THE ROLE of KYNURENINE, a TRYPTOPHAN METABOLITE THAT INCREASES with AGE, in MUSCLE ATROPHY and LIPID PEROXIDATION) by Helen Eliz

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THE ROLE OF KYNURENINE, A TRYPTOPHAN

METABOLITE THAT INCREASES WITH AGE, IN MUSCLE

ATROPHY AND LIPID PEROXIDATION)

Helen Elizabeth Kaiser

Submitted to the Faculty of the Graduate School

of Augusta University in partial fulfillment

of the Requirements of the Degree of

(Doctor of Philosophy in Cellular Biology and Anatomy)

April

2020

ACKNOWLEDGEMENTS

I wish to express my deepest gratitude to my mentor Dr. Mark Hamrick. His support and advice were invaluable to my learning and development as a scientist. I would like to especially recognize Dr. Hamrick’s honesty to data, and kindness to everyone around him. It was an honor to work as his student, and to be a part of this project. I would like to thank my committee members, Drs. Carlos Isales, Meghan McGee-Lawrence, and

Yutao Liu, for their continued support, guidance, and helpful suggestions. They have helped to shape and focus this project, and have augmented my experience as a student during the development of my dissertation. Additionally, I want to recognize and thank

Dr. Sadanand Fulzele for sharing his wealth of knowledge in techniques, and always keeping his door open for student’s questions. Next, a most heartfelt thanks to my lab mates: Andrew Khayrullin, Bharati Mendhe, Ling Ruan and Emily Parker. This project has been a team effort, all of them have been fundamental to its success.

I further want to show my appreciation for the rest of the department of

Cellular Biology and Anatomy at Augusta University. The histology core members

Donna Kumiski and Penny Roon for their direct help in sectioning, and contributions to this project. Also Cotton Mitchell, Diane Riffe, and Donna Tuten for their administrative support, and their constant encouragement.

I thank Drs. Watsky and Cameron and the Graduate School team for allowing me the opportunities and resources to grow as both a leader and professional over the years, aiding me with career-developing opportunities and travel support to present my work at

Experimental Biology in 2018, and The International Conference of Frailty and

Sarcopenia in 2019.

I thank my family for their unwavering support as I've gone through the highs and lows of my professional degree. I am especially grateful to my husband, Jacob Kaiser. His unconditional love, support, and encouragement over the years have been vital to this journey. Finally, to my dogs: Gordon, Mica and Oti for their love and helpful suggestions while writing this thesis.

ABSTRACT

HELEN KAISER

The Role of Kynurenine, a Tryptophan Metabolite that increases with age, in

Muscle Atrophy and Lipid Peroxidation.

(Under the direction of DR. MARK HAMRICK)

Loss of mobility and independence are risk factors for falls and mortality, and drastically reduce the quality of life among older adults. The cellular and molecular mechanisms underlying loss of muscle mass and strength with age (sarcopenia) are not well- understood; however, heterochronic parabiosis experiments show that circulating factors are likely to play a role. Kynurenine (KYN) is a circulating tryptophan metabolite that is known to increase with age and is implicated in several age-related pathologies. Here I tested the hypothesis that KYN contributes directly to muscle loss with aging. Results indicate that that KYN treatment of mouse and human myoblasts increased levels of reactive oxygen species (ROS) two-fold, and significantly increased lipid peroxidation enzymes. Small-molecule inhibition of the Aryl hydrocarbon receptor (Ahr), an endogenous KYN receptor, in vitro did not prevent KYN-induced increases in ROS, and homozygous Ahr knockout in vivo did not protect mice from KYN-induced stress, suggesting that KYN can directly increase ROS independent of Ahr activation. In vivo, wild-type mice treated with KYN had reduced skeletal muscle strength, size, and increased oxidative stress and lipid peroxidation. Old wild-type mice treated with 1MT, a small molecule that suppresses KYN production by IDO1, showed an increase in muscle

fiber size, peak muscle strength, and oxidative stress. Protein analysis identified mitochondrial lipid peroxidation as a downstream mechanism that is increased upon

KYN treatment. Lipid peroxidation enzymes increased with KYN have been shown to produce H2O2 outside of the electron transport chain. Our data suggest that IDO inhibition may represent a novel therapeutic approach for the attenuation of sarcopenia and possibly other age-associated conditions associated with KYN accumulation such as bone loss and neurodegeneration

KEY WORDS: (Muscle, sarcopenia, tryptophan, kynurenine, lipid peroxidation, muscle atrophy, aging,

Table of Contents

Chapter Page

1 Introduction and specific aims ...... 1

1.1 Statement of a problem ...... 1

1.2 Specific aims ...... 4

1.3 Innovation ...... 5

2 Literature review ...... 6

2.1 Healthspan vs. Lifespan ...... 6

2.2 Sarcopenia ...... 7

2.3 ROS and muscle atrophy ...... 11

2.4 Circulating factors ...... 14

2.5 Kynurenine in health and disease ...... 16

3 Materials and Methods ...... 28

3.1 Cell culture ...... 28

3.2 MTT assay ...... 28

3.3 AmplexTM Red assay ...... 29

3.4 Total Antioxidant Capacity assay...... 29

3.5 Animal experimental design ...... 30

3.6 Histological staining ...... 31

3.7 Proteomics ...... 32

3.8 Western blots ...... 32

3.9 PCR array plates ...... 33

3.10 In vivo functional muscle testing ...... 33

3.11 Ex vivo muscle testing ...... 34

3.12 Open field activity assays ...... 36

3.13 Cytokine array panels ...... 36

3.14 Statistical analysis...... 37

4 Results ...... 38

4.1 KYN treatment increases ROS levels in C2C12 mouse and primary

human myoblasts...... 38

4.2 Ahr inhibition did not rescue KYN effects in vitro...... 40

4.3 KYN treatment produces an aged-muscle phenotype in young female .43

4.4 Ahr knockout weakly atenuated KYN effects in female mice...... 46

4.5 1-MT treatment decreased oxidative stress and attenuates muscle

atrophy old female mice ...... 48

4.6 Peak muscle strength is attenuated with KYN ...... 50

4.7 PCR arrays reveal structural muscle proteins are decreased by KYN .52

4.8 KYN did not change intracellular lipid accumulation ...... 54

4.9 Fiber type proteins are reduced with KYN treatment ...... 56

4.10 Proteomic analysis of KYN treated and 1-MT treated skeletal muscle

reveals differential expression of mitochondrial very long chain acyl-

coa dehydrogenase ...... 58

4.11 Old female mice treated with 1-MT reduced activity 30 days after 1-

MT treatment ...... 61

4.12 Muscle and serum cytokines were altered with 30 days of KYN

treatment...... 63

5 Discussion ...... 66

6 Summary ...... 74

References ...... 76

Appendices ...... 99

A. Abbreviations ...... 99

List of Tables

Table 1. Exogenous and endogenous ligands of the aryl hydrocarbon receptor and their potential impact on age-related diseases...... 27

List of Figures

Figure 1 The central hypothesis...... 3

Figure 2. Factors leading to and protein anabolism and catabolism...... 8

Figure 3 Oxidative species contribute to lipid peroxidation...... 12

Figure 4 The kynurenine pathway of tryptophan breakdown...... 17

Figure 5. Kynurenine pathway with exercise and age. ophan into kynurenine...... 20

Figure 6. KYN-AHR signal transduction pathway...... 23

Figure 7. Factors regulating Ahr gene expression...... 25

Figure 8 Aurora functional muscle testing system...... 35

Figure 9. ROS is increased in mouse and human myoblasts with KYN compared to VEH treatment...... 39

Figure 10. Ahr inhibition in vitro...... 42

Figure 11 Muscle loss is increased in young mice treated with KYN...... 44

Figure 12. Skeletal muscle lipid peroxidation is increased in young mice with KYN treatment...... 45

Figure 13. Ahr knockout in female mice...... 47

Figure 14. Skeletal muscle H2O2 and morphology changes after 1-MT treatment in aged female mice...... 49

Figure 15. Peak muscle strength was decreased with KYN treatment in young mice and increased with 1-MT treatment in old mice...... 51

Figure 16. Structural protein mRNA was decreased with KYN treatment in young female mice...... 53

Figure 17. Intracellular lipid content did not change with KYN treatment in male or female mouse quadriceps...... 55

Figure 18. Fiber type changes in human myotubes treated with KYN...... 57

Figure 19. Lipid peroxidation enzymes are differentially expressed with KYN or 1-MT treatment...... 60

Figure 20. Open field activity after KYN or 1-MT treatment in young and old, male and female mice...... 62

Figure 21. Inflammatory cytokines changed with KYN treatment...... 65

Figure 22. Proposed model for KYN’s effects on lipid peroxidation, ROS generation, and age-related muscle atrophy...... 73

I. INTRODUCTION AND SPECIFIC AIMS

1.1 Statement of a problem

The population of adults 65 years and older is growing rapidly in the United

States and in countries around the world (United Nations, 2013). Aging is associated with a loss of muscle mass and strength called sarcopenia, which increases the risk for falls and debilitating bone fractures (Rubenstein, Josephson, & Robbins, 1994; Tinetti, 1994).

The cellular and molecular processes leading to sarcopenia are not well understood, which presents the aging field with a significant knowledge gap. The goal of this study is to address this gap by defining the cellular and molecular mechanisms underlying the progression of sarcopenia. Kynurenine (KYN) is a circulating tryptophan metabolite that increases with age and is correlated with increased mortality in older adults (Pertovaara et al., 2006). Elevated circulating levels of KYN are also found to be associated with osteoporosis and Alzheimer’s disease (Breda et al., 2016; Refaey et al., 2017), as well as with bone loss, coronary artery disease, cachexia, and oxidative stress (Eussen et al.,

2015; B. J. Kim et al., 2019; Laviano, Meguid, Preziosa, & Rossi Fanelli, 2007).

The central hypothesis (Figure 1) is that an increase in KYN with age causes an imbalance in lipid peroxidation pathways, leading to oxidative stress and consequent muscle atrophy. This hypothesis is supported by 1) studies demonstrating a role for KYN in stimulating bone loss (B. J. Kim et al., 2019; Refaey et al., 2017), 2) research showing that KYN may contribute to coronary artery disease with aging (Eussen et al., 2015) as well as cancer-induced muscle wasting (Laviano et al., 2007), and 3) studies showing

increased KYN can change lipid metabolism, as lipid peroxidation is increased in adults with sarcopenia (Pratico, 2002; Schnyder & Handschin, 2015)

Figure 1 The central hypothesis: (A.) As people age and systematic inflammation is increased, IDO is activated to convert tryptophan to kynurenine, which contributes to further oxidative stress and lipid peroxidation, weakening cellular membranes, and (B.) increasing overall muscle atrophy.

1.2 Specific Aims

Specific Aim 1: Test the hypothesis that age-associated increases in KYN impairs muscle function and decreases muscle mass by activating ubiquitin ligases and inhibiting protein synthesis. First, mouse and human primary myoblasts were treated with KYN to examine gene and protein expression changes. Next young mice were treated with KYN to determine effects on muscle fiber size, muscle mass, and muscle contractile force. Aged mice were then treated with the IDO inhibitor 1-MT to inhibit KYN production and to determine effects of 1-MT treatment on muscle fiber size, muscle mass, and muscle contractile force, and in vitro expression of genes involved in muscle atrophy, and protein synthesis was examined in all in vitro and in vivo experiments. Protein was measured using proteomic arrays, and western blot for protein synthesis and fiber type proteins (Mhc type I, IIa, IIb).

Specific Aim 2: Test the hypothesis that KYN increases oxidative stress and lipid peroxidation in skeletal muscle cells. The effects of KYN on ROS production in human primary myoblasts were determined using AmplexTM red and lipid peroxidation assays. Lipid peroxidation enzyme expression (Mvlcad, Mlcad) was measured in C2C12 mouse myoblasts and human myoblasts using PCR and western blot. Circulating inflammatory markers were measured from the serum of mice treated with KYN or 1-MT vs. mice treated vehicle control (VEH). Finally, ROS and lipid peroxidation was measured in vivo in skeletal muscle by measuring hydrogen peroxide, 4-hydroxynonenal

(4HNE), and malondialdehyde (MDA) levels KYN and 1-MT treated mice.

1.3 Innovation

There are a number of novel and innovative aspects to this study. First, this was the first study exploring a role for the KYN pathway in skeletal muscle. Second, there are no FDA approved drugs currently used to treat sarcopenia. The proposed IDO inhibitor,

1-MT, is currently in ongoing phase 1 clinical trials for several types of cancer (Soliman et al., 2014). Repurposing an IDO inhibitor for sarcopenia would be feasible, and could capitalize on many studies already completed in toxicity, dosage, safety, and effects

(Coto Montes et al., 2017; Kakimoto, Tamaki, Cardoso, Marana, & Kowaltowski, 2015).

Additionally, this study used several innovative experimental approaches including functional testing of muscle in vivo, in vitro analysis of KYN, Ahr interactions, and proteomic analysis of muscle in a mouse model of sarcopenia. These innovative approaches shed new light on a pathway unexplored in skeletal muscle and provides potential therapeutic targets for the treatment of sarcopenia.

II. LITERATURE REVIEW

2.1. Healthspan versus Lifespan

The population of people 60 years of age or older is expected to increase from 8% of the world’s population in 2013 to an estimated 21% of the world’s population by 2050.

This increase means that in the next 40 years there will be more than 2 billion people over 60 years of age (United Nations, 2013). Extending the healthspan of the world population so that aged people can remain disease-free and independent is paramount toward easing the burden of medical costs and increasing the quality of life. A significant reason aged people lose independence is a fall, with an estimated 30% of people over 65 falling yearly (Rubenstein et al., 1994; Tinetti, 1994). After a fall in-hospital and postdischarge mortality rates are significantly higher than aged people with blunt trauma injuries. Those with a fall are also more likely to lose independence (Gerrish et al., 2018).

Falls in aged people often cause fractures and other trauma-related injuries

(Faulkner et al., 2009). Falls can also induce fall-associated anxiety, in which the person who fell no longer partakes in activities, further facilitating a loss of health and accelerating depression. Approximately half of people over 65 who fall are injured from a fall. Furthermore, Female gender and fear of falling were significant risk factors for future falls (Gazibara et al., 2017; Hoang, Jullamate, Piphatvanitcha, & Rosenberg,

2017). Risk factors for falling are multifactorial and include gait changes, loss of balance, slower reflexes and lower limb muscle strength (Martin et al., 2013; Menant et al., 2012;

Pijpers et al., 2012; Scheffer et al., 2013). The loss of lower limb muscle quadriceps femoris strength significantly increases the risk of falls (Ahmadiahangar et al., 2018).

Multiple studies have shown that 1-year mortality rates after falls and fractures are

significantly higher in those with sarcopenia (Deren et al., 2017; Y. K. Kim et al., 2018;

Mitchell, Collinge, O'Neill, Bible, & Mir, 2018). Furthermore, hospitalization rates increase significantly in those with sarcopenia (Zhang et al., 2018).

2.2 Sarcopenia

The loss of skeletal muscle homeostasis is a major contributor to the age-related decline in independence. The loss of muscle mass and power with age is called sarcopenia (Santilli, Bernetti, Mangone, & Paoloni, 2014). Sarcopenia occurs in over one-third of people over 70 years of age (Brown, Harhay, & Harhay, 2016) and is a multifactorial disease with unknown causes. Sarcopenia can be characterized by several broad symptoms such as generalized muscle atrophy, increases in systemic inflammatory cytokines, replacement of muscle fibers with fibrotic factors and fat, and loss of muscle power (Ryall, Schertzer, & Lynch, 2008).

The balance of skeletal muscle is maintained through life by the interactions of protein anabolic factors, otherwise known as “myogenes”, and protein catabolic factors, known as “atrogenes” (Figure 2). In conjunction with skeletal muscle satellite cells these myogenes and atrogenes build new skeletal muscle and breakdown old skeletal muscle, respectively. In skeletal muscle atrophy models of disuse and acute disease the ubiquitin ligase atrogenes Murf1 and Mafbx are highly upregulated (Attaix et al., 1998).

Meanwhile, myogenes such as Myogenin and MyoD that stimulate differentiation of muscle stem cells into mature myotubes are arrested by P53 during stress, resulting in decreased satellite cell differentiation and fusion (Z. J. Yang et al., 2015).

Figure 2. Factors leading to and protein anabolism and catabolism. The balance of skeletal muscle homeostasis is maintained by a network of environmental, self-induced, and molecular factors. Resistance training, and a leucine-enriched protein diet are the only known treatments for sarcopenia. High levels of protein catabolism can occur due to the actions of MyoD and myogenin, muscle differentiation factors. In contrast, Low levels of vitamin D, low protein intake, lowered testosterone and estrogen levels are attributed to a loss in muscle strength. Skeletal muscle atrogenes, Murf1 and Mafbx are upregulated with muscle loss. Several circulating factors, IFN-γ, TNF-α, KYN/TRP ratio and transcription factor NF-κβ are seen upregulated with muscle loss during aging.

After the second decade of life, reduced vitamin D levels and lower protein and calorie intake are correlated with weaker muscles (Houston et al., 2008). Both men and women lose testosterone and estrogen levels, respectively, which contribute to declines in muscle mass (Baumgartner, Waters, Gallagher, Morley, & Garry, 1999; Samson et al., 2000).

Males and females can both become sarcopenic, but the pathologies seem to have some differences (Tay et al., 2015). For example, men have a higher rate of skeletal muscle loss, and women with skeletal muscle loss experience higher rates of mortality than men

(Batsis, Mackenzie, Barre, Lopez-Jimenez, & Bartels, 2014; Payette et al., 2003). One study attributed these differences to myostatin levels in men catabolizing skeletal muscle, whereas lowered IGF-1 in women led to less skeletal muscle growth in women (Tay et al., 2015). Another group, showed no increases in myostatin with age in women (Bergen et al., 2015). Anderson et al. proposed that sarcopenia is a common disease, but hormone differences in men and women effect the course of the disease (Anderson, Liu, & Garcia,

2017).

Older patients with skeletal muscle loss have increased levels of the pro- inflammatory cytokines TNF-α and IL-6 (Meng & Yu, 2010). Il-6 has also been suggested as a predictor for sarcopenia (Aleman, Esparza, Ramirez, Astiazaran, &

Payette, 2011). TNF-α and IL-6 are mediated by the transcription factor nuclear factor-κβ

(NF- κβ), which responds to reactive oxidative species and directs transcription of pro- inflammatory cytokines on a short-term basis. But during chronic stress, activation of

NF- κβ remains elevated (Yu & Chung, 2006). Furthermore, TNF-α itself activates NF-

κβ, creating a feed-forward loop that requires external factors to resolve (Handel,

McMorrow, & Gravallese, 1995). Normal skeletal muscle functions closely with the

immune system, as it is capable of regenerating after injury to be capable of exerting more force than before injury. Growth factors secreted by immune cells are essential for muscle stem cells (satellite cells) to proliferate and differentiate (Saini, McPhee, Al-

Dabbagh, Stewart, & Al-Shanti, 2016) yet immune cells secrete fewer growth factors with age (Domingues-Faria, Vasson, Goncalves-Mendes, Boirie, & Walrand, 2016).

These dysregulated pathways provide promise for several therapeutic pathways that could be targeted, but more work needs to be done. Currently resistance training and leucine- enriched protein diet are used to manage sarcopenia, but no FDA approved medications are available to treat sarcopenia (Abate et al., 2007).

Skeletal muscle is composed of skeletal muscle cells (myofibers) that can exhibit different metabolic and functional characteristics. These are colloquially referred to as fast-twitch (type 2) and slow-twitch (type 1) muscle fibers. Fast-twitch fibers are primarily anaerobic, with high glycolysis and low mitochondrial content, whereas slow- twitch fibers are primarily aerobic, with high mitochondria content (Y. Wang & Pessin,

2013). Several studies in the 1970’s were performed to analyze the fiber type composition in different types of athletes to determine fiber function. They found that sprinters had muscles made up of mostly fast-twitch fibers, and endurance runners had more slow-twitch fibers (Costill et al., 1976; Fink, Costill, & Pollock, 1977; Saltin,

Henriksson, Nygaard, Andersen, & Jansson, 1977).

Muscles in un-trained people tend to be comprised of a general pattern. Quadriceps femoris is typically 50% slow-twitch and 50% fast-twitch, interestingly, slow-twitch fibers tend to have larger cross-sectional areas, but fast-twitch fibers have about 2x the strength per area (Young, 1984). In the mouse, soleus is primarily slow-twitch, and

extensor digitorum longus (EDL) is primarily fast-twitch (Schiaffino & Reggiani, 2011;

Sher & Cardasis, 1976). A muscle fiber type shift has been observed with age, with older people losing the number and cross sectional area of fast-twitch fibers (Deschenes, 2004;

Nilwik et al., 2013). However, a loss in slow-twitch fibers has also been seen with age, with less significant of a change than fast-twitch (Kramer et al., 2017).

2.3 ROS and muscle atrophy

Several cellular functions produce toxic reactive oxygen species, many as a by-product from the electron transport chain (Ingram et al.) (Li et al., 2013). The ETC consists of a series of enzymatic redox complexes in the inner mitochondrial membrane that pass H+ ions to eventually synthesize stored energy in the form off adenosine triphosphate (ATP) and H2O (Li et al., 2013). Most oxygen passed through the ETC is reduced to H2O, however approximately 1% of oxygen is incompletely reduced and escapes the ETC as

− − O2 (Li et al., 2013). The antioxidant enzyme superoxide dismutase can convert O2 to

H2O2. Then H2O2 can be converted to H2O and O2 by glutathione peroxidase (Figure 3), the Fenton reaction or the Haber-wells reactions (Misra, Sarwat, Bhakuni, Tuteja, &

− Tuteja, 2009). O2 and H2O2 can both produce toxic levels of damage in cells, with H2O2 being used by epithelial cells as host defense (van der Vliet & Janssen-Heininger, 2014).

For instance, H2O2 is produced heavily in wound responses to fight potential infection

(Gauron et al., 2013).

Figure 3 Oxidative species contribute to lipid peroxidation. Electrons that leak from the electron transport chain react with oxygen to produce superoxides. Superoxides are reduced by superoxide dismutase (SOD), and glutathione peroxidase (GPx). These reactive oxygen species (outlined in red) all contribute to lipid damage and peroxidation, which produces 4HNe.

Despite antioxidant systems, and endogenous uses for ROS, a build-up of cellular

ROS has been consistently shown to occur in all aging tissues and is correlated with disease. While oxidative stress consistently is a symptom of aging, antioxidant therapies have been disappointing. Antioxidants supplemented after exercise in healthy young men impair the beneficial effects of exercise by blocking downstream signaling cascades induced by endogenous antioxidants (Ristow et al., 2009). Skeletal muscle contractions produce upregulated levels of ROS regardless of the type and duration of muscle use

(Vollaard, Shearman, & Cooper, 2005). In trained muscle ROS can be beneficial at high levels, whereas in untrained muscle ROS is beneficial at low levels and detrimental at high levels (Radak, Taylor, Ohno, & Goto, 2001; M. A. Smith & Reid, 2006). The beneficial effects include increased mitochondrial biogenesis, insulin sensitivity, and higher antioxidant capacity, while detrimental effects include mitochondrial dysfunction,

DNA mutations, lipid peroxidation and muscle degradation (Radak et al., 2001; M. A.

Smith & Reid, 2006).

Furthermore, the duration of exposure to ROS has differential effects: short and low exposure to H2O2 increases contractile force, while chronic exposure inhibits contractile force (Andrade, Reid, Allen, & Westerblad, 1998; Prochniewicz, Spakowicz,

& Thomas, 2008). Muscle contraction increases ROS levels, but also increases antioxidant systems simultaneously, with increases in superoxide mutase, and glutathione peroxidase seen with exercise training (Gore et al., 1998). These studies show that acute increases in ROS are helpful and protective, but chronic increases in ROS can lead to tissue damage.

Lipid peroxidation is the process by which lipids are broken down and weakened by oxidative stress (Figure 3). In this process, an oxidative species will accept an electron from a double bond in a polyunsaturated fatty acid, creating a lipid peroxide (Pratico,

2002). With age, ROS accumulation consequently contributes to lipid peroxidation

(Pratico, 2002). Skeletal muscle has previously been shown to be susceptible to lipid peroxidation damage. In a model of Duchenne’s muscular dystrophy in which dystrophin, an integral membrane anchoring protein is mutated, lipid peroxidation pathways are upregulated and contribute to the pathogenesis of the disease (Messina et al., 2006).

However, exercise is an effective protectant against age-related lipid peroxidation. Long term exercise training lowers lipid peroxidation with age. (Barranco-Ruiz et al., 2017)

And even in a rat model of obesity, exercise lowers lipid peroxidation (Morris et al.,

2008). As with all oxidative stress, it is important to recognize that a regulated balance of lipid peroxidation is required for healthy cellular processes (Peternelj & Coombes, 2011).

Several studies have shown a beneficial effect for normal levels of lipid peroxidation, which may be a mechanism to aid in cellular proliferation and differentiation (Barrera,

Pizzimenti, & Dianzani, 2008). Intracellular lipid droplet accumulation is seen during aging and skeletal muscle loss, but whether lipid peroxidation upregulation occurs due to increased lipid accumulation or through other pathways remains to be tested (Gueugneau et al., 2015) (Rivas et al., 2016).

2.4 Circulating factors

The search for the “fountain of youth” is a rapidly expanding area of research that has inspired a search for aging biomarkers that could clinically predict predisposition to

age-related disease. In the 1950s several groups used parabiosis experiments to connect two mice by their circulatory systems, and found that irradiated mice could survive longer when paired with non-irradiated mice (Binhammer, Schneider, & Finerty, 1953;

Finerty, Binhammer, & Schneider, 1952). More recently experiments using heterochronic parabiosis, in which an old mouse is paired with a young mouse have provided insight to age-related factors. When the muscle of an aged mouse in a pair was wounded, the aged mouse could regenerate muscle faster than an unpaired aged mouse (Conboy et al., 2005).

Villeda et al expanded on these experiments and showed that young mice mimicked age- related cognitive impairments when paired with an old mouse. (Villeda et al., 2011;

Villeda et al., 2014). Mice must be genetically similar, and living in the same environment before they can be considered as candidates for parabiosis (Scudellari,

2015). These studies together suggested that some factor/s found in the circulation are increased with age, so that identifying and targeting these factors is a promising therapeutic intervention for age-related disease. Established markers for general muscle weakness include serum creatine concentration, 3-methylhistidine, type VI collagen, agrin, and myostatin (Kemp et al., 2018).

The search for aging biomarkers carries a dual purpose. First, to clinically predict accelerated aging in patients, and second, to find targets to combat the decline of function that comes with age. Several candidates have been identified that are consistently associated with age, frailty, muscle loss, and inflammation. In a recent study of 180 patients in Italy the top candidates that were associated with declines in grip strength, gait speed overall frailty were Neopterin, tryptophan, nitrite and C-reactive protein (Valdiglesias et al., 2018). Another study measured Interleukin-6 levels in adults

70-85yo that had been sedentary for the previous months. The top metabolite associated with high Il-6 levels was KYN, a metabolite of tryptophan (Lustgarten & Fielding, 2017).

Most recently, 166 adults spanning 20 to 97 years were studied to compare inflammation and frailty with serum markers. The top candidates associated with age and frailty status were KYN (<.0001), and the KYN/ tryptophan ratio (<.0001) (Reyhan Westbrook et al.)

(Figure 2). These studies point to KYN as a possible target for inflammation and ageing.

2.5 KYN in health and disease

Under normal conditions, approximately 10% of tryptophan is broken down into the serotonin pathway which produces serotonin and melatonin (Stone & Darlington, 2002).

The other 90% of tryptophan is broken down in the KYN pathway by tryptophan 2,3- dioxygenase (TDO) in the liver and by indoleamine 2, 3, dioxygenase 1 (IDO1) extrahepatically (Stone & Darlington, 2002) (Figures 4 and 5). The KYN pathway is outlined in Figure 4. IDO1 is found in the gut, endothelium, brain, bone, skeletal muscle, and lymph tissues in low concentrations during healthy conditions (Fallarino et al., 2002).

IFN-γ induces the transcriptional activation of IDO1 and IDO2 primarily, but is also activated by interferon alpha, interferon beta, and lipopolysaccharide (LPS), (Grohmann,

Fallarino, & Puccetti, 2003; Mellor & Munn, 2004; Munn et al., 1999). Tryptophan activation of IDO arrests T-cell proliferation (Munn et al., 1999), and also suppresses infectious microorganisms (A. Muller et al., 2009). Tryptophan depletion was seen along with upregulated P53 in alloreactive T cells with activated IDO (Eleftheriadis et al.,

2014). IDO suppresses T-cells, closely regulates the growth in the local

Figure 4 The kynurenine pathway of tryptophan breakdown. Tryptophan can broken down into serotonin, or into the kynurenine pathway. Enzymes that degrade tryptophan are shown in blue TDO, tryptophan-2,3- dioxygenase; IDO, indoleamine-2,3-dioxygenase; KATI, kynurenine aminotransferase I; KATII, kynurenine aminotransferase II; KMO, kynurenine-3-monooxygenase; KYNU, kynureninase; HAAO, 3- hydroxyantrhanilic acid oxidase; QPRT, quinolinate phosphoribosyl transferase. Figure and caption borrowed with permission from (Formisano et al., 2017)

microenvironment, and can allow tumors to escape immune control (Munn & Mellor,

2007). Maternal KYN serum levels acutely increase, and further KYN metabolite increases are seen in the fetus suggesting an important role for KYNs during development (Goeden et al., 2017). Furthermore, the KYN pathway has been implicated in immune suppression at the fetal-maternal barrier during pregnancy (Munn et al.,

1998). The proposed mechanism was thought to be tryptophan depletion, but more recent evidence suggests KYN metabolites may play a stronger role (Badawy, Namboodiri, &

Moffett, 2016; Zadori, Klivenyi, Plangar, Toldi, & Vecsei, 2011).

The KYN pathway is upregulated in inflammation and has further been linked to a wide variety of age-related pathologies. Whether systemic increases in inflammation simply increase IDO activity, or KYN itself induces age-related pathologies remain to be determined. KYN pathway over-activation has been implicated in Alzheimer’s disease, parkinson’s disease, Huntington’s disease, depression, schizophrenia, cardiovascular disease, osteoporosis, diabetes, inflammatory bowel disease and rheumatoid arthritis

(Agudelo et al., 2014; Breda et al., 2016; Oxenkrug, 2013; Refaey et al., 2017; Williams,

2013; Wolf et al., 2004; Wonodi & Schwarcz, 2010).

In addition to the previously mentioned tryptophan starvation, and T-cell suppression models, further molecular mechanisms have been explored. Agudelo et al., identified

PGC-1α as a modulator of the KYN pathway (Agudelo et al., 2019). Agudelo further demonstrated that PGC-1α increases the transcription of KYN aminotransferase’s

(KATs), converting KYN to kynurenic acid, a metabolite that can activate a Gpr35, to further increase PGC-1α expression (Agudelo et al., 2018). PGC-1α is a transcription factor induced by exercise that acts to support mitochondrial health, fatty acid oxidation,

and energy metabolism (Schnyder & Handschin, 2015). In primary mouse myotubes,

10µM KYN reduced skeletal muscle oxidative consumption rate significantly, and in a muscle specific knockout of PGC-1α respiration rates were reduced further (Agudelo et al., 2019). The significance of these findings is that KYNA is important for downstream positive effects in skeletal muscle induced by exercise, but KYN buildup in the absence of exercise is detrimental to muscle. A summarization of the KYN pathway is described in Figure 4.

Figure 5. Kynurenine pathway with exercise and age: Inflammatory cytokines arising from systematic inflammation (IFN-γ, IFN-α, IFN-β,), and LPS activated IDO to convert Tryptophan into kynurenine. Kynurenine is involved in T-cell suppression, tryptophan starvation, and oxidative stress. KATs are upregulated with exercise and convert kynurenine to neuroprotective kynurenic acid. This may also temporarily deplete the de novo NAD+ synthesis. Without KATs, kynurenine is converted to neurotoxic quinolinic acid, and then by QAPRT into NAD+. NAD+ is also depleted with age, which could be in part due to a decrease in QAPRT and a buildup of kynurenine and quinolinic acid.

Inhibition of KYN synthesis has shown promise for therapeutic intervention for re-establishing immune surveillance in a number of cancers. A small-molecule, selective inhibitor of IDO, 1-methyl-tryptophan (1-MT) was first shown to be as competitive antagonist inhibitor of IDO in the early 90s (Cady & Sono, 1991). Early work showed promise for reducing colon tumor growth, and has been in clinical trials both alone, and with Docetaxel (Hou et al., 2007; Soliman et al., 2016). 1-MT is non-toxic, with an ic(50) of 10nM (Jia et al., 2008; Liu et al., 2010). Further studies have shown that 1-MT is therapeutic alone or in combination with other established chemotherapeutic agents in

Lung cancer, Bladder cancer, Melanoma, or ovarian cancer (Friberg et al., 2002; Hou et al., 2007; Inaba et al., 2009; H. J. Yang et al., 2010) 1-MT use in musculoskeletal pathologies or aging has been more limited. 1-MT use in a mouse model of rheumatoid arthritis lowered inflammatory cytokine levels, and alleviated arthritis symptoms (Scott et al., 2009). 1-MT use in skeletal muscle has not been studied, however some work on the

KYN pathway has sought to understand how KYN may be affecting age-related disease mechanistically.

KYN has been shown to be detrimental in several age-related diseases, but the pathology downstream of KYN remains unclear. KYN is an endogenous ligand of the

Ahr (Bessede et al., 2014; Mezrich et al., 2010). The Ahr is cytosolic and bound several chaperone proteins including heat shock protein 90. Once ligand activated, the bound Ahr translocates to the nucleus and binds to aryl hydrocarbon receptor nuclear transporter

(Arnt). The Ahr-Arnt complex then can interact with histone deacetylaces and chromatin remodeling proteins to promote transcription of xenobiotic response elements (XRE)

(Noakes, 2015). These XRE are called so because they include Cyp1a1, Cyp1a2, drug

metabolizing enzymes that can digest foreign agents, producing oxidative stress (Nebert,

Dalton, Okey, & Gonzalez, 2004; Senft et al., 2002). Ahr is activated strongly by dioxin

(agent orange), and the negative health effects of agent orange are attributed to activation of Ahr; mice deficient in Ahr are protected from many effects of dioxin (Fernandez-

Salguero, Hilbert, Rudikoff, Ward, & Gonzalez, 1996). Ahr inhibition has been tested using a potent and highly selective antagonist CH-223191 (S. H. Kim et al., 2006). While toxic xenobiotic agents such as dioxin are well-established agonists of Ahr, KYN was the first endogenous Ahr ligand to be discovered. Endogenous KYN was found to be upregulated in brain tumors, suppressing the normal immunological response thereby promoting tumor growth and malignant progression (Opitz et al., 2011).

Tryptophan and KYN both enter the cell via the same transporter, solute carrier family 7 member 5 (Slc7a5, or Lat1; (Sinclair, Neyens, Ramsay, Taylor, & Cantrell,

2018)) (Figure 6).Once inside the cell KYN binding to Ahr causes translocation to the nucleus (Opitz et al., 2011; Yamamoto et al., 2019). Ahr binding and transcriptional activation by KYN have previously been shown to promote the expression of both TGF beta 1 and indoleamine 2, 3-dioxygenase 1 (IDO1) (Nuti et al., 2014; Vogel, Goth, Dong,

Pessah, & Matsumura, 2008).

Figure 6. KYN-AHR signal transduction pathway. Circulating KYN can enter the cell through the LAT1 transporter and bind to Ahr. Ahr dissociates from HSP90 and translocated to the nucleus. In the Nucleus Ahr binds to ARNT and can bind to XRE to transcribe downstream CYP1A1 and other xenobiotic response element genes.

The induction of IDO1 by Ahr is significant, since IDO1 metabolizes tryptophan to

KYN, thus forming a positive feedback loop. In addition, Ahr has also been observed to positively regulate Slc7a5 expression (Brauze et al., 2017; S. Kim, Dere, Burgoon,

Chang, & Zacharewski, 2009; Tomblin et al., 2016). The increase in Slc7a5 with Ahr activation by KYN presumably facilitates further KYN entry into the cell, representing yet another “feed forward” mechanism of Ahr pathway activation in the setting of inflammation (Figure 7).

While the bulk of literature on IDO1-KYN-Ahr signaling reveals mechanisms by which this pathway is involved in inflammatory responses and oxidative stress in the settings of cancer and dioxin exposure, it is also clear that Ahr can serve a protective function in certain contexts (Dalton, Puga, & Shertzer, 2002). This is most obvious from the phenotype of mice lacking Ahr, which show a number of abnormalities noted above including liver and lung dysfunction. Importantly, Ahr has recently been highlighted as playing a key role in regulating organismal aging and lifespan. One group suggested that Ahr activation promoted aging by increasing vascular stiffness in mice, which was further supported by data in human subjects showing a positive association between Ahr expression and vascular stiffness (Eckers et al., 2016). These authors also documented increased lifespan in C. elegans worms lacking functional Ahr. On the other hand, Bravo-Ferrer and colleagues observed that Ahr expression decreased significantly with age in the mouse brain, and that mice lacking Ahr showed elevated circulating levels of inflammatory cytokines and reduced lifespan (Bravo-Ferrer et al., 2019). Their work is consistent with the data from studies on hematopoietic stem cells by Singh et al. and

Bennett et al. who found that aged Ahr knockout mice showed decreased survival,

Figure 7. Factors regulating Ahr gene expression. The Ahr promoter region contains multiple binding regions for NK kappa B (NF-kB). KYN, a ligand of Ahr, is produces by the degradation of tryptophan by the enzyme IDO1, which is itself induced by inflammatory stimuli and interferon gamma expression. The Ahr itself activates expression of the amino acid transporter SLC7A5, which transports both KYN and tryptophan into the cell. Positive feedback mechanisms are indicated in blue, and include Ahr upregulation of IDO1, further increasing KYN concentrations, and SLC7A5 expression increasing KYN transport to further activate Ahr nuclear translocation.

splenomegaly, and increased circulating white blood cells (Bennett, Singh,

Unnisa, Welle, & Gasiewicz, 2015; Singh et al., 2014; Singh, Garrett, Casado, &

Gasiewicz, 2011). These findings are also in general agreement with earlier research documenting liver fibrosis and cardiac hypertrophy in Ahr null mice (Vasquez et al.,

2003).

These somewhat contradictory findings regarding Ahr function and organismal aging are further complicated by studies showing that, while ligands such as KYN and dioxin may have pathological effects via Ahr, other Ahr ligands have been identified such as quercetin that are well-documented to have anti-aging functions (Table 1) (Dietrich,

2016). Thus, ligand-specific effects may exist such that downstream effectors of Ahr signaling such as cytochrome P450 1a1 enzyme (Cyp1a1) may be activated with particular ligands but other ligands may activate different pathways such as the antioxidant nuclear factor erythroid 2–related factor 2 (Nrf2) (Dietrich, 2016; Larigot,

Juricek, Dairou, & Coumoul, 2018; Nuti et al., 2014). In addition, Ahr ligand specificity and affinity can differ between mice and humans, further complicating observed relationships among Ahr activation, inhibition, and disease processes (Murray, Patterson,

& Perdew, 2014). Ahr can modulate ROS accumulation by activating Cyo1a1, which increases ROS (Senft et al., 2002), but this is balanced by the direct and indirect activation of Nrf2 (Dietrich, 2016).

Table 1 Exogenous and endogenous ligands of the aryl hydrocarbon receptor and their potential impact on age-related diseases. References: 1(Refaey et al., 2017) 2(Kaiser et al., 2019) 3(B. J. Kim et al., 2019) 4(Enoki et al., 2016) 5(Watanabe et al., 2017) 6(Kirkland & Tchkonia, 2017) 7(Ashida, Fukuda, Yamashita, & Kanazawa, 2000) 8(Ciolino & Yeh, 1999) 9(Jin et al., 2018) 10(Ciolino, Daschner, & Yeh, 1998).

III. MATERIALS AND METHODS

3.1 Cell culture

C2C12 cells were obtained from ATCC (ATCC® CRL-1772™) and primary human myoblasts were obtained from Gibco. Both cell lines were cultured in Dulbecco’s modified Eagle’s medium (DMEM) (Gibco, USA) containing 10% fetal bovine serum

(Gibco, USA) and 1% penicillin-streptomycin (Gibco, USA) Myoblasts were seeded in the media and maintained at 37 °C in a 5% CO2 cell incubator (Thermo, USA) until

70%-80% confluence. KYN concentrations of 1µM, and 10µM were chosen based on serum concentrations in found in healthy vs. pathological humans (Pertovaara et al.,

2006). Ahr inhibition was performed using the Ahr antagonist CH-223191 (Sigma

C8124) at 10µM. Ahr inhibition was given 2 hours prior to treatments.

3.2 MTT assay

Myoblast viability after KYN treatment was determined using a MTS assay (Promega

CellTiter 96® AQueous One MTS Cell Proliferation Assay). C2C12 myoblasts were plated in a 96-well plate at initial seeding density of 2500 cells/cm2 or 5000 cells/cm2.

After 24 hours myoblasts (N= 8 per group) were treated with 1xPBS, 5µM KYN, 10µM

KYN or 40µM KYN for 24 hours, and 48 hours. After treatment, myoblasts were washed with PBS 2x, and 20 μl of MTS assay buffer (MTS (CellTiter 96® AQueous One

Solution Reagent, Promega) was added in 100 μl of media. The myoblasts were kept in at

37°C in a humidified, 5% CO2 incubator for 2 hours, then optical density was read at 490

nm using a spectrophotometer. Percent cellular viability was determined by normalizing optical density values to 6 averaged control wells, then multiplying by 100.

3.3 AmplexTM red assay

A fluorometric method was used to measure H2O2 in C2C12 myoblasts (n=6 per treatment), and primary human myoblasts from middle-aged females (n=4 per treatment) treated with KYN using an AmplexTM red assay kit. Once confluent myoblasts were treated with 1xPBS, 1µM KYN, or 10µM KYN for 24 hours. After treatment, media was removed and cells were suspended in sodium phosphate buffer (0.05 m, pH7.4, 100 mL) and plated in triplicate in a flat-bottom 96-well plate. The reaction was started by adding

AmplexTM red reagent, horse-radish peroxidase and p-tyramine. After 30 min incubation in the dark, the production of H2O2 was quantified at 37ºC in a multi-detection microplate fluorescence reader (Syner-gy H1, BioTek Instruments) based on the fluorescence generated at an emission wavelength of 590 nm upon excitation at 545 nm. The specific final fluorescence emission was calculated against a standard curve of H2O2 incubated simultaneously.

3.4 Total Antioxidant capacity assay

The Total antioxidant capacity (TAC) was measured with an Antioxidant Assay kit

(Cayman Chemicals). In this assay .C2C12 myoblasts were treated with VEH or 10µM

KYN in triplicate (n=6 each) for 24 hours. Then cells were exposed to 2.2΄-azino-di 3- ethylbenthiazoline- sulfonate (ABTS) and then methmyoglobin. ABTS was then oxidized

to ABTS+ by methmyoglobin. Trolox, an analogue of vitamin E, was used to create a standard curve, and TAC of myoblasts were plotted against the standard curve.

3.5 Animal Experimental Design

All aspects of the animal research were conducted in accordance with the guidelines set by the Augusta University Institutional Animal Care and Use Committee (AU-IACUC) under an AU-IACUC–approved Animal Use Protocol. For KYN and 1-MT treatment studies 6-8 month-old (young adult) and 22-24-month-old (old) C57BL/6 mice were obtained from the aged rodent colony at the National Institute of Aging (NIA, Bethesda,

MD, USA). For Ahr studies, young adult female Ahr whole body knockout mice were obtained from (Taconic #9166). For KYN treatment and Ahr-KO studies mice received daily intraperitoneal (I.P.) injections of vehicle (phosphate-buffered saline [PBS]) or L-

KYN (Sigma; #K8625) at 10 mg/kg body weight for 4 weeks (n = 10 per group). For 1-

MT studies, old, C57BL/6 mice were used. Mice were split into 3 groups (n = 20 per group): Old VEH (sterile H2O, 1% DMSO, 0.20 mL injection), old low 1-MT (10mg/kg

1-MT, 0.20mL injection), and old high 1-MT (100mg/kg 1-MT 0.20 injection). No acute adverse effects were detected with injected KYN of 1-MT. KYN doses were chosen based on previous work in bone (Refaey et al., 2017). 1-MT was dissolved in 1% DMSO and rotated on low heat for 3hrs to dissolve. 1-MT doses were chosen based on toxicology work on 1-MT in rats and dogs (Jia et al., 2008). Mice were euthanized using

CO2 overdose followed by thoracotomy according to AU-IACUC–approved animal protocols. Right quadriceps were fixed in 10% formalin and stored in 70% ethanol for paraffin embedding and histology. Left quadriceps were frozen in liquid nitrogen for

protein and gene expression analysis, and right tibialis anterior were used for AmplexTM red assay immediately.

3.6 Histological staining

Quadricep femoris muscles were fixed in 10% buffered formalin, then paraffin embedded, and sectioned at 6-8µm.Sections were deparaffinized, rehydrated, and non- specific binding was blocked by 0.3% H2O2 in TBS. Sections were then incubated overnight at room temperature with a rabbit polyclonal anti-Laminin (dilution 1:1000,

Sigma-Aldrich, USA) and Rabbit-anti-4-HNE (Alpha diagnostic HNE11-S, dilution

1:50), and ChromPure Bovine IgG (Jackson, 001-000-003, dilution 1:50). The Laminin sections were washed with phosphate-buffered saline (PBS, pH 7.4), and incubated 1hr at room temperature with a goat anti-Rabbit Alex fluor 488 conjugated secondary

(Invitrogen, A11008). 4-HNE and igG sections were incubated with a polyvalent secondary antibody, followed by streptavidin solution (Abcam ab93697). 4HNE and IgG were visualized using DAB Liquid Chromogen Solution (Sigma D3939), and counterstained with hematoxylin (Fisher 245-677). Muscle fiber size was determined by creating a grid on using ImageJ (Fiji) software and measuring one muscle fiber in each voxel. One section was selected at random from each mouse (N=10 per group), and 10 muscle fibers per section were measured, then an average fiber diameter per mouse was taken. Percentage of 4HNE-positive staining was measured using Photoshop. All measurements were performed by an investigator blinded to group assignment.

Frozen sections were used for Oil red O. Slides were incubated at room temperature for 10 minutes. Then fixed in 10% buffered formalin for 1 hour at room

temperature. Next, slides were washed with deionized water three times, and stained with

Oil red O solution (Sigma-Aldrich) for 1 hour at room temperature. Finally, slides were rinsed in deionized water, washed in running tap water for five minutes, and mounted microscopy using fluoroshield mounting medium with DAPI (Abcam). Confocal images were acquired using a Zeiss LSM 780 confocal microscope (Carl Zeiss Microscopy,

Peabody, MA, USA) fitted with a 63X/1.2NA water immersion lens. The Zen 2012 software package (Carl Zeiss) was used for image capture.

3.7 Proteomics

In order to identify protein candidates, proteomics were run on three quadriceps muscle samples per each group: young KYN, young VEH, old 1-MT (high dose), old VEH.

Quadriceps muscles were homogenized and protein was run at the Augusta University proteomics core using an Orbitrap Fusion™ Tribrid™ mass spectrometer. All proteins with a two-fold or greater difference were chosen from each group and the protein candidate that was differentially expressed in the separate treatment groups identified.

3.8 Western Blots

For western blots, human myoblasts were lysed in radioimmununoprecipitation assay

(RIPA) buffer (Tecnova) containing 1% protease inhibitor cocktail (Sigma). Protein concentrations were obtained using a BCA Protein Assay Kit (Sigma). Protein was run in

SDS–polyacrylamide gels and transferred using electrophoresis onto a nitrocellulose membrane (Bio-Rad). Blots were incubated with a rabbit polyclonal anti-VLCAD

(ab155138) overnight at 4°C. After washing with 1× PBS and blocking with 5% milk in

1× PBS, blots were incubated with HRP-conjugated anti-rabbit secondary antibody

(Santa Cruz Biotechnology) for 1 h, followed by developing with the ECL Plus Western

Blotting Detection System (GE Healthcare). Chemiluminescence signals were captured on autoradiographic blue films (Bioexpress). Films were scanned and the densitometric values for the proteins of interest were corrected using β-actin with Image J Software

(NIH).

3.9 PCR array plates

Quadriceps muscles from young VEH mice, and three young KYN (n=3/group) were sonicated, and RNA was isolated using Trizol reagent (Invitrogen) according to manufacturer’s instructions. Total RNA was purified using an RNeasy Mini Kit (Qiagen).

1µg of total RNA was then reverse transcribed using the First Strand Synthesis Kit

(Qiagen) and subsequently loaded into Skeletal Muscle Myogenesis and Myopathy RT2

Profiler PCR Arrays (Qiagen). PCR was run at the following conditions: 10 min at 95°C,

45 cycles of 15s at 95°C, and 1 min at 60°C. Fold change was calculated by determining the ratio of mRNA levels to control values using the ΔCt method (2−ΔΔCt). All data were normalized to an average of six housekeeping genes: Actb, B2m, Gapdh, Gusb,

Hsp90ab1, and Mgdc. PCR conditions are the following: hold for 10 min at 95°C, followed by 45 cycles of 15 s at 95°C and 60 s at 60°C.

3.10 In vivo muscle function testing

Muscle peak twitch was measured using a whole mouse testing apparatus (1300A,

Aurora Scientific Inc, Aurora, ON, Canada) and a force transducer (Aurora Scientific Inc,

Canada) (Figure 6.). This apparatus provides torque measurements in (milliNm) of tetanic contraction while the animal is alive and with normal vasculature, innervation, and muscle orientation. Animals were maintained under anesthesia with a CO2 and oxygen breathing cone. Animals were placed on a 37°C platform and the right hind foot was stabilized to a foot lever with cloth tape at 20° of plantar flexion. Needle electrodes were placed under the skin below the knee to stimulate the peroneal nerve. From torque measurements, the specific muscle force is obtained by normalizing the absolute force values (milliN) to the animal’s body weight. Peak muscle twitch values were selected from each animal.

3.11 Ex Vivo muscle testing

For ex vivo muscle testing, an outbred strain of mice were used. Untreated CD-1 female mice were sacrificed, and fresh extensor digitorum longus (EDL were dissected and immediately placed in Ringer’s solution with or without 100µM or 200µM kyn. A surgical suture was tied to each tendon of an EDL, and around a hook in a bath of

Ringer’s solution using the ex vivo muscle testing apparatus (1300A, Aurora Scientific

Inc, Aurora, ON, Canada) and a force transducer (Aurora Scientific Inc, Canada) (Figure

7.). Of note, the amplitude was set to 100x higher than the in vivo system. Muscle peak twitch was then recorded. Tetanic contractions (350ms train) at 10 to 250 Hz were elicited to obtain a force-frequency curve, with a 2-minute rest between each contraction.

Results of single stimulations were collected in torque (milliNm).

Figure 8 Aurora functional muscle testing system: A) In vivo mice are anethsitized with a breathing cone. The hind leg is stabilized by a knee stabilizer, and by taping the foot to a foot plate on the force detector. Two electrodes are inserted into the anterior portion of the hind leg to stimulate the peroneal nerve and activate the tibialis anterior. (B.) Ex vivo, the EDL is suspended in a bath of ringer’s solution and tendons are sutured to a stabilized hook on one side, and a force detector on the other side. Electrodes are suspended in the bath to stimulate the muscle.

3.12 Open field activity

Young and old mice treated with KYN and 1-MT, respectively were tested in an open apparatus measuring 40cm x 30cm, with walls 15cm high. The walls and floor of the apparatus were white. A GoPro camera was placed above the apparatus. Each mouse was allowed to acclimate to the apparatus for 5 minutes, then removed from the apparatus.

Activity data on each mouse was measured three times for 5 minutes with at least 30 minutes in-between each test. Data was averaged for each mouse across each day. All mice were tested before treatment, treated with KYN or 1-MT for 30 days, and tested after treatment. Old mice treated with 1-MT were also tested 30 days after the last day of treatment. Data was analyzed using Kinovea software to track distance traveled.

3.13. Cytokine array panels

Tibialis anterior muscles, and serum were collected from young male and female mice treated with 10mg/kg of KYN for 30 days or VEH. Skeletal muscle was homogenized in cell lysis buffer (Bio-Plex kit, BioRad), spun for 20 minutes at 10k rpm, then the supernatant was collected and frozen. Serum was incubated at room temperature for 30 minutes, then centrifuged for 20 minutes at 10k rpm, the supernatant was collected and frozen. Cytokine levels in serum or skeletal muscle lysate into the media was quantified using the LEGENDplex™ (mouse inflammation panel) bead-based immunoassay

(BioLegend, San Diego, CA, USA) according to the manufacturer’s instructions. Data were acquired using four-color BD FACSCalibur (BD Biosciences, San Jose, CA, USA) and analyzed using the LEGENDplex Data Analysis software (BioLegend). (Sanz et al.,

2017)

3.14 Statistical Analysis

For all experiments with more than 2 groups, an ANOVA and a post hoc Tukey’s HSD test for differences between means was used. Results for all experiments with 2 groups except the functional muscle testing were determined using a 2-sample independent t-test to compare differences between means of groups. Functional muscle testing results were determined using a repeated measures t-test. Biostatistician Maribeth Johnson was consulted to verify datasets.

IV RESULTS

4.1 KYN treatment increases ROS levels in C2C12 mouse and primary human myoblasts.

In order to determine the effect of KYN on ROS production in myoblasts, H2O2 levels were measured from C2C12 mouse myoblasts that were treated with KYN for two hours.

H2O2 levels were measured using an Amplex™ red assay. The Amplex™ red reagent is cleaved by H2O2 to produce resorufin which fluoresces at approximately 560 nm. H2O2 levels are measured by the amount of signal emitted after a 30 minute incubation with

Amplex™ red reagent. H2O2 was increased two-fold with KYN treatment at only 1μm

(Figure 9A) The higher dose (10μm) produced no further increase beyond the lower 1μm dose. Primary human myoblasts were also treated with KYN in the same manner. The higher dose was significantly higher than VEH (Figure 9B).

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Figure 9. ROS is increased in mouse and human myoblasts with KYN compared to VEH treatment. (A.) H2O2 levels are increased (P<.01) in mouse C2C12 myoblasts after 24 hours of KYN treatment (n=6) at 1µM and 10µM compared to VEH treatment (n=6). (B.) H2O2 levels are increased (P<.05) in Human primary myoblasts after 24 hours of KYN treatment (n=4) at l0µM compared to VEH treatment. Data are presented as mean ± 95% confidence interval. * P<.05, ** P<.01, *** P<.001.

4.2 Ahr inhibition did not rescue KYN effects in vitro.

KYN is a ligand for the Ahr (Mezrich et al., 2010). To determine if the changes demonstrated in skeletal muscle by KYN is due to KYN-activation of Ahr, KYN was tested in C2C12 myoblasts for 24 hours with and without the highly selective Ahr inhibitor CH-223191. An AmplexTM red assay (Figure 10A) showed that all treatments had significantly higher H2O2 than VEH control (ANOVA; p<.001) KYN treated with

CH-223191 had more H2O2 than KYN alone (P<0.05). An MTT assay was performed to measure cellular viability (Figure 10B) Cellular viability was unchanged in KYN compared to VEH with and without CH-223191. Downstream Ahr product Cyp1a1 was measured in C2C12 myoblasts treated with KYN with and without CH-223191 (Figure

10C), and no differences were seen with or without CH-223191. A total antioxidant assay was performed on KYN with or without CH-223191. There was no difference in antioxidant capacity between VEH and KYN alone. KYN treated with CH-223191 had higher antioxidant capacity than KYN alone (P<0.05). C2C12 myoblasts treated with

CH-223191 alone had a higher antioxidant capacity than VEH alone (P<.05) (Figure

10D).

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TM Figure 10. Ahr inhibition in vitro. Amplex red measuring H2O2 in C2C12 cells treated with KYN for 24 hours with or without CH-223191 (CH) (A.), MTT assay measuring cellular viability (B.). CYP1A1 assay measuring CYP1A1 expression (C.) Total antioxidant capacity (D.). Data are presented as mean ± 95% confidence interval. * P<.05, ** P<.01, *** P<.001.

4.3 KYN treatment produces an aged-muscle phenotype in young female mice.

To test the effect of increased KYN on skeletal muscle in vivo young and old C57/BL6 mice were given intraperitoneal injections with KYN (10 μg/kg Sigma k3750) or VEH

(1xPBS) daily for 4 weeks. Mice were sacrificed and quadriceps femoris muscles were removed. Quadriceps were weighed and used for histological sections and proteomics.

Histological sections were further stained with either trichrome to measure fiber size, or

4HNE, a marker for lipid peroxidation, which is found to increase with increased ROS production. Quadriceps weight relative to body weight was reduced (10% NS) in young treated muscle compared to control (Figure 11A) and muscle fiber size was significantly lower (P<.01) in young mice treated with KYN compared to VEH (Figures 11B and C).

Positive 4HNE signal was significantly increased (P<.05) in young mice treated with

KYN compared to young controls (Figures 12A and B). Skeletal muscle fibrosis levels were measured from young mice treated with KYN using trichrome staining. Blue staining was normalized to red and blue (total) staining. There was no change in fibrosis with KYN treatment (Not pictured.

Figure 11 Muscle loss is increased in young mice treated with KYN. After 4 weeks of I.P. injections of KYN or VEH (n=10 ea. group) average quadriceps wt. was reduced by 10% in young mice (A.), and fiber size was significantly reduced (ANOVA, P<0.01) in young mice treated with KYN, or Old mice with or without KYN (B. and C.) Figure C. demonstrates Cross sections of quadriceps muscle fibers stained with laminin to visualize fiber size. Representative fibers are indicated with *. Data are presented as mean ± 95% confidence interval. * P<.05, ** P<.01, *** P<.001

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Figure 12. Skeletal muscle lipid peroxidation is increased in young mice with KYN treatment. 4HNE staining is significantly increased (ANOVA, P<.05) in young KYN and Old with and without KYN compared to Young VEH (A. and B.) (Scale bar 100µm) Data are presented as mean ± 95% confidence interval. * P<.05, ** P<.01, *** P<.001.

4.4 Ahr knockout weakly attenuated KYN effects in female mice.

To further explore the relationship between Ahr and KYN in skeletal muscle, female global Ahr KO mice (6-8 months old) were treated with KYN (10 mg/kg) or VEH

(1xPBS) by intraperitoneal injections daily for 4 weeks. Quadriceps muscle weight was significantly lower (P<.0001) in Ahr KO mice treated with KYN compared to VEH treatments (Figure 13A). Muscle fiber size was not significantly different in Ahr KO mice with KYN treatment compared to VEH (Figure 13B). There was not a significant difference in H2O2 in Ahr KO mice compared to VEH (Figure 13C). 4HNE staining was not significantly different in Ahr KO mice with KYN treatment compared to VEH

(Figure 13D). A two-factor ANOVA was run with genotype (WT, Ahr KO) and treatment

(VEH, KYN) as the two factors to determine whether absence of the Ahr would significantly impact lipid peroxidation assessed by 4HNE staining. We found no significant genotype*treatment interaction for either muscle fiber size (F=0.10, P=0.75) or 4HNE staining intensity (F=0.54, P=0.47), indicating that loss of the Ahr receptor did not significantly alter the response of muscle to KYN treatment. However, the trends between Figures 11 and 12 and Figure 13 suggest there may be a weak protective effect of Ahr knockout in female mice treated with KYN.

Figure 13. Ahr knockout in female mice. Global Ahr knockout mice treated with KYN for 30 days (n=10). KYN treatment of Ahr knockout mice significantly decreased quadriceps mass (A.). Fiber size of quadriceps muscles from Ahr knockout mice decreased slightly but not significantly with KYN treatment (B.). H2O2 from quadriceps of Ahr knockout mice treated with KYN did not change with KYN treatment compared to VEH (C.). 4HNE staining of quadriceps muscles did not change with KYN treatment. Data are presented as mean ± 95% confidence interval. * P<.05, ** P<.01, *** P<.001.

4.5 1-MT treatment decreased oxidative stress and attenuates muscle atrophy in old female mice.

Old female C57/BL6 mice were given intraperitoneal injections with a low dose

(10µg/kg), high dose (100µg/kg) or VEH (1xPBS) daily for 4 weeks to examine the levels of oxidative stress and muscle atrophy induced by KYN production. Amplex™ red assays were performed on quadriceps femoris (Figure 14A). H2O2 measured by

Amplex™ red assay showed significantly less (P<.05) H2O2 in old mice treated with

100mg/kg 1-MT compared to VEH controls. Quadriceps femoris muscles were weighed and processed for histological sections to test the effect of decreased KYN on skeletal muscle. Skeletal muscle plasma membranes were stained with laminin to measure fiber size. Quadriceps weight relative to body weight was significantly increased in mice treated with a high dose of 1-MT compared to vehicle treated mice (Figure 14B). Muscle fiber size was significantly increased compared to vehicle treated mice in both treatment groups (Figures 14C and D). Fibrosis levels were measured with trichrome staining in old mice treated with 1-MT. Blue staining was normalized to red and blue (total) staining.

There was no change in fibrosis with 1-MT treatment (Not pictured).

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b 1500

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a 500

u Q 0 g g H /k /k E g g m m V 0 0 ld 1 0 1 O T T -M 1 -M 1 ld d O l O Figure 14. Skeletal muscle H2O2 and morphology changes after 1-MT treatment in aged female mice. H2O2 (A.) levels were significantly decreased in mouse quadriceps muscles after 4 weeks of 100mg/kg 1-MT treatment. (B) Quadriceps muscle mass was significantly increased in mice treated with 1-MT compared to VEH. Muscle fiber size was significantly increased in mice treated with high dose 1-MT compared to VEH (C and D), visualized with laminin staining. Representative muscle fibers are marked with stars. (Scale bar 100µm) Data are presented as mean ± 95% confidence interval. * P<.05, ** P<.01, *** P<.001.

4.6 Peak muscle strength is attenuated with KYN treatment.

To test the strength of mice with increased KYN, or inhibition of KYN production, an

Aurora functional muscle testing system was used. Young male and female mice treated with KYN, or old male and female mice treated with 1-MT were anesthetized, and electrodes were inserted to stimulate the peroneal nerve, and measure the strength of the anterior muscles of the leg; tibialis anterior, extensor digitorum longus and extensor hallucis longus. Functionally, stimulation of the peroneal nerve, dorsiflexes the foot taped to a foot plate that measures force pulled. Functional, in vivo assessment of muscle contractile force showed that young KYN-treated male mice lost significant peak muscle strength after 4 weeks of treatment (Figure 15A) (P<.05). Muscle peak contractile force was significantly higher in old male mice treated with 100mg/kg 1-MT after 4 weeks of treatment than before treatment (Figure 15B) (P<.05). Young female mice treated with 4 weeks of KYN did not change muscle strength (Figure 15C), but old female mice treated with 4 weeks of 1-MT showed triple the peak muscle strength compared to old female controls (Figure 15D) (P<.01). Ex vivo Muscle testing was performed to test muscle strength removed from confounding factors of nerve stimulation and vascular supply. The extensor digitorum longus was removed from untreated mice, and immediately hung in a bath of Ringer’s solution supplemented with 100µm or 200µm of KYN. The bath was stimulated to induce muscle contraction. Results show that EDLs dissected from untreated mice showed no significant difference from stimulation in Ringer’s solution supplemented with 100µm or 200µm of KYN (Not pictured).

Figure 15. Peak muscle strength was decreased with KYN treatment in young mice and increased with 1- MT treatment in old mice. Muscle peak contraction significantly lowered in young mice treated with KYN for 4 weeks after treatment than before treatment (P<.05). In old mice treated with 1-MT for 4 weeks, muscle peak contraction was significantly higher after treatment than before (P<.05). Data are presented as mean ± 95% confidence interval. * P<.05, ** P<.01, *** P<.001.

4.7 PCR arrays reveal structural muscle proteins are decreased with KYN treatment.

To explore the mechanism by which KYN effects skeletal muscle, PCR arrays on genes known to affect muscle strength and size were performed. PCR arrays were run on quadriceps femoris from three young VEH and young KYN female mice (Figure 16).

Skeletal Muscle: Myogenesis & Myopathy (PAMM-099) PCR arrays (SABiosciences

Corp.) were used to profile muscle genes important for differentiation, structure, and disease. All significant changes are shown in figure 15. mRNA coding for structural proteins Alpha-actinin-3 (Actn3), Caveolin 1 (Cav1), myosin heavy chain 1 (Myh1), myosin heavy chain 2 (Myh2), Troponin 3, (Tnnt3) and titan (Ttn) were significantly decreased with KYN treatment. Calpain 2 catalytic subunit (Capn2), a calcium activated protease was also significantly decreased with KYN treatment. Individual real time PCRs were run on quadriceps femoris homogenates from young VEH and Young KYN treated female mice on genes known to build or break down skeletal muscle: Murf1, Mafbx, P53,

Mtorc11, and Ps6k1, compared to controls 18s and Gapdh. All revealed no significant

changes after KYN treatment.

Figure 16. Structural protein mRNA was decreased with KYN treatment in young female mice: mRNA for Actn3, Capn2, Cav1, Myh1, Myh2, Tnnt3, and Ttn were significantly (P<.05) decreased with four weeks of KYN treatment in the quadriceps femoris of young female mice (n=3 per group) Data are presented as mean ± SEM. * P<.05, ** P<.01, *** P<.001

4.8 KYN did not change intracellular lipid accumulation.

Sarcopenia is associated with lipid accumulation in skeletal muscle (Ryall et al., 2008).

To test the amount of lipid accumulation with increases of KYN, Frozen sections were taken of quadriceps femoris from young male and female mice treated with KYN or

VEH. Intracellular lipid was stained with oil red O, a fat-soluble dye to stain neutral triglycerides and lipids. There was no difference observed in intracellular lipid content in

VEH compared to KYN treatment for male or female groups (Figures 17A and B). There was more lipid accumulation in females compared to males (P<.01).

17A Female VEH Female KYN

20x

FemaleMale VEH VEH FemaleMale KYN KYN

y t

B. i ✱✱ s

n 1.0

g 0.8

i a

t 0.6

R 0.4

a 0.2

e M 0.0 H N H N E Y E Y V K V K e e l l le le a a a a m m e e M M F F

Figure 17. : Intracellular lipid content did not change with KYN treatment in male or female mouse quadriceps: (A) Oil red O (red) in skeletal muscle quadriceps of male and female mice Nucleus stained blue with dapi. Quantified in (B.) Data are presented as mean ± 95% confidence interval. * P<.05, ** P<.01, *** P<.001.

4.9 Fiber type proteins are reduced with KYN treatment.

Skeletal muscle is composed of myofibers that are primarily slow twitch, type 1, and use aerobic, oxidative pathway for energy, or fast twitch, type 2, that use anerobic glycolytic pathway for energy. The distribution of fiber types is known to change with age. To test the effect of KYN on fiber type distribution, human myotubes in vitro were treated with

10µm KYN for 24 hours, and protein was collected to run western blots on myosins expressed primarily in fast or slow twitch fibers. Fast twitch myofibers were measured with myosin heavy chain II (Myh II), and slow twitch myofibers were measured with myosin heavy chain I (Myh I). Both fast twitch and slow twitch myosin content was significantly reduced with KYN treatment (P<0.05) (Figures 18A and B).

18A. VEH KYN 10 µM

MYH II

MYH I

Gapdh

MYH II MYH I B. C. ✱ ✱

2.0 2.0

1.5 1.5

e e

g g

n n

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h h

C C

1.0 d 1.0

l l

o o

F F

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0.0 0.0 VEH KYN VEH KYN 10M 10M

Figure 18. Fiber type changes in human myotubes treated with KYN. (A.) Western blots for Myh II (fast twitch) and Myh I (Slow twitch) on human myotubes treated with 10µm KYN. Both Myh I (B.) and Myh II (C.) were significantly reduced in human myotubes treated with 10µm KYN compared to VEH controls. All results were normalized to Gapdh. Data are presented as mean ± 95% confidence interval. * P<.05, ** P<.01, *** P<.001.

4.10 Proteomic analysis of KYN treated and 1-MT treated skeletal muscle reveals differential expression of very long chain acyl-coa dehydrogenase.

To determine protein expression changes in young skeletal muscle with increases in KYN, or aged skeletal muscle with inhibition of KYN production, proteomic analysis was run on quadriceps femoris tissue from female young mice treated with KYN, and female aged mice treated with 1-MT. Both groups were compared to age matched VEH treated controls.

Proteins that were decreased with KYN and increased with 1-MT or increased with KYN and decreased with 1-MT were selected (Figure 19A). The top protein candidate that was differentially expressed with KYN and 1-MT compared to age-matched VEH controls was mitochondrial very long chain acyl-coa dehydrogenase, (Mvlcad) (Figure 19B). Mvlcad was significantly increased at 1μM and 10μM KYN treatment of primary human myoblasts compared to VEH controls (Figures 19C and D).

Figure 19. Lipid peroxidation enzymes are differentially expressed with KYN or 1-MT treatment. (A) Proteins that were increased with KYN treatment and decreased with 1-MT treatment in female mice compared to age-matched controls. (B) Proteomics run on quadriceps muscle homogenates from young mice treated with KYN or aged mice treated with 1-MT showed that Mvlcad was significantly increased with KYN treatment in young mice. (C) Representative western blots from primary human cells that showed a significant increase in Mvlcad with KYN treatment. (D) Mvlcad western blot results were normalized to Gapdh controls and quantified. Data are presented as mean ± 95% confidence intervals. * P<.05, ** P<.01, *** P<.001.

4.11 Old female mice treated with 1-MT reduced activity 30 days after 1-MT treatment.

Open field activity tests were run on young mice treated with KYN for 30 days (Figure

20A) old mice treated with 1-MT for 30 days, and old mice 30 days after the last 1-MT treatment (Figures 20B and C) to determine if strength changes in skeletal muscle might result from activity changes. Neither male nor female mice treated with KYN changed activity levels after KYN treatment. In addition, there were no significant differences in old females immediately after 1-MT treatment, but after 30days, female mice moved significantly less than pretreatment (P<.01), and less than immediately after treatment

(P<.05). Males however, demonstrated a slight decrease in activity 30 days after 1-MT treatment, but no time points were significant.

20A . Males P=0.34

0.8

t w

0.6

/ e

c 0.4

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0.0 Females t t Males B n C en ✱✱ e m tm t P=0.17 . 3000 a e.a 4000 re tr ✱ t st

re o ) ) p p

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D D

0 0 t se t T se n T o n o e M d e M d m - m - t 1 T t 1 T a d a d re 0 -M re 0 -M t 3 1 t 3 1 - r t - r t re e s re e s P ft la P ft la A A er er ft ft a a d d 0 0 3 3

Figure 20. Open field activity after KYN or 1-MT treatment in young and old, male and female mice. (A.) Young male and female mice did not differ in open field activity data after four weeks KYN treatment. (B.) Old female mice treated with 1-MT did not differ in open field activity immediately after the last treatment, however 30 days after the last treatment, activity decreased significantly from pretreatment (P<.01), and from posttreatment (P<.05). (C.) Old male mice treated with 1-MT showed some decrease in open field activity 30 days after the last treatment, but differences were not significant. Data are presented as mean ± 95% confidence intervals. * P<.05, ** P<.01, *** P<.001.

4.12 Muscle and serum cytokines were altered with 30 days of KYN treatment.

Inflammatory cytokine arrays were run on tibialis anterior lysate, and serum taken from male and female mice treated with 30 days of KYN or VEH. Il-1α was increased in female muscle treated with KYN compared to VEH (P<0.05) (Figure 21A) with no difference between male groups. Female serum had decreased Il-1αcompared to VEH

(P<0.05) (Figure 21B) with no difference between male groups. Il-27p70 was decreased in male and female serum compared to VEHs (P<0.001) (Figure 21C) Il-10 is decreased in female serum (P<0.050 (Figure 21D), and Il-17α is decreased in female serum (P<.01) with no change in male serum (Figure 21E).

21A B. Muscle lysate Serum lysate ✱ 15 ✱ 10

8 l

l 10 m

m 6

a 1

1 4

l I I 5 2

0 0

H g H g H g H g E /k E /k E /k E /k V g V g V g V g F m M m F m M m 0 0 0 0 1 1 1 1 N N N N Y Y Y Y K K K K F M F M

C. Serum lysate D. Serum lysate ✱✱ ✱

800 ✱✱✱✱ 800

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600 l 600

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- l

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0 0 g g g g H k H k H k H k E / E / E / E / V g V g V g V g m m m m F 0 M 0 F 0 M 0 1 1 1 1 N N N N Y Y Y Y K K K K F M F M

E Serum lysate ✱✱ . 200

l 150

a 100

- l I 50

0 g g H /k H /k E g E g V m V m F 0 M 0 1 1 N N Y Y K K F M

Figure 21. Inflammatory cytokines changed with KYN treatment. Il-1α is increased in female muscle treated with KYN compared to VEH (P<0.05) (A), and decreased in female serum compared to VEH (P<0.05) (B.) Il-27p70 is decreased in male and female serum compared to VEHs (P<0.001) (C.) Il-10 is decreased in female serum (P<0.050 (D.), and Il-17α is decreased in female serum (P<.01). Data are presented as mean ± 95% confidence intervals. * P<.05, ** P<.01, *** P<.001.

V. DISCUSSION

The cellular and molecular processes leading to sarcopenia are incompletely understood.

Several factors may contribute to loss of muscle mass and strength with age including lack of physical activity, dietary protein deficiency, circulating inflammatory cytokines, and oxidative stress. Skeletal muscle has the ability to activate antioxidant proteins to quickly repair exercise-induced oxidative damage; however, these mechanisms are attenuated with age, causing an imbalance in ROS (Vasilaki, McArdle, Iwanejko, &

McArdle, 2006). Accumulation of ROS has in particular been suggested to induce age- related declines in muscle, however the mechanism leading to increases in ROS in skeletal muscle have not been clearly defined (F. L. Muller, Lustgarten, Jang,

Richardson, & Van Remmen, 2007; N. T. Smith, Soriano-Arroquia, Goljanek-Whysall,

Jackson, & McDonagh, 2018). We addressed this knowledge gap by examining the cellular and molecular mechanisms underlying the age-associated accumulation of reactive oxygen species. KYN is a circulating tryptophan metabolite that increases with age and is correlated with frailty and increased mortality in older adults (Pertovaara et al.,

2006; Valdiglesias et al., 2018). Elevated circulating levels of KYN are also found to be associated with osteoporosis and Alzheimer’s disease (Bandres et al., 2000; Braidy,

Guillemin, Mansour, Chan-Ling, & Grant, 2011; Srivastava, 2016). In the presence of

IFNγ, IDO converts tryptophan to KYN and IFNγ and IDO activity have both been shown to increase with age (Badawy, 2017; Dai & Zhu, 2010; Mezrich et al., 2010).

As IDO activity is increased, the essential amino acid tryptophan is depleted from the tissue microenvironment and its metabolism is directed away from serotonin synthesis and toward the KYN pathway (Badawy, 2017; Dai & Zhu, 2010). Our study

addressed the hypothesis that increased KYN with age contributes muscle atrophy and oxidative stress. In culture, mouse (C2C12) myoblasts and primary human myoblasts were treated with 1 µM, and 10µM KYN for 24 hours, then H2O2 levels were measured

TM using a fluorometric Amplex red assay. KYN significantly increased H2O2 levels at both doses in C2C12 myoblasts, and at 10µM in human myoblasts. We further speculate the oxidative effects of KYN may be due to downstream breakdown and metabolites of

KYN and these metabolites may function as free radical generators that contribute to cellular ROS (Laviano et al., 2007).

We found that KYN did not decrease quadriceps weight in young mice but did decrease muscle fiber size. This discrepancy could be due to experimental design in weighing whole muscle, vs. sectioned muscle fibers. PCR array data identified several structural muscle proteins that were downregulated with KYN treatment such as Myh,

Myh2, Tnnt3 and Ttn suggesting that reduction in fiber size with KYN treatment may be due to a loss of protein anabolism rather than in increase in protein catabolism. Western blot data showed KYN decreased Myh I and Myh II. These results are consistent with those of several groups that have shown protein anabolism is impaired with aging

(Atherton et al., 2016; Cuthbertson et al., 2005; Moore et al., 2015; Wall et al., 2015). We observed an increase in the oxidative stress markers H2O2 and 4HNE in young mice treated with KYN. We noted that the levels of these oxidative stress markers were similar to those of aged mice and did not continue to increase with further KYN treatment in old mice. We speculate that KYN may reach a threshold, such that additional exogenous

KYN in animals that already have high KYN levels may yield no effects (KYN resistance), but further work to test this hypothesis is needed. We also demonstrated that

inhibition of IDO helped to preserve muscle mass and function in aged mice, further suggesting that the KYN pathway plays an important role in muscle health.

The loss of skeletal muscle size with KYN has been outlined here, but sarcopenia is defined by both the loss of skeletal muscle mass and strength (Santilli et al.,

2014). To test the strength of skeletal muscle we used an Aurora functional muscle testing system. Classically, skeletal muscle strength is tested by hang time and grip strength, but these tests depend on a behavioral component that can make it difficult to demonstrate repeatable results (Martinez-Huenchullan et al., 2017). The Aurora functional muscle testing system uses electrodes to stimulate a nerve and activate the subsequent muscle while the animal is anesthetized. Stimulations were administered in 2 minute intervals to the peroneal nerve to stimulate the tibialis anterior muscle. The foot of the mouse is firmly secured on a lever, and the amount of force exerted on the lever in dorsiflexion is recorded. Peak strength was recorded in both male and female mice before treatment, and after 30 days of treatment. Young male mice lost significant peak strength after 30 days of KYN treatment, and old male and female mice gained significant peak strength after 30 days of 1-MT treatment. Of note, female mice treated with KYN for 30 days did not lose strength. Whether these results are due to a protective effect from female hormonal regulation and cycling, or if fiber size loss is not directly affecting female peak strength remains to be determined.

To further dissect physiologically how KYN is affecting muscle strength, ex vivo muscle testing using the Aurora functional muscle testing system was utilized. In this experiment, the EDL was chosen to be a small enough muscle for the testing system, with sufficiently long tendons to suture to the machine. The EDL was placed in a bath of

ringer’s solution with and without KYN and simulated through electrodes placed directly in the bath. At very high concentrations of 100µM, and 200µM KYN, there was no difference in EDL force. CD-1 mice were used as they are outbred and more robust than

C57bl/6 mice. In order to validate these findings, the experiment should be repeated with skeletal muscle from C57bl/6 mice.

Open field activity tests were performed on young and old male and female mice to determine the rate of KYN-induced activity changes in altered muscle fiber size. Open field activity tests are a relatively non-invasive way to record movement and anxiety in a reproducible manner (Seibenhener & Wooten, 2015). Young mice will move significantly more than old mice in an open field test, and females tend to move more than males (Tatem et al., 2014). Open field activity tests were performed on mice before treatment, after treatment, and old mice were given a 30 day rest period, and tested again.

Young mice treated with KYN did not change activity levels. Old female mice exhibited less activity after 30 days of treatment with 1-MT, and even less after a rest period. Old male mice did not significantly change activity but lost some activity over the course of the experiment. The loss of activity with age is well documented in male and female mice, as well as some loss of activity with repeated testing (L. Wang, Tang, Howell,

Ruiz, & Spurney, 2012). It is possible the mice could lose activity levels faster without 1-

MT treatment, but these results do suggest the effect of KYN in skeletal muscle is not due to decreased activity, or due to increased anxiety levels. Further experiments measuring 24 hour cage activity should be performed to account for changing the environment of the mice, and testing them during the day, despite the nocturnal nature of mice.

KYN is a ligand of the Ahr and is involved in immunosuppression (Seok et al., 2018). Previous work on skin and vascular aging have suggested a potential for

Ahr activation in the aging of various tissue types (Eckers et al., 2016; Qiao et al., 2017).

Ahr is a xenobiotic drug response element that acts as a transcription factor and once activated it stimulates expression of Cyp1a1, which can further increase oxidative stress

(Fukunaga, Probst, Reisz-Porszasz, & Hankinson, 1995; Seok et al., 2018). We found that Ahr inhibition did not protect muscle cells from the detrimental effects of KYN treatment. Surprisingly, when myoblasts were treated with KYN and an Ahr small molecule inhibitor CH-223191, there were significantly higher levels of H2O2 than with

KYN alone. CH-223191 is a highly specific inhibitor capable of blocking KYN’s interactions with Ahr in cells such as bone marrow derived murine dendritic cells

(BMDCs); however, until now, the effects of KYN’s interaction with Ahr has not been explored in skeletal muscle (Mezrich et al., 2010). Ahr knockout mice treated with KYN had significantly lower quadriceps weight compared to VEH treated controls, but there was no significant differences between VEH and KYN-treated mice in muscle fiber size,

H2O2, or 4HNE staining in Ahr knockout mice. This could again be due to a threshold effect as Ahr knockout mice carry a number of systemic health problems, including vascular stiffness, high inflammatory cytokine load, and higher mortality rates (Bravo-

Ferrer et al., 2019; Eckers et al., 2016). Overall, these results suggest that KYN-induced skeletal muscle loss with age may occur through more than one pathway. The role of Ahr should further be explored using skeletal-muscle specific knockout mice and determining the effects of downstream KYN metabolites on Ahr activation in skeletal muscle.

Maintenance of skeletal muscle throughout life is dependent upon a balance of protein synthesis and protein catabolism (Atherton et al., 2016; Cuthbertson et al., 2005;

Moore et al., 2015; Wall et al., 2015). We examined proteins from young mice treated with KYN, and old mice treated with 1-MT, and found that lipid peroxidation products were differentially expressed with manipulation of the KYN pathway. Branched chain amino acid aminotransferase, mitochondrial very long chain acyl-coa dehydrogenase, mitochondrial long chain acyl-coa dehydrogenase, isocitrate dehydrogenase, and isocitrate dehydrogenase subunit gamma 1. The largest fold change difference was mitochondrial lipid peroxidation enzyme, Mvlcad, which we identified as a potential downstream target mechanism for KYN’s contribution to sarcopenia. Mvlcad has been previously shown to produce H2O2 (Kakimoto et al., 2015). Furthermore, Montes et. al demonstrated that lipid peroxidation markers can serve as indicators of sarcopenia, and

Bellanti et. al found that that lipid peroxidation products form aldehyde-protein hybrids that are increased in sarcopenic adults (Bellanti et al., 2018; Coto Montes et al., 2017).

Exercise has been previously reported to increase skeletal muscle ROS as well as

Mvlcad, suggesting that in the setting of acute inflammation transient, elevated ROS and

Mvlcad levels may have beneficial effects on skeletal muscle; however, it is likely that chronically elevated ROS and Mvlcad resulting from prolonged KYN exposure may ultimately have detrimental effects on muscle, which would explain the previous associations among aging, inflammation, circulating KYN, Mvlcad, ROS and sarcopenia noted above (Horowitz, Leone, Feng, Kelly, & Klein, 2000). The proposed model described here is outlined in figure 23.

To test the effect of KYN on inflammatory cytokines, a cytokine panel was run on tibialis anterior lysate, and serum from young male and female treated for 30 days with KYN or VEH. Interestingly female mice showed more inflammatory changes with KYN treatment than males, with inflammatory Il-1α increased in female muscle but decreased in female serum whereas male Il-1α matched female VEH. We speculate this result may be due to localization of Il-1α to damaged skeletal muscle. Females also showed a decrease in anti-inflammatory Il-10, and an increase in inflammatory Il-17α in serum (Abusleme & Moutsopoulos, 2017; Saraiva & O'Garra, 2010). Interestingly, kynurenic acid has been shown to decrease Il-17α in dendritic cells, and kynurenine has been shown to increase Il-10 in corneal epithelial cells (Matysik-Wozniak et al., 2017;

Salimi Elizei, Poormasjedi-Meibod, Wang, Kheirandish, & Ghahary, 2017). Furthermore,

Il-10 levels have been shown to increase in serum in sarcopenic adults (Rong, Bian, Hu,

Ma, & Zhou, 2018). The discrepancy between these studies and our results may be a difference between mice and human responses, or mechanisms outside of the KYN pathway. Furthermore, the downstream effects of these inflammatory cytokines likely further contribute to muscle atrophy (Sharma & Dabur, 2018). Both males and females had a highly significant decrease in Il-27p70, an inflammatory modulator (Yoshida &

Hunter, 2015). The discrepancy between male and female cytokine results may be due to hormonal differences and will need to be further explored.

Figure 22. Proposed model for KYN’s effects on lipid peroxidation, ROS generation, and age-related muscle atrophy. Aging causes an increase in systemic oxidative stress, and skeletal muscle antioxidant pathways are unable to meet the increased demand. Inflammatory cytokines like IFNγ activate IDO to convert tryptophan to KYN. KYN then induces the upregulation of mitochondrial mVLCAD which degrades lipid and produces H2O2. The overexpression of ROS leads to further inflammation and muscle atrophy. 1-MT is an antagonist of IDO that can inhibit tryptophan’s conversion into KYN, thereby attenuating KYN-induced tissue dysfunction with aging.

VI SUMMARY

In this study I tested the central hypothesis that an increase in KYN with age causes an imbalance in lipid peroxidation pathways, leading to oxidative stress and consequent muscle atrophy. This hypothesis was tested by using two specific aims:

Specific aim 1 tested the hypothesis that age-associated increases in KYN impair muscle function and decrease muscle mass by activating ubiquitin ligases and inhibiting protein synthesis. This hypothesis was mostly supported. Specifically, KYN impaired muscle function and decreased muscle mass in young mice. Inhibition of IDO by 1-MT attenuated this effect in old mice. No evidence that KYN activated ubiquitin ligases was found, however structural muscle proteins Myh1, Myh2, Tnnt3 and Ttn were reduced.

Specific aim 2 tested the hypothesis that KYN increases oxidative stress and lipid peroxidation in skeletal muscle cells. This hypothesis was supported. KYN increased

H2O2 and mitochondrial lipid peroxidation enzymes in mouse and human myoblasts.

Quadriceps femoris in mice treated with KYN showed increases in Mvlcad. These results were further attenuated in old mice by 1-MT treatment. Lipid peroxidation enzymes

Mlcad and Mvlcad were identified as upregulated with KYN treatment and contributory to H2O2 levels. In addition, inflammatory cytokines Il-1α, Il-17α, Il27p70 and Il-10 were dysregulated with KYN treatment in young mice.

This study demonstrated a critical role for KYN in increasing oxidative stress, lipid peroxidation, and altering the inflammatory microenvironment, contributing to the age-related loss of muscle mass and strength. My work establishes a promising

therapeutic target for sarcopenia that may provide further evidence to support inhibition of IDO for age-related conditions such as osteoporosis, and Alzheimer’s disease.

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APPENDIX A: Abbreviations

AA: Anthranillic acid Ahr: Aryl hydrocarbon receptor CA: Cinnabarinic acid CYP1A1: cytochrome P450 1A1 enzyme GPx: Glutatione peroxidase IDO: Indolemine 2, 3, dioxygenase KYN: Kynurenine Mafbx: Muscle atrophy F-box MHC: myosin heavy chain Murf1: Muscle RING finger 1 NA: Nicotinic acid Nico: Nicotinamide Nrf2: antioxidant nuclear factor erythroid 2–related factor 2 PA: Picolinic acid P53: Tumor protein 53 QA: Quinolinic acid ROS: Reactive oxidative species SLC7A5: solute carrier family 7 member 5 SOD: Super oxide dismutase mVLCAD: mitochondrial very long chain acyl-coa dehydrogenase 1-MT: 1-methyl tryptophan