with an exercise training program.

2

Introduction

Cardiovascular disease (CVD) is the leading cause of death for both men and

women in the United States. One of the most common and effective preventative

treatments to combat CVD is statin therapy. Statins work by inhibiting HMG-CoA

reductase, the rate-limiting enzyme in the cholesterol biogenesis pathway. High

cholesterol levels in the blood (hypercholesterolemia) increase the risk of CVD via an

increase in atherosclerosis within the circulatory system. As such, statins are one of the

most widely prescribed pharmaceutical agents administered to prevent the development

of CVD. However, a relatively common side effect of statin usage is the occurrence of

skeletal muscle myopathy (Sinzinger 2002; Ucar 2012). The mechanism behind the

relationship between statins and myopathy remains poorly understood, and elucidating

the underlying molecular mechanisms is of prime importance to reduce the incidence of

statin-induced myopathy.

Different mechanisms have been proposed to explain the mechanistic basis for

statin-induced myopathy. The primary theory supported by literature is an increase in

mitochondrial dysfunction corresponding with a depletion of Coenzyme Q10 in the muscle

(Duncan 2008, Vaklavas 2009). This mitochondrial co-factor serves as an electron carrier

within the electron transport chain. Reductions in Coenzyme Q10 can result in an

abnormal accumulation of reactive oxygen species (ROS) within the mitochondria, which

can ultimately induce cellular apoptosis, DNA damage, and enzyme inactivation (Slimen

2014). Coenzyme Q10 is generated downstream of mevalonate (a major product in the

early stages of cholesterol biogenesis). Thus, statin usage inhibits the tissue’s normal

ability to synthesize Coenzyme Q10, which can result in enhanced mitochondrial

3

dysfunction and altered cellular respiration. This dysfunction could be responsible for the

skeletal muscle myopathy and myalgia experienced by statin users.

Cholesterol is vital to the stability and structure of the skeletal muscle membrane,

or sarcolemma. Statin-induced disruptions to cholesterol synthesis may significantly

impair muscle membrane flexibility, allowing for damage as a result of contraction and

mechanical strain (Parker 2012). Thus, a reduction in cholesterol represents an

alternative explanation for statin-induced myopathy.

A main preventative measure recommended for individuals at risk for CVD is to

undertake an aerobic exercise training program (American Heart Association 2013).

Aerobic exercise training has repeatedly been demonstrated to improve cardiovascular

function as well as reduce high blood pressure and the risk for CVD (Gielen 2015).

Despite the fact that exercise is an important component of the prescription for CVD,

studies suggest that exercise in combination with statin therapy may exacerbate

myopathy. Parker et al. (2012) and others have demonstrated that exercise in

combination with statin therapy can increase the prevalence of muscle pain and reduce

engagement in physical activity. Therefore, further studies are necessary to provide a

better understanding of the biological basis for this condition.

Our lab recently conducted a study that attempted to address the mechanisms

that underlie statin-induced myopathy and the impact of exercise on this condition

(Boppart lab, unpublished results, manuscript in progress). Hypercholesterolemic mice

(ApoE-/-) received either statin therapy or saline then were subjected to statin treatment

after two weeks of running wheel activity (accustomed exercise group, total of 4 weeks of

exercise), statin treatment concurrent with exposure to running wheel activity (novel

4

exercise group, 2 weeks of exercise), or remained sedentary throughout the treatment.

Statin treatment resulted in significant myopathy as assessed by total running wheel

activity, hindlimb grip strength, and maximal isometric force. Exercise, either accustomed

or novel, did not provoke further deficits in activity or force. However, gene expression of

an important ubiquitin ligase, atrogin-1, was significantly elevated in the exercise groups

in combination with statin administration. Systemic inflammation, as assessed by serum

amyloid A, was also significantly elevated in the novel exercise group in combination with

statins. Therefore, these results suggest the ability for exercise to stimulate the

degradation of myofibrillar protein during statin administration.

The purpose of this study was to extend the results of our primary investigation

and determine the extent to which exercise can increase myofiber damage and atrophy

in the presence of statin therapy. We hypothesized, based on previous results, that

exercise would provoke a significant increase in both myofiber damage and atrophy, and

that these early events may contribute to exacerbation of myopathy observed with long-

term statin administration.

Methods

Study Design. 8 week-old male ApoE knockout mice (ApoE-/-) (Jackson Laboratory, Bar

Harbor, ME) were randomized to six groups (n=10/group). Mice were first assigned to

one of three groups: no exercise, voluntary novel exercise (initiation of exercise at 2

weeks, concomitant to initiation of statin or placebo), or voluntary accustomed exercise

(exercise starting 2 weeks prior to statin or placebo administration and during treatment).

Exercise was administered through the use of a running wheel. Mice were injected with

5

either simvastatin (20 mg/kg/day) or an equivalent volume saline. Groups are designated

as Sedentary+Saline (n=3), Sedentary+Statin (n=3-4), Novel+Saline (n=3), Novel+Statin

(n=3-4), Accustomed+Saline (n=3), and Accustomed+Statin (n=2). Due to the original

samples being used for the initial study, the number of samples available for this study

was limited.

Immunofluorescence. Gastrocnemius-soleus muscle complexes previously frozen in

isopentane were divided at the midline along the axial plane, and the distal half was

embedded in OCT (Tissue-Tek; Fischer Scientific). Three transverse cryosections per

sample (10μm non-serial sections, each separated by a minimum of 40 μm) were cut for

each histological assessment using a CM3050S cryostat (Lecia, Wezlar, Germany).

Sections were placed on microscope slides (Superfrost; Fischer Scientific, Hanover Park,

IL) and stored at -80°C before staining.

To assess skeletal muscle damage, sections were stained with anti-IgG antibodies

and dystrophin to distinguish individual muscle fibers. The frozen tissue sections (10μm)

were fixed in ice cold acetone and blocked with AffiniPure Fab Fragments Goat Anti-

Mouse IgG (H+L) (Jackson Immunoresearch Cat #115-007-003) (1:20 dilution) in 5%

bovine serum albumin (BSA) solution for 1 hour at room temperature. Sections were then

incubated with a FITC-conjugated mouse anti-IgG (Vector F1200) (1:100 dilution) and

rabbit anti-mouse dystrophin (Abcam ab15277) (1:100 dilution) in 1% BSA solutions

overnight at 4°C. This was followed by a secondary antibody incubation with AlexaFluor

633 goat anti-rabbit (Invitrogen) (1:100 dilution) in 1% BSA solution for 1 hour at room

temperature. Sections were then briefly incubated with 4’,6-diamidino-2-phenylindole

(DAPI) (1:20000), to stain myonuclei. Sections were washed with 1% BSA for 5 minutes

6

multiple times between each of the incubation periods to remove excess antibodies.

Vectashield (Vector Laboratories) was applied to the samples and slides were sealed with

a coverslip and nail polish. Stained slides were stored at 4°C until imaging.

Myofiber typing stains used antibodies specific for four types of Myosin Heavy

Chain (MHC) isomers: MHC 1, 2A, 2B, 2X. Stains were prepared to detect two isomers

at a time: MHC 2B and MHC 2A were paired together and MHC 2X and MHC 1 were

paired together. All samples were first fixed in ice-cold acetone and then blocked in a

0.5% BSA and 0.5% Triton-X solution containing AffiniPure Fab Fragments Goat Anti-

Mouse IgG (H+L) (Jackson Immunoresearch Cat #115-007-003) (1:10 dilution) for 1 hour

at room temperature. 1x PBS was used to perform multiple washes between incubations.

The MHC 2B and MHC 2A stain consisted of a primary antibody incubation with a

mouse IgM anti-type 2b MHC antibody (BF-F3 concentrate, Developmental Studies

Hybridoma Bank, University of Iowa) (1:50 dilution), mouse IgG1 anti-type 2a MHC

antibody (Sc-71 supernatant, Developmental Studies Hybridoma Bank, University of

Iowa) (1:50 dilution), and rabbit anti-mouse dystrophin (Abcam ab15277) (1:100 dilution).

The antibodies were prepared in a solution of 1x PBS-containing 0.5% BSA and 0.5%

Triton-X and incubated for 1 hour at room temperature. Secondary antibody incubation

conditions included AMCA-conjugated anti-mouse IgM µ-chain specific (Jackson

Immunoresearch Cat #115-155-075) (1:100 dilution), Alexa 488-conjugated anti-mouse

IgG subclass 1 (Jackson Immunoresearch Cat #115-545-205) (1:100 dilution), and Alexa

Fluor 633 goat anti-rabbit (Invitrogen) (1:200 dilution). The antibodies were prepared in a

solution of 1x PBS-containing 0.5% BSA and 0.5% Triton-X and incubated for 1 hour at

room temperature.

7

The MHC 2X and MHC 1 stains consisted of a primary antibody incubation of

mouse IgG2b anti-type 1 MHC antibody (BA-D5 supernatant, Developmental Studies

Hybridoma Bank, University of Iowa) (1:20 dilution), mouse IgM anti-type 2x MHC

antibody (6H1 supernatant, Developmental Studies Hybridoma Bank, University of Iowa)

(1:20 dilution), and rabbit anti-mouse dystrophin (Abcam ab15277) (1:100 dilution). The

antibodies were prepared in a solution of 1x PBS-containing 0.5% BSA and 0.5% Triton-

X and incubated for 1 hour at room temperature. Secondary antibody incubation

conditions included Alexa 488 anti-mouse IgM µ-chain specific (Jackson

Immunoresearch Cat #115-155-075) (1:100 dilution), Alexa 350-conjugated anti-mouse

IgG2b (Invitrogen Cat #A21140) (1:100 dilution), and Alexa Fluor 633 goat anti-rabbit

(Invitrogen) (1:200 dilution). The antibodies were prepared in a solution of 1x PBS-

containing 0.5% BSA and 0.5% Triton-X and incubated for 1 hour at room temperature.

Vectashield was applied to the samples and slides were sealed with a coverslip and nail

polish. Stained slides were stored at 4°C until imaging.

Image Capture and Analysis. All stained tissue samples were visualized using the 10X

objective on an upright inverted fluorescent microscope (Zeiss, Thornwood, NY) and an

excitation source (X-Cite 120, EXFO, Ontario, Canada) with the appropriate filters. Image

capture was performed through the Zeiss AxioCam digital camera and Axiovision capture

software (Zeiss, Thornwood, NY).

For the identification of IgG+ muscle fibers and centrally located nuclei, entire

sections were captured in multiple images at 10X magnification using the upright inverted

fluorescent microscope. One to three sections per animal sample were analyzed

depending on the quality of the individual section and the quality of the stain. Images were

8

captured on multiple color channels that were complementary to the specific stain and

then analyzed as a singular image. The number of total IgG+ muscle fibers were counted

as a factor of the total area of the sections using Adobe Photoshop (Adobe Photoshop

CC 2015). Centrally located nuclei and total number of fibers were counted using the Cell

Counter function in the ImageJ software (NIH, Bethesda, Maryland).

For the evaluation of the muscle fiber-typing stain, three to five multichannel

images were captured of each sample using the appropriate filter, depending on the

quality of the section and quality of the stain. They were taken using the 10X objective on

the upright inverted fluorescent microscope. The area of approximately 150 individual

fibers of a specific fiber type was calculated using Adobe Photoshop to provide the total

cross sectional area (CSA) of that particular fiber type.

Statistical Analysis. All values are presented as mean ± SEM. Two-way ANOVA was

used to test significant differences between the separate exercise groups and statin

treatment (statin*exercise interaction effects) for all histological measures. One-way

ANOVA was used to test differences between groups within each fiber type distribution

range. Least significant difference (LSD) post-hoc analysis was performed to examine

group differences. All statistical analyses were performed using SPSS Ver. 22 (IBM,

Chicago, IL). Differences were considered significant at p≤0.05.

9

Results

Assessment of Myofiber Damage. Total IgG+ fibers per mm2 of muscle tissue were not

significantly different between groups (Figure 1A). The percentage of CLN+ fibers

between treatment groups was not significantly different (Figure 1B).

Assessment of Myofiber Atrophy. Total and fiber type-specific changes in mean CSA

are reported. The percentage of the different fiber types (Type 1, 2A, 2X, 2B) was also

evaluated to determine if statin administration or the combination of statins with exercise

modified fiber composition. Type 1 myofibers were not readily observed within the stained

muscle sections, and thus, not able to be analyzed. The average CSA of all myofibers

was not significantly different between groups (Figure 2A). The mean CSA of individual

Type 2 fiber types was also not different among the groups (Figures 2B-D). The Type 2

fiber size distribution was plotted to assess atrophy, or the percentage of small fibers in

each group. For Type 2A fibers of sizes between 501-1000 µm2, there was a significant

interaction effect (statin x exercise, p=0.02), with the Sedentary+Statin group possessing

a significantly lower proportion of fibers within this range compared to all other groups

(Figure 3A). There were no significant main or interaction effects for the Type 2X fiber

sizes (Figure 3B), but for Type 2B fibers (Figure 3C) a significant statin x exercise

interaction effect was detected for fiber sizes ranging from 1001-1500 µm2 and 2501-3000

µm2 (p=0.04 for both). Within the 1001-1500 µm2 range the Accustomed+Statin group

had a significantly larger proportion of fibers compared to Sedentary+Statin,

Novel+Saline, and Accustomed+Saline. Interestingly, a significant decrease in the

percentage of fibers ranging greater than 3000 µm2 (compared to Sedentary+Saline,

10

Accustomed+Saline) was noted for Type 2B fibers in the accustomed exercise group

receiving statin treatment (p < 0.05) (Figure 3C).

The percentage of Type 2X and 2B fibers revealed a trend toward an interaction

effect of statin and exercise (Type 2X, p=0.07; Type 2B, p=0.06) (Figure 4B-C), but no

significant main effects were observed. The percentage of Type 2A fibers showed no

significant differences among any of the groups (Figure 4A).

Discussion

The purpose of this study was to determine the extent exercise could increase

myofiber damage and atrophy when combined with statin therapy. Upon completion of

the study, we found that there were no significant increases in global myofiber damage,

nor changes to myofiber repair and regeneration. We also examined various indices of

myofiber hypertrophy, including mean CSA, mean fiber type-specific CSA, myofiber size

distribution, and the percentage of Type 2 fibers present. Mean CSA was not altered with

exercise or statin treatment, but Type 2B fiber size was reduced in hypercholesterolemic

mice who underwent exercise prior to statin administration. Overall, the findings from this

study suggest that exercise training may increase skeletal myofiber degradation and

increase susceptibility to myopathy with long-term statin use in hypercholesterolemic

individuals.

Statin-induced skeletal myopathy is theorized to stem from mitochondrial

dysfunction and subsequent impaired energy generation and an accumulation of ROS.

The specific impact on skeletal muscle structure and morphology is less known. In order

to determine the impact of exercise and statins on myofiber sarcolemmic integrity, IgG

11

infiltration into myofibers was assessed. Our findings found no significant difference

between groups (Figure 1A); therefore, sarcolemmic disruption of the myofiber likely does

not contribute to the significant decreases in strength observed in the original study

(Boppart lab, unpublished data, manuscript in progress). In order to corroborate these

findings, the quantity of centrally located nuclei (CLN) per 100 myofibers was evaluated

(Figure 1B) in order to assess the cell’s ability to regenerate and repair damaged muscle

myofibers. Dystrophic muscle often displays CLN in response to rounds of degeneration

and regeneration of muscle fibers (Rahimov 2013) so an increased amount of CLN in a

given area will indicate the myofibers regenerative state (Matsumoto 2007). Similar to the

IgG results, CLN per muscle fiber was not significantly different among groups suggesting

a lack of regeneration and repair, which agrees with the lack of fiber damage observed.

This absence of myofiber damage and regeneration provides evidence that statin-

associated myopathy occurs under a separate mechanism than physical myofiber

structural damage, which is unaffected by short-term endurance exercise.

As myofiber damage was unlikely to be responsible for the strength changes

observed with exercise and statin treatment, we hypothesized that alterations to CSA and

fiber type size would contribute to the strength losses previously observed. Werning et al.

(1989) demonstrated that running exercise results in an increase in CSA of muscle fibers

due to the concurrent increase in muscular loading. However, we found no significant

differences in mean CSA or in the mean Type 2 fiber CSA between groups (Figure 2A-

D).

Finding no increase in the average CSA of myofibers, we chose to investigate if

the fiber size distribution of the Type 2 fibers was altered, perhaps indicating a degree of

12

atrophy. Type 2 fibers are classified into 3 separate sub-groups: Type2A, 2X, and 2B.

Type 2A fibers are fast-twitch fibers with a high oxidative potential, similar to Type 1/slow-

twitch fibers. Type 2X fibers are fast-twitch high glycolytic intermediate muscle fibers,

which possess low quantities of mitochondria, but a large pool of glycolytic enzymes.

Finally, Type 2B fibers are a fast-glycolytic fiber type, characterized by large myofiber size

and glycogen content (Pette 2005). The greatest percentage of Type 2A fibers were

grouped in the 501-1000μm2 range (Figure 3A). Interestingly, the Sedentary+Statin group

had a significantly reduced percentage of fibers within this range compared to all other

groups. This suggests that statins affect this particular fiber range specifically and

exercise maintains this population of myofibers. For Type 2X fibers, the greatest

percentage of all myofibers fell within the same 501-1000μm2 as Type 2A fibers but with

no significant differences between groups within any of the size ranges (Figure 3B).

The most significant size alterations occurred in the Type 2B fibers (Figure 3C).

This fiber type was affected variably by both treatment and exercise within different

ranges, specifically within the 1001-1500μm2 and >3000μm2 ranges. The

Accustomed+Statin group demonstrated various size ranges with an increased

percentage of fibers in the 1001-1500μm2 range, but a significantly reduced percentage

in the large fiber size ranges. This finding suggests the possibility that only select fiber

types are impacted by statins, and that in conjunction with exercise, these changes can

be exacerbated. A separate study using rats provided evidence that statins affect muscle

myofibers in a manner dependent on their oxidative or glycolytic metabolic nature, with

glycolytic fibers being the most sensitive to statin usage (Westwood 2005). This finding

agrees with the data that Type 2B fibers are affected the most. As such, the myofiber size

13

distribution illustrates a diverse effect that both statins and prior exposure to exercise

have on fiber size, especially in Type 2B fibers.

Given the diverse range of sizes present for each fiber type and lack of average

fiber type CSA differences, we examined the percentage of the different Type 2 fibers in

order to determine if a particular type was specifically altered by statins and exercise.

Interestingly, there was a trend for statins and exercise together to alter the percentage

of Type 2X and 2B fibers within skeletal muscle. Overall, these findings raise the

possibility that statins and exercise interact to preferentially effect larger fast-twitch and

glycolytic fibers versus smaller slow-twitch and oxidative fibers.

This study provides a unique perspective on the combined effect of statins and

exercise on myofiber structure and morphology. Our findings indicate that reductions in

strength associated with short-term statin use and exercise are not caused by increase

myofiber damage; however, we present interesting data in regards to the effect on

myofiber size and the percentage of Type 2 fibers, specifically in Type 2B fibers within

skeletal muscle. These fibers and their respective sizes are vital for maximal strength

production to perform any physically demanding task, as there has been a strong

correlation shown between the CSA and maximum strength of a muscle (Seitz 2016).

Furthermore, two important aspects of Type 2 fibers are their size and significant motor

unit innervation. Without substantial neural activation, these fibers are unable to utilize

their full force production potential. While the number of samples used in the study are

relatively low, our findings demonstrate a limited impact on fiber CSA and muscle

damage, leading us to hypothesize that disruptions to the neuronal innervation of Type 2

fibers could contribute to reduced strength production in hypercholesterolemic mice.

14

Further investigations into how statins affect other aspects of skeletal muscle, such as

neuronal innervation, are warranted in order to more fully understand the complex

relationship between statins, exercise, and myopathy.

15

Figure 1

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19

Figure Legends

Figure 1. The effect of statin treatment and exercise on myofiber damage and regeneration. (A) The ratio of IgG+ myofibers per area of muscle (mm2) was assessed to determine the amount of myofiber damage that occurred during a combination of statin and exercise therapy. (B) Percentage of myofibers displaying a centrally-located nucleus, a hallmark of damage and repair/regeneration. No significant differences were detected between groups. Figure 2. The effect of exercise and statin administration on mean cross sectional area. (A) The mean myofiber CSA, as well as the mean CSA for (B) Type 2A, (C) Type 2X, and (D) Type 2B fibers were assessed. No significant differences were detected between groups. Figure 3. Fiber type-specific size distribution is differentially affected by a combination of exercise and statin therapy. The percentage of (A) Type 2A, (B) Type 2X, and (C) Type 2B myofibers categorized by size. The combination of statin and exercise altered Type 2X and Type 2B myofiber size. *p<0.05 compared to all other

groups; ǂ†*p<0.05 compared to Sedentary+Statin, Novel+Saline, Accustomed+Saline; ǂ

*p<0.05 compared to Sedentary+Saline, Accustomed+Saline. Figure 4. The percentage of Type 2 fibers is selectively affected by a combination of exercise and statin therapy. The percentage of (A) Type 2A, (B) Type 2X, and (C) Type 2B fibers was unaffected by statin or exercise alone; however, a trend for a statin x exercise interaction effect was observed for Type 2X (p=0.07) and Type 2B (p=0.06).

20

Acknowledgements

I would like to thank Michael Munroe for his guidance and support throughout the entire

project. He has been extremely helpful as a mentor and an important factor in the

completion of my senior thesis. I would also like to thank my faculty advisor, Dr. Marni

Boppart, for her support and providing me with the opportunity to complete this work. I

would also like to acknowledge Dr. Hae R. Chung for completing the initial study and

performing all the functional tests with mice. I would also like to thank Ziad Mahmassani

and Slav Dvoretskiy for their assistance with this project. This work was supported by a

grant from the Center for Health Aging and Disease (to MDB).

21

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