TFAM enhances fat oxidation and attenuates high fat diet ......2019/05/03 · 1 TFAM enhances fat...

1

TFAM enhances fat oxidation and attenuates high fat diet induced insulin resistance in skeletal muscle

Jin-Ho Koh1, Matthew L. Johnson1, Surendra Dasari2, Nathan K. LeBrasseur3, Ivan Vuckovic4, Gregory C. Henderson1, Shawna A. Cooper1, Shankarappa Manjunatha1, Gregory N. Ruegsegger1, Gerald I. Shulman5, Ian R. Lanza1 and K. Sreekumaran Nair1,*

1. Division of Endocrinology and Metabolism, Mayo Clinic, Rochester, MN2. Department of Health Sciences Research, Mayo Clinic, Rochester, MN3. Department of Physical Medicine & Rehabilitation, Mayo Clinic, Rochester, MN4. Mayo Clinic Regional Comprehensive Metabolomics Core, Mayo Clinic, Rochester, MN5. Department of Medicine and Cellular and Molecular Physiology, Yale University, New

Haven, CT

*Corresponding authorK. Sreekumaran Nair, M.D., Ph.D.Professor of MedicineDivision of Endocrinology and MetabolismMayo Clinic College of Medicine200 First St SWRochester, MN 55905Tel: 507-255-2415Fax: 507-255-4828Email: [email protected]

Running Title: Muscle TFAM & insulin resistance

of 54 Diabetes

Diabetes Publish Ahead of Print, published online May 14, 2019

mailto:[email protected]

2

ABSTRACT

Diet induced insulin resistance (IR) adversely affects human health and life span. We show that

muscle specific overexpression of human mitochondrial transcription factor A (TFAM)

attenuates high fat diet (HFD)-induced fat gain and IR in mice in conjunction with increased

energy expenditure and reduced oxidative stress. These TFAM effects on muscle are shown to be

exerted by molecular changes that are beyond its direct effect on mitochondrial DNA replication

and transcription. TFAM augmented muscle tricarboxylic acid cycle and citrate synthase

facilitating energy expenditure. TFAM enhanced muscle glucose uptake despite increased fatty

acid (FA) oxidation in concert with higher β-oxidation capacity to reduce accumulation IR-

related carnitines and ceramides. TFAM also increased pAMPK expression explaining enhanced

PGC-1α, and PPARβ and reversing HFD-induced GLUT4 and pAKT reductions. TFAM-induced

mild uncoupling is shown to protect mitochondrial membrane potential against FA-induced

uncontrolled depolarization. These coordinated changes conferred protection to TFAM mice

against HFD-induced obesity and IR while reducing oxidative stress with potential translational

opportunities.

KEYWORDS: TFAM, mitochondria, insulin action, high fat diet, fatty acid oxidation,

uncoupling, PGC-1α, PPARβ, muscle and oxidative stress

of 54Diabetes

3

INTRODUCTION

An estimated 9% of the United States population has diabetes and an additional 86 million

representing 1 in 3 adults have prediabetes - a risk factor for type 2 diabetes mellitus (T2DM)(1).

Skeletal muscle insulin resistance (IR) is a cardinal feature of T2DM and it is often present years

prior to T2DM onset (2). Ameliorating IR could prevent or reverse T2DM. Altered mitochondrial

function has been shown to cause IR. For example, excessive mitochondrial H2O2 (mtH2O2)

emissions following high fat diet (HFD) results in IR via attenuation of insulin signaling and

glucose uptake (3). In IR women, abnormal mtH2O2 emission when normalized by aerobic

exercise training improved insulin sensitivity (4). Moreover, age-related IR occurs with reduced

mitochondrial DNA (mtDNA) abundance (5) and maintaining high abundance of mtDNA and

reduced oxidative stress ameliorate age-related IR (6). Overall it seems that efficient

mitochondrial function can improve insulin sensitivity. Lipid accumulation in skeletal muscle (7)

or metabolic overload in mitochondria (8) have also been shown to occur with IR, and it has been

shown that incomplete β-oxidation contribute to skeletal muscle IR (9). These studies support a

notion that enhances mitochondrial oxidative capacity and efficiency, especially the upregulation

of mtDNA abundance and reduction of oxidative stress ameliorate IR.

Mitochondrial transcription factor A (TFAM) is a key regulator of mitochondrial DNA

replication, repair and gene transcription (10). An ex vivo study demonstrated that overexpressing

TFAM in cardiomyocytes decreased mtH2O2 (11). In contrast, adipose tissue-specific deletion of

TFAM decreased ROS, increased mitochondrial fuel oxidation, and protected mice against

obesity and IR (12). These paradoxical studies suggest that TFAM’s role in regulating

mitochondrial fuel metabolism and ROS emission is tissue specific and TFAM’s effect on

skeletal muscle is currently unknown.

of 54 Diabetes

4

Hence, we sought to determine if muscle-specific overexpression of human TFAM in mice

(transgenic, Tg) can increase mtDNA abundance, β-oxidation and mitochondrial respiration,

decrease mtH2O2 generation, and prevent HFD-induced IR. We further investigated various

molecular and bioenergetics pathways by which TFAM influence insulin sensitivity and

oxidative stress in skeletal muscle. These mechanistic insights provide opportunities for potential

novel therapeutic approaches to counteract IR, and prevent and treat many metabolic disorders,

including T2DM.

RESEARCH DESIGN AND METHODS

Study procedures are outlined here, but more details are provided in the supplement.

Human TFAM transgenic animals. Human TFAM (TFAM) cDNA and muscle specific

creatinine kinase (MCK) promoter construct was inserted into a TA cloning vector

(pcDNA3.1/NT-GFP-TOPO). Purified vector was directly injected into pronucleus of single-

celled FVB mice embryos and transgenic (Tg) embryos were implanted in a pseudopregnant

mother. Sexually mature F1 Tg progeny were derived and utilized for all analyses. Male animals

of both Tg and wild were assigned to either chow diet (CHO) or high fat diet (HFD) for twelve

weeks. We chose only male animals to avoid confounding effects of fluctuating female hormones

on our outcomes.

Body composition. Body composition was measured, after an overnight fast, using EchoMRI

(Echo Medical Systems) and dual-energy X-ray absorptiometry (GE Medical-Lunar).

Metabolic and physical activity measurements. Measurements were performed in both fed and

fasted states. Oxygen consumption (VO2) and carbon dioxide production (VCO2) were measured

using an 16-chamber open-circuit cage calorimetry system (Columbus Instruments). Respiratory

of 54Diabetes

5

exchange ratio (RER) and energy expenditure was computed using standard equations.

Ambulatory levels were measured using infrared photocell beam interruption method.

Insulin sensitivity. Mice were fasted for 5hr and peripheral insulin sensitivity was measured

using a euglycemic clamp (120-130 mg/dL) maintained for 120 mins. Mice were primed with

16mU/kg bolus of Humulin R followed by 2.5mU/kg/min continuous infusion. Blood was

collected for every 10mins for glucose and insulin measurements (as described above).

Mitochondrial function. Mitochondria were isolated from quadriceps muscle and analyzed by

high-resolution respirometry (13). Mitochondria were added to a 2 mL chamber (Oxygraph-2K)

followed by sequential additions of glutamate, malate, ADP, succinate, and inhibitors.

Mitochondrial membrane integrity was verified with cytochrome C.

TFAM myotube cultures. Differentiated C2C12 myoblasts were infected with adenovirus bearing

TFAM (AD-TFAM, Applied Biological Materials Inc.) in either presence or absence of 0.2mM

palmitate. A similar protocol was followed to generate empty vector (EV) myotubes. All

myotubes were differentiated for 5 days prior to collecting them for various analyses.

Hydrogen peroxide measurements in muscle and myotubes. Quadriceps muscle mitochondria

were isolated, treated with glutamate and malate, and H2O2 emission was measured using

Amplex Red oxidation as previously published (14). Myotubes were suspended in respiration

medium and membranes were permeabilized using digitonin. Glutamate and malate were added

and H2O2 was measured as described above.

Muscle DNA oxidative damage and redox measurements. Mice gastrocnemius muscle was

homogenized and the adduct biomarker 8-oxo-2’deoxyguanosine (8-oxo-dG) was measured as

previously described (6). Muscle redox state was assessed by measuring oxidized (GSSG) and

reduced (GSH) glutathione using a commercial kit (Cayman Chemicals).

of 54 Diabetes

6

mRNA measurements. Semi-quantitative RT-PCR was utilized to measure PGC-1α, CS,

NDFAU9, SHDH, UQRCR1, UQRCR2, CYCS, MT-CO1, COX1, COX4, APT5A, and UCP3

genes.

Western blots. Protein abundance was measured with antibodies (Abs) against PPARβ, ATP

synthase subunit α (ATPsyn), Succinate-ubiquinone oxidoreductase (SUO), ND ubiquinone

oxidoreductase (ND), ubiquinone-cytochrome c oxidoreductase Core subunit 1 (Core1) and 2

(Core2), cytochrome c oxidase subunit (COX) I and IV, caspase 3, PGC-1α, UCP3, c-Myc-Tag,

pS473-AKT, pT308-AKT, total AKT, total AMPK, pAMPK, GLUT4, PPARα, SOD2, Catalase,

NRF-1, MEF-2A, CaMKKβ, citrate synthase (CS), and cytochrome c (Cyt c).

Immunoprecipitation studies. Anti-Myc or anti-ubiquitin antibody was mixed with magnetic

beads for 1.5hr at RT. 250ug protein from each group were added in antibody and beads mixture

and then rotated overnight. Ab-antigen complexes were captured on magnetic beads and probed

using western blot analysis for Cyt C, ND, Core1 or COX1 Ab.

Muscle lipid measurements. Acylcarnitines (C0, C2, C3, C4, C5, C8, C12, C14, C16, C18, and

C18:1), ceramides (cer14, Cer16, Cer18, Cer24, Cer24:1, sphinganine [SPA], sphingosine [SPH]

and sphingosine-1-phosphate [S1P]), saturated (16:0/16:0) diacylglycerols and unsaturated

diacylglycerols (18:0/18:2, 18:0/18:1, 18:1/18:1, 18:2/18:2, 16:0/18:1) were extracted from

gastrocnemius muscle and analyzed via LC-MS as previously described (15-17).

Muscle energy metabolite NMR studies. Tissue chunks were weighed, homogenized, and

metabolites were extracted. 1H spectra were recorded using 600MHz NMR spectroscopy.

Metabolites were identified and quantified using Chenomx software and expressed as µmol per g

of tissue analyzed.

of 54Diabetes

7

Membrane potential (Δψm) studies. TFAM or empty vector was inserted into differentiated

myotubes. After 5 days, cells were preloaded with TMRM (tetramethylrhodamine methyl ester)

and Hoechst 33342. Labeled cells were resuspended in imaging buffer supplemented with either

glucose or palmitate for 2 hours. Cells were imaged continuously for 480secs in 10sec intervals.

During the imaging study, 2M FCCP was added at 180secs and 8M FCCP was added at

300secs. TMRM intensity was calculated as Δt=(tX-t0)/t0 X 100, where tx=fluorescence intensity

at any time point, t0 = baseline fluorescence.

DNP studies. After 5 days of cell differentiation, C2C12 cells were incubated for 24 hour with

3M (final concentration) of 2,4-Dinitrophenol (DNP) (kindly provided by Dr. Geisler, Mitochon

Pharmaceuticals, Inc; Blue Bell, PA 19422) or DMSO as control in differentiation medium.

Statistical tests. Data were analyzed using either Student’s t-test, One-way ANOVA, two-way

ANOVA, Mann-Whitney rank sum test wherever appropriate. Statistical significance was

denoted as * for p<0.05, ** for p<0.01, *** for p<0.001and **** for p<0.0001.

RESULTS

Muscle specific TFAM overexpression increases mtDNA copy number and metabolic rate

while attenuating HFD-induced fat accumulation and insulin resistance.

We generated Tg FVB mice with skeletal muscle specific overexpression of human TFAM (Fig.

1A). Muscle TFAM mRNA, protein (Fig. 1B), mtDNA copy number (Fig. 1C), and

mitochondrial-encoded mRNAs (Supp. Fig. 1) were higher in Tg mice compared to wild type

(Wt) littermates.

Mice on twelve weeks high fat diet (HFD; 60% calorie from fat) consumed more calories than

chow mice but no differences in caloric intake were observed between Wt and Tg mice (Supp.

Fig. 2). However, Tg mice had lower body weight and lean mass than Wt mice irrespective of the

of 54 Diabetes

8

diet (Figs. 1D and 1E). Moreover, Tg mice on HFD gained less body weight and fat mass than

Wt mice on HFD (Fig. 1E-F). Based on indirect calorimetry, we observed that Tg mice on HFD

had higher metabolic rate during both day and night when compared to Wt on HFD mice,

irrespective of whether they were in fasted or fed state (Fig. 1G). The VO2 and VCO2

measurements mirrored the metabolic rate differences observed between Tg and Wt mice (Fig.

1H), although respiratory exchange ratio (RER) was unaltered by TFAM (Supp. Fig. 3).

Curiously, HFD increased nightly activity of Tg mice than Wt mice (Fig. 1I) and this

phenomenon was absent with chow diet (Supp. Fig. 4) and it is possible that these increased

activity contributed to nightly increase in energy expenditure. These increased activity levels can

be potentially attributed to higher mtDNA abundance and functional capacity (18).

Hyperinsulinemic euglycemic clamp showed that HFD significantly reduced the insulin

sensitivity in Wt mice when compared to chow diet (Figs. 1J-1K). However, Tg mice on HFD

demonstrated lesser decline in insulin sensitivity than Wt HFD mice based on glucose infusion

rate during hyperinsulinemic euglycemic clamp and muscle glucose uptake (Figs 1J-1K). We did

not observe any difference in endogenous glucose production during the euglycemic clamp (Supp.

Fig. 5), indicating that the effects of TFAM in insulin sensitivity were restricted to peripheral

tissues. An intraperitoneal insulin tolerance test indicated that HFD in Wt decreased insulin

sensitivity whereas Tg mice on chow or HFD had same levels of insulin sensitivity as Wt chow

mice (Supp. Fig. 6). Muscle glucose uptake also decreased in both Wt and Tg mice with HFD,

however, Tg mice on HFD had higher glucose uptake than Wt HFD mice (Fig. 1L) indicating

that TFAM attenuated HFD induced decline in muscle glucose uptake. Blood glucose levels of

Tg HFD mice were lower than Wt HFD mice (Fig. 1M).

of 54Diabetes

9

Collectively, these results data indicate that skeletal muscle overexpression of TFAM increased

mtDNA abundance and metabolic activity while attenuating HFD-induced weight gain and loss

of insulin sensitivity.

TFAM increases mitochondrial lipid oxidative capacity.

We studied the bioenergetic characteristics of muscle mitochondria to understand the potential

link to increased metabolic rate and attenuated fat accumulation. Chow Tg mice had significantly

higher state 3 respiration when using palmitoyl-carnitine and malate (PCM) as substrate, and

lower state 3 respiration when using glutamate and malate (GM) as substrates, when compared to

Wt chow mice normalizing for protein content (Fig. 2A) as well as when normalized to the tissue

weight (Supp. Fig. 7). Mitochondrial respiratory efficiency, measured as respiratory control ratio

(RCR), was significantly higher in Tg compared to WT with PCM and significantly lower with

GM substrates (Fig. 2B). Similar phenomena were observed in Tg mice on HFD (Figs. 2C & 2D).

These data revealed that TFAM decreases coupling efficiency when respiration is supported

through carbohydrate-based substrates but increases coupling efficiency when respiration is

supported by fatty acid (FA) substrates. Overall, these results support that Tg mice exhibit

marked shifts in skeletal muscle mitochondrial bioenergetics whereby fat oxidative capacity is

increased, complex I supported respiratory capacity is decreased.

TFAM enhances skeletal muscle energy metabolism.

We observed higher amounts of ADP and AMP and lower amounts of ATP and creatine

phosphate (CrP) in skeletal muscle of Tg mice compared to Wt mice regardless of diet (Figs. 2E-

2K) and higher AMP:ATP ratio is known to increase activation of AMPK (19). In contrast to the

lower ATP in Tg mice, we observed higher levels of citrate, citrate synthase (CS) protein, CS

activity and other tricarboxylic acid (TCA) cycle metabolites when compared to Wt mice in both

of 54

For Peer Review Only

Diabetes

10

chow and HFD groups (Figs. 2L-2S). Energy demands of high metabolic rate, like during

exercise, increases TCA cycle activity, and decreases CrP (20), as observed in Tg mice who has

high energy demand of muscle. TFAM also increased hexokinase enzyme activity (Fig. 2T) and

lactate, indicating TFAM increased glycolysis (Fig. 2U). Additionally, glycogen synthase

phosphorylation was increased by TFAM and HFD, indicating impairment in glycogen storage

(Supp. Fig. 8). Increased malate and fumarate as well as aspartate and glutamate with reduced

glutamine suggests increased aspartate-malate shuttle and increased conversion of glutamine to

glutamate to utilize in the TCA cycle. These data also suggest that TFAM increased the aspartate-

malate shuttle to increase NAD+ import into the mitochondrial matrix. Carnosine, an inhibitor of

electron transport chain in animals (21), was also lower in Tg mice compared to Wt mice in both

chow and HFD groups (Fig. 2V). The contrasting findings of lower ATP levels and higher levels

of TCA activity in the skeletal muscle of Tg mice when combined with the fact that Tg mice had

higher metabolic rate and activity levels suggest that Tg mice consume ATP at a higher level.

TFAM lowers skeletal muscle ROS emissions and oxidative stress.

HFD is known to increase mtH2O2 production (3). As expected, Wt mice on HFD had the highest

mtH2O2 emissions followed by Wt mice on chow (Fig. 3A). Of interest, Tg mice had the lowest

mtH2O2 emissions irrespective of the diet (Fig. 3A). This is further supported by reduced ROS

mediated DNA damage (Fig. 3B) and lower GSSH/GSH ratio indicates lower oxidative stress

(Fig. 3C) regardless of diet. Concomitantly, Tg mice had higher levels of endogenous anti-

oxidant defense enzymes (superoxide dismutase 2, catalase and cytochrome C [Cyt c]) when

compared to Wt mice regardless of diet (Fig. 3D). These data indicate that muscle TFAM can

increase FA oxidation while lowering ROS emissions and enhance anti-oxidant defense.

of 54


Diabetes

11

Next, we sought to determine the potential mechanism(s) by which TFAM influences ROS

production. Irrespective of the diet, Tg mice had higher levels of PPAR (Fig. 3E), which can

regulate catalase (22) and SOD2 (23). Tg HFD mice had the highest levels of Cyt C when

compared to rest of the groups (Fig. 3D) and Cyt C under non-apoptotic conditions can act as a

ROS scavenger (24). We found TFAM decreased mtH2O2, increased SOD2 and catalase (Figs.

3F-3G) and blocked the release of cleaved caspase (Fig. 3H) with palmitate treatment in myotube.

Palmitate treatment increased cytosolic Cyt C levels in empty vector (EV) cells when compared

to sham treatment whereas mitochondrial Cyt C levels did not change (Fig. 3I). In contrast,

TFAM overexpression increased the mitochondrial Cyt C levels regardless of palmitate or sham

treatment while reducing the cytosolic Cyt C levels to basal level when treated with palmitate

(Fig. 3I).

Since Tg mice had higher levels of activated AMPK (pAMPK) regardless of diet (Fig. 3E), we

hypothesized that TFAM may be using pAMPK to hold Cyt C in the mitochondria to act as a

ROS scavenger, and found that activated AMPK binds to Cyt C (Fig. 3J). The net effect of

reduced ROS emission and increased anti-oxidant defense by TFAM also attenuated the ROS

induced loss of GLUT4 observed with PM treatment (Fig. 3G). All of these data support a notion

that muscle TFAM controls ROS via SOD2, catalase and Cyt C via PPAR and pAMPK (Fig.

3K).

TFAM increases mitochondrial β-oxidative capacity and prevents accumulation of insulin

resistance-associated bioactive lipids in skeletal muscle.

We measured β-oxidation and found HFD significantly increased LCAD, a primary -oxidation

enzyme (Fig. 4I) and lower medium-chain (C5 to C12) and long-chain (C16 to 18) acylcarnitines,

however, HFD did not increase short-chain (C0 to C4) in Wt. These data indicate that HFD

of 54


Diabetes

12

induces an incomplete -oxidation. We also found that, irrespective of diet, Tg mice had lower

very long-, long and medium-chain acyl-carnitines (Figs. 4D-H) and higher levels of LCAD (Fig.

4I). However, Tg mice had higher levels of short-chain acylcarnitines when compared to Wt mice

in both chow and HFD groups (Figs. 4A-4B). These data suggest that TFAM overexpression in

skeletal muscle activates a complete β-oxidation. Diacylglycerol (DAGs) have been reported to

reduce insulin sensitivity in skeletal muscle (25). We found that TFAM influences abundance of

DAGs (Supp. Fig. 9). As reported by Ameti et al (26) that DAGs are higher in exercise-trained

muscle corresponding to higher insulin sensitivity, we also found that muscle DAGs in TFAM Tg

muscle are higher concurrent to higher insulin sensitivity. These data indicate that the cellular

DAG content cannot explain skeletal insulin sensitivity induced by TFAM.

HFD has been shown to increase abundances of intramyocellular diacylglycerol as well as

ceramides (27), and certain classes of ceramides are associated with skeletal muscle IR (28).

Further, a decrease in palmitoylcarnitine (Fig. 4F), a precursor of palmitoyl-CoA-based ceramide

synthesis, in Tg mice suggested that TFAM may affect muscle ceramide levels. We found Tg

mice had higher levels of sphingosine (SPH), sphinganine (SPA), -1-phosphate (S1P) regardless

of the diet (Fig. 4J). S1P has been shown to increase muscle’s ability to use FA as a fuel for

energy needs (29). Cer14, Cer16, Cer24:1, and Cer24 were higher in Tg mice on chow when

compared to Wt mice on chow diet whereas HFD did not significantly change their levels

between Tg and Wt mice (Figs. 4K-4L,4N-4O). In contrast, Cer18 is associated with IR (28), was

higher in Wt mice on HFD when compared to chow diet whereas Tg mice on HFD had levels

similar to that of Wt or Tg mice on chow diet (Fig. 4M). These data suggest that, even though

overexpression of TFAM increases intramyocellular de novo ceramide synthesis, TFAM

of 54


Diabetes

13

differentially regulates Cer18 levels, a potent inhibitor of insulin signaling, and TFAM control β-

oxidation to block accumulation of incompletely oxidized lipids that is IR factor.

Collectively, these data indicate that high fat diet induces an incomplete β-oxidation. However,

skeletal muscle TFAM enhances both β-oxidation pathway and de novo ceramide synthesis

pathway to increase FA oxidation while attenuating the accumulation of IR associated bioactive

lipids (Fig. 4O).

TFAM overexpression remodels electron transport chain in a posttranslational manner and

attenuates fatty acid-induced membrane depolarization

High levels of free fatty acid (FFA) were shown to alter ATP synthesis in skeletal muscle (30).

However, the precise effect of HFD on electron transport chain (ETC) enzymes has never been

studied. Further, Tg mice on HFD had increased FA oxidative capacity, decreased ROS and

increased energy metabolism (Figs. 2-4), which would require adaptation of the ETC by TFAM

to accommodate the high fuel flux. HFD in Wt mice increased protein expression of ND and

CORE2 (Fig. 5A) and decreased ATP synthase α (ATPsyn) when compared to chow diet (Fig.

5A). However, Tg mice had significantly increased protein expression of ATPsyn and SUO

proteins when compared to Wt mice in both chow and HFD groups (Fig. 5A). These data suggest

that TFAM overexpression can attenuate HFD-induced loss of complex V. Tg mice had lower

protein expression of ND, CORE1, UCP3 and COX1 proteins in both diet groups compared to

Wt (Fig. 5A). Ubiquitination of ND, CORE1, and COX1 proteins also increased in Tg mice (Fig.

5B). These data suggest the TFAM overexpression downregulates complex I, complex III and

complex IV, which pumps proton into mitochondrial intermembrane space, via ubiquitination.

mRNA measurements showed that protein changes induced by TFAM in ETC pathway are

of 54


Diabetes

14

mediated in a posttranslational manner via protein ubiquitination rather than direct transcriptional

changes (Fig. 5C).

The asymmetric reformation of key ETC complexes by TFAM is likely to alter the mitochondrial

membrane potential (31). Further, TFAM mice showed different coupling efficiency with fatty

acid or glucose as fuel substrate (Figs. 2B & 2D). At resting state, TFAM myotubes have lower

tetramethylrhodamine methyl ester (TMRM) intensity (i.e. surrogate for Δψm) in the

mitochondria than EV when using glucose as substrate and higher Δψm when using palmitate as

substrate (Fig. 5D). When a small amount (2µM) of an ionophore (FCCP), which is an uncoupler

disrupting ATP synthesis, was added, mitochondria significantly depolarized in EV cells treated

with palmitate when compared to TFAM cells (Figs. 5D & 5E). In contrast, when glucose was

being used as fuel substrate, TFAM cells lost significant Δψm compared to EV cells with 8µM

FCCP, however, Δψm was protected by TFAM when 2µM FCCP was added (Figs. 5D & 5E).

Time resolved changes in the Δψm are shown in Figs 5F and 5G. These results indicate that

TFAM induces Δψm mild uncoupling, perhaps via decreased ND, Core1 and COX1 (Fig. 5A).

Simultaneously, TFAM is also protecting Δψm, perhaps via ATP synthase reversal. This is

supported by previous studies showing that Δψm protection (i.e. mild uncoupling) could be

mediated via ATP synthase reversal (Fig. 5H) when mitochondrial ETC is altered (32) or

glycolysis is increased (33). In addition, a previous study showed that an increased Cyt C

(observed with TFAM overexpression; Fig. 4D) is associated with mild Δψm uncoupling (34). All

of these data suggest that Δψm coupling efficiency in TFAM overexpressed muscle is higher with

fatty acid than glucose, showing the preference of fatty acid as the energy source in TFAM

muscle.

of 54


Diabetes

15

Muscle TFAM overexpression attenuates HFD-induced insulin resistance by increasing

glucose uptake and disposal via pAMPK or PPARβ or PGC-1α

A previous study suggested that HFD-induced insulin resistance cannot be explained on the basis

of changes in electron chain complexes (35). Next, we sought to identify the mechanism on how

TFAM preserves insulin sensitivity under HFD conditions. We measured skeletal muscle pAKT

in response to insulin in Wt and Tg mice on either chow or HFD. HFD attenuated the pAKT in

Wt mice in response to insulin when compared to chow diet (Fig. 6A). In contrast, Tg mice

preserved the insulin mediated pAKT under HFD state (Fig. 6A), that is associated with

enhanced energy metabolism (Fig. 2), antioxidant buffering system (Fig. 3) and higher β-

oxidation (Fig. 4) by TFAM. Further, TFAM overexpression results in an increase GLUT4 (key

regulator of glucose transport) expression, which is directly regulated by MEF2A (36). Further,

NRF-1, upstream of MEF2A, is increased by PPARβ (21), these factors also were upregulated by

TFAM overexpression irrespective diet. We also found pAMPK, which translocate GLUT4 to

uptake glucose, is activated by TFAM. We also observed increased expression of PGC-1α protein

in Tg mice in both diet groups (Fig. 6B), and this pathway also may increase GLUT4 expression

and translocation (37). Thus our results indicate that TFAM alters pAMPK or PPARβ or PGC-1α

pathways (Fig. 6C) and offer additional mechanistic explanation of enhanced insulin sensitivity

and glucose uptake observed in Tg mice on HFD (Fig. 1J-1L).

DISCUSSION

The current study demonstrates that human-TFAM overexpression in skeletal muscle has unique

and profound impact on multiple molecular pathways regulating energy metabolism. These

effects are distinct from what has been reported in other tissues when TFAM is overexpressed (11;

12) or PGC-1α is overexpressed in muscle (38). Most of the molecular changes noted are beyond

of 54


Diabetes

16

the direct effects of TFAM as transcription factor of mitochondrial biogenesis. Importantly, Tg

mice increased fatty acid oxidation but warded off HFD-induced decline in glucose disposal, thus

counteracting the well-known glucose-fatty acid cycle (39). A notable effect of TFAM

overexpression is the enhancement energy expenditure in conjunction with higher mitochondrial

fatty acid (FA) oxidative capacity which contributed to reduced fat accumulation and metabolites

of incomplete fat oxidation. TFAM also despite enhanced fatty acid oxidation reduced oxidative

stress in mice on HFD. We used hTFAM in the current study that has >70% sequence homology

to that of mouse and DNA binding domains were conserved. The objective was to understand the

biological effects of hTFAM but the effects related to the species differences cannot be fully

excluded.

The most striking effect of TFAM was on energy metabolism in muscle by enhancing

fatty acid oxidation and reducing accumulation of fat and incomplete metabolites of fatty acid

oxidation. These changes likely contributed to higher insulin sensitivity as noted previously (28).

We further addressed the potential molecular underpinnings on how TFAM preserved muscle

glucose uptake on HFD. As expected (3), 12-week HFD in Wt mice reduced glucose uptake in

muscle (Fig. 1L) in conjunction with reduced Akt phosphorylation. TFAM overexpression

increased AMPK activity in conjunction MEF2A expression that likely contributed to increased

GLUT4 expression (43) (Figs. 3E & 6B). Experiments in cell lines show a direct link between

AMPK activation and AKT phosphorylation (40). Our results support that reduction of AMPK

activation by HFD (Fig. 3E) led to a decrease in AKT activation (Fig. 6A), which seems to be

counteracted by increased AMPK and AKT activation by TFAM (Fig. 3E & 6A). This indicates a

novel role for TFAM in skeletal muscle glucose uptake via AMPK. Of interest, activity levels in

Tg mice on HFD increased especially in the night which has been reported to occur with

of 54


Diabetes

17

enhanced mitochondrial capacity (18) and this increased activity also could contribute to insulin

sensitivity and activation of AMPK. Some of the TFAM effects are similar to hyperthyroid state

although unlike hyperthyroid state oxidative stress is lower in TFAM mice. Another important

effect of TFAM was reduction in ROS emission and oxidative stress even with HFD and high

fatty acid oxidation (Fig. 3A) that also could explain enhanced insulin sensitivity in Tg mice).

Chronic overproduction of ROS increases oxidative stress, damages DNA, increases apoptosis as

well as triggering inflammation which is a key feature in T2DM and metabolic syndrome (41).

Previous studies showed that IR induced by HFD can be mitigated by elimination of ROS (42;

43). In the current study, TFAM appears to utilize two different mechanisms to prevent HFD-

induced overproduction of ROS: TFAM increased anti-oxidant enzymes SOD2 and catalase (Fig.

4D) and increased mitochondrial abundance of Cyt C (Fig. 3I), a potent ROS scavenger. TFAM

increased PPARβ (Fig. 3E), which can upregulate SOD2 and catalase (22). Hence, the increased

insulin sensitivity observed in TFAM HFD mice may also be related to anti-oxidant buffers via

PPARβ and eliminating ROS.

TFAM is a mitochondrial transcription factor. TFAM overexpression in muscle increased -

oxidation of FAs (Fig. 4), TCA cycle (Fig. 2) and some nuclear encoded protein expression (Figs

5 & 6). All of these effects cannot be directly regulated by TFAM by its well-known direct effect

on mtDNA as many proteins in TCA cycle and betaoxidation are encoded by nuclear DNA.

Membrane potential (Δψm) depolarization is known to elevate the cytoplasmic Ca2+ level (44),

and we found TFAM overexpression induces Δψm depolarization with glucose substrate (Fig. D-

G) and an increase CaMKKβ expression (Fig. 6B), indicating that TFAM signals to nuclear by

Ca2+/CaMKKβ pathway to induce nuclear encoded proteins such as PGC-1α and PPARβ (Figs

6B & 3E). PPARβ activation results in an increase lipid oxidation and glucose metabolism as

of 54


Diabetes

18

previously reported (45) and improves insulin sensitivity via PGC-1α (46). HFD promotes fatty

acid oxidation (Fig. 4) and increased PPARα expression in muscle (Fig. 6) and muscle-specific

PPARα overexpression without increasing activity (i.e. muscle contraction) has been reported to

increases β-oxidation and decreases glucose uptake and oxidation (47). By enhancing TCA cycle

based on TCA cycle substrates (Fig. 2) TFAM thus seems to have counteracted the imbalance

between β-oxidation and TCA cycle following HFD (Fig. 4) and prevented accumulation of acyl-

CoAs, their respective acyl-carnitines, and perhaps other as yet unidentified metabolites that

could contribute to mitochondrial failure (9) and insulin resistance. TFAM overexpression in

skeletal muscle results in an increases in PGC-1α (Fig. 6) which also controls to cope with fatty

acid load by coordinately regulating β-oxidation, TCA cycle and ETC activity (48). The

mitochondria in Tg mice muscle prefer fatty acid than glucose to produce ATP (Fig. 2 and 5), this

is likely related to PPARβ and PGC-1α enhancement by TFAM (Fig. 3E and 6B). PPARβ and

PGC-1α increased the use of fatty acid by pyruvate dehydrogenase kinase 4 (PDK4) (49; 50),

PPARβ also is reported to mediate glucose uptake and glycolysis (45), thus, TFAM

overexpression in muscle results in an increases β-oxidation as well as glycolysis as noted by

increased lactate (Fig. 2). These effects are associated with PPARβ enhanced mitochondrial

coupling efficiency and Δψm in Tg muscle seem to be specific with fatty acid (Fig. 5). Thus, HFD

induced defect in glucose uptake was attenuated by TFAM (Fig. 1K & 1L), possibly related to

TFAM preference for fatty acid than glucose for muscle fuel (Fig. 2A-2D, 5D-5G).

Δψm is an important factor for maintaining cellular energy homeostasis, and small reductions in

Δψm induced by mild uncoupling were shown to decrease H2O2 emissions (51; 52) and produce a

natural antioxidant effect (53). Hence, mild mitochondrial uncoupling can be potentially

therapeutic in disorders induced by oxidative stress. We found that DNP, an established

of 54


Diabetes

19

uncoupler, induced activation of AKT and AMPK and expression of GLUT4 (Fig. 6D) thus

contributing to insulin sensitivity but had no effect on PPARβ or any mitochondrial enzymes (Fig.

6E). Generally, mitochondrial uncoupling increases proton leak, resulting in an increase

AMP/ATP ratio and induce various energy sensing pathways (54). TFAM overexpression,

however, did not increase FA-induced proton leak, but rather decreased expression of UCP3 (Fig.

5A). TFAM also prevented FA-induced depolarization of mitochondria (Figs. 5D-5G). We

elucidated two potential mechanisms by which TFAM could be maintaining Δψm when

challenged with FA. Firstly, TFAM is inducing mild uncoupling by asymmetrically changing

mitochondrial enzymes (Fig. 5A). Decreasing complex I, II and IV would lower Δψm but

enhanced complex V (ATP synthase) likely enhanced ATP production. Secondly, Cyt C can

regulate Δψm and reduce H2O2 emission through AMPK activation (34). In the current study,

TFAM decreased H2O2 emission and increased Cyt C which was bound to pAMPK (Figs. 3A &

3J). Thus, our results indicate that TFAM controls mitochondrial Δψm uncoupling via Cyt

C/AMPK and reformation of ETC.

In conclusion, the current study demonstrated that muscle specific overexpression of human

TFAM has hitherto unknown beneficial metabolic effects. TFAM overexpression attenuated

HFD-induced loss of glucose uptake and insulin sensitivity despite increasing FA oxidation.

TFAM also prevented the oxidative stress that occurs on HFD by enhancing endogenous

antioxidant defense. TFAM enhanced TCA cycle and ATP synthase and by pathways involving

AMPK, PPARβ and PGC-1αTFAM also caused Δψm uncoupling as potential mechanisms by

which TFAM is countering many adverse effects of HFD. These identified pathways are rich in

therapeutic targets that can ameliorate insulin resistance and oxidative stress and potentially treat

obesity.

of 54


Diabetes

20

Author Contributions

K.S.N designed, supervised the study, analysis and data interpretation. G.C.H, S.C, I.R.L, G.I.S,

I.V, N.K.L, M.L.J, J.H, G.N.R. and S.D conducted various aspects of the study. M.L.H

performed mitochondrial phenotyping studies. G.C.H, S.A.C and NK conducted animal studies.

Yale Core Lab (G.I.S) performed hyperinsulinemic clamp and muscle glucose uptake

measurements. J.H., S.D, and I.V conducted metabolite measurements. J.H. conducted all cell

line studies and molecular phenotyping studies. S.D conducted statistical analyses. J.H., S.D. and

K.S.N. drafted the manuscript and all authors contributed to the final version of the manuscript.

Acknowledgements: K.S.N. is the guarantor of this work and, as such, had full access to the data

in the study and takes responsibility for the integrity of the data and the accuracy of the data

analysis. The study was supported by David Murdock Dole Professor ship (KSN), Grants from

National Institute of Health (RO1 DK41973-KSN, TRDK007352-GCH, DK007198-MLJ), Mayo

Clinic Metabolomics Core (supported by U24 DK100469). We also gratefully acknowledge

skillful technical support of Katherine Klaus and Dawn Morse.

Data and Resource Availability:

The datasets generated during and/or analyzed during the current study are available from the

corresponding author upon reasonable request. No applicable resources were generated or

analyzed during the current study.

Conflict of interest statement

The authors have declared that no conflict of interest exists.

of 54


Diabetes

21

REFERENCES

1. Dall TM, Yang W, Halder P, Pang B, Massoudi M, Wintfeld N, Semilla AP, Franz J, Hogan PF: The economic burden of elevated blood glucose levels in 2012: diagnosed and undiagnosed diabetes, gestational diabetes mellitus, and prediabetes. Diabetes Care 2014;37:3172-3179

2. DeFronzo RA, Tripathy D: Skeletal muscle insulin resistance is the primary defect in type 2 diabetes. Diabetes Care 2009;32 Suppl 2:S157-163

3. Anderson EJ, Lustig ME, Boyle KE, Woodlief TL, Kane DA, Lin CT, Price JW, 3rd, Kang L, Rabinovitch PS, Szeto HH, Houmard JA, Cortright RN, Wasserman DH, Neufer PD: Mitochondrial H2O2 emission and cellular redox state link excess fat intake to insulin resistance in both rodents and humans. J Clin Invest 2009;119:573-581

4. Konopka AR, Asante A, Lanza IR, Robinson MM, Johnson ML, Dalla Man C, Cobelli C, Amols MH, Irving BA, Nair KS: Defects in mitochondrial efficiency and H2O2 emissions in obese women are restored to a lean phenotype with aerobic exercise training. Diabetes 2015;64:2104-2115

5. Short KR, Bigelow ML, Kahl J, Singh R, Coenen-Schimke J, Raghavakaimal S, Nair KS: Decline in skeletal muscle mitochondrial function with aging in humans. Proc Natl Acad Sci U S A 2005;102:5618-5623

6. Lanza IR, Zabielski P, Klaus KA, Morse DM, Heppelmann CJ, Bergen HR, 3rd, Dasari S, Walrand S, Short KR, Johnson ML, Robinson MM, Schimke JM, Jakaitis DR, Asmann YW, Sun Z, Nair KS: Chronic caloric restriction preserves mitochondrial function in senescence without increasing mitochondrial biogenesis. Cell Metab 2012;16:777-788

7. Yu C, Chen Y, Cline GW, Zhang D, Zong H, Wang Y, Bergeron R, Kim JK, Cushman SW, Cooney GJ, Atcheson B, White MF, Kraegen EW, Shulman GI: Mechanism by which fatty acids inhibit insulin activation of insulin receptor substrate-1 (IRS-1)-associated phosphatidylinositol 3-kinase activity in muscle. The Journal of biological chemistry 2002;277:50230-50236

8. An J, Muoio DM, Shiota M, Fujimoto Y, Cline GW, Shulman GI, Koves TR, Stevens R, Millington D, Newgard CB: Hepatic expression of malonyl-CoA decarboxylase reverses muscle, liver and whole-animal insulin resistance. Nature medicine 2004;10:268-274

9. Koves TR, Ussher JR, Noland RC, Slentz D, Mosedale M, Ilkayeva O, Bain J, Stevens R, Dyck JR, Newgard CB, Lopaschuk GD, Muoio DM: Mitochondrial overload and incomplete fatty acid oxidation contribute to skeletal muscle insulin resistance. Cell Metab 2008;7:45-56

10. Ekstrand MI, Falkenberg M, Rantanen A, Park CB, Gaspari M, Hultenby K, Rustin P, Gustafsson CM, Larsson NG: Mitochondrial transcription factor A regulates mtDNA copy number in mammals. Hum Mol Genet 2004;13:935-944

11. Ikeda M, Ide T, Fujino T, Arai S, Saku K, Kakino T, Tyynismaa H, Yamasaki T, Yamada K, Kang D, Suomalainen A, Sunagawa K: Overexpression of TFAM or twinkle increases mtDNA

of 54


Diabetes

22

copy number and facilitates cardioprotection associated with limited mitochondrial oxidative stress. PLoS One 2015;10:e0119687

12. Vernochet C, Mourier A, Bezy O, Macotela Y, Boucher J, Rardin MJ, An D, Lee KY, Ilkayeva OR, Zingaretti CM, Emanuelli B, Smyth G, Cinti S, Newgard CB, Gibson BW, Larsson NG, Kahn CR: Adipose-specific deletion of TFAM increases mitochondrial oxidation and protects mice against obesity and insulin resistance. Cell Metab 2012;16:765-776

13. Lanza IR, Nair KS: Functional assessment of isolated mitochondria in vitro. Methods in enzymology 2009;457:349-372

14. Lanza IR, Blachnio-Zabielska A, Johnson ML, Schimke JM, Jakaitis DR, Lebrasseur NK, Jensen MD, Sreekumaran Nair K, Zabielski P: Influence of fish oil on skeletal muscle mitochondrial energetics and lipid metabolites during high-fat diet. Am J Physiol Endocrinol Metab 2013;304:E1391-1403

15. Chace DH, DiPerna JC, Mitchell BL, Sgroi B, Hofman LF, Naylor EW: Electrospray tandem mass spectrometry for analysis of acylcarnitines in dried postmortem blood specimens collected at autopsy from infants with unexplained cause of death. Clinical chemistry 2001;47:1166-1182

16. Blachnio-Zabielska AU, Persson XM, Koutsari C, Zabielski P, Jensen MD: A liquid chromatography/tandem mass spectrometry method for measuring the in vivo incorporation of plasma free fatty acids into intramyocellular ceramides in humans. Rapid communications in mass spectrometry : RCM 2012;26:1134-1140

17. Blachnio-Zabielska AU, Zabielski P, Jensen MD: Intramyocellular diacylglycerol concentrations and [U-(1)(3)C]palmitate isotopic enrichment measured by LC/MS/MS. J Lipid Res 2013;54:1705-1711

18. Chow LS, Greenlund LJ, Asmann YW, Short KR, McCrady SK, Levine JA, Nair KS: Impact of endurance training on murine spontaneous activity, muscle mitochondrial DNA abundance, gene transcripts, and function. Journal of applied physiology (Bethesda, Md : 1985) 2007;102:1078-1089

19. Hardie DG: AMP-activated protein kinase: a key system mediating metabolic responses to exercise. Medicine and science in sports and exercise 2004;36:28-34

20. Constable SH, Favier RJ, McLane JA, Fell RD, Chen M, Holloszy JO: Energy metabolism in contracting rat skeletal muscle: adaptation to exercise training. The American journal of physiology 1987;253:C316-322

21. Macarini JR, Maravai SG, Cararo JH, Dimer NW, Goncalves CL, Kist LW, Bogo MR, Schuck PF, Streck EL, Ferreira GC: Impairment of electron transfer chain induced by acute carnosine administration in skeletal muscle of young rats. BioMed research international 2014;2014:632986

of 54


Diabetes

23

22. Fan W, Waizenegger W, Lin CS, Sorrentino V, He MX, Wall CE, Li H, Liddle C, Yu RT, Atkins AR, Auwerx J, Downes M, Evans RM: PPARdelta Promotes Running Endurance by Preserving Glucose. Cell Metab 2017;25:1186-1193 e1184

23. Wang P, Liu J, Li Y, Wu S, Luo J, Yang H, Subbiah R, Chatham J, Zhelyabovska O, Yang Q: Peroxisome proliferator-activated receptor {delta} is an essential transcriptional regulator for mitochondrial protection and biogenesis in adult heart. Circ Res 2010;106:911-919

24. Pereverzev MO, Vygodina TV, Konstantinov AA, Skulachev VP: Cytochrome c, an ideal antioxidant. Biochem Soc Trans 2003;31:1312-1315

25. Morino K, Petersen KF, Shulman GI: Molecular mechanisms of insulin resistance in humans and their potential links with mitochondrial dysfunction. Diabetes 2006;55 Suppl 2:S9-s15

26. Amati F, Dube JJ, Alvarez-Carnero E, Edreira MM, Chomentowski P, Coen PM, Switzer GE, Bickel PE, Stefanovic-Racic M, Toledo FG, Goodpaster BH: Skeletal muscle triglycerides, diacylglycerols, and ceramides in insulin resistance: another paradox in endurance-trained athletes? Diabetes 2011;60:2588-2597

27. Holland WL, Brozinick JT, Wang LP, Hawkins ED, Sargent KM, Liu Y, Narra K, Hoehn KL, Knotts TA, Siesky A, Nelson DH, Karathanasis SK, Fontenot GK, Birnbaum MJ, Summers SA: Inhibition of ceramide synthesis ameliorates glucocorticoid-, saturated-fat-, and obesity-induced insulin resistance. Cell Metab 2007;5:167-179

28. Bergman BC, Brozinick JT, Strauss A, Bacon S, Kerege A, Bui HH, Sanders P, Siddall P, Wei T, Thomas MK, Kuo MS, Perreault L: Muscle sphingolipids during rest and exercise: a C18:0 signature for insulin resistance in humans. Diabetologia 2016;59:785-798

29. Nguyen-Tran DH, Hait NC, Sperber H, Qi J, Fischer K, Ieronimakis N, Pantoja M, Hays A, Allegood J, Reyes M, Spiegel S, Ruohola-Baker H: Molecular mechanism of sphingosine-1-phosphate action in Duchenne muscular dystrophy. Disease models & mechanisms 2014;7:41-54

30. Brehm A, Krssak M, Schmid AI, Nowotny P, Waldhausl W, Roden M: Increased lipid availability impairs insulin-stimulated ATP synthesis in human skeletal muscle. Diabetes 2006;55:136-140

31. Brennan JP, Southworth R, Medina RA, Davidson SM, Duchen MR, Shattock MJ: Mitochondrial uncoupling, with low concentration FCCP, induces ROS-dependent cardioprotection independent of KATP channel activation. Cardiovasc Res 2006;72:313-321

32. Chinopoulos C, Gerencser AA, Mandi M, Mathe K, Torocsik B, Doczi J, Turiak L, Kiss G, Konrad C, Vajda S, Vereczki V, Oh RJ, Adam-Vizi V: Forward operation of adenine nucleotide translocase during F0F1-ATPase reversal: critical role of matrix substrate-level phosphorylation. FASEB journal : official publication of the Federation of American Societies for Experimental Biology 2010;24:2405-2416

of 54


Diabetes

24

33. Baffy G, Derdak Z, Robson SC: Mitochondrial recoupling: a novel therapeutic strategy for cancer? British journal of cancer 2011;105:469-474

34. Mahapatra G, Varughese A, Ji Q, Lee I, Liu J, Vaishnav A, Sinkler C, Kapralov AA, Moraes CT, Sanderson TH, Stemmler TL, Grossman LI, Kagan VE, Brunzelle JS, Salomon AR, Edwards BF, Huttemann M: Phosphorylation of Cytochrome c Threonine 28 Regulates Electron Transport Chain Activity in Kidney: IMPLICATIONS FOR AMP KINASE. The Journal of biological chemistry 2017;292:64-79

35. Han DH, Hancock CR, Jung SR, Higashida K, Kim SH, Holloszy JO: Deficiency of the mitochondrial electron transport chain in muscle does not cause insulin resistance. PLoS One 2011;6:e19739

36. Sparling DP, Griesel BA, Weems J, Olson AL: GLUT4 enhancer factor (GEF) interacts with MEF2A and HDAC5 to regulate the GLUT4 promoter in adipocytes. The Journal of biological chemistry 2008;283:7429-7437

37. Michael LF, Wu Z, Cheatham RB, Puigserver P, Adelmant G, Lehman JJ, Kelly DP, Spiegelman BM: Restoration of insulin-sensitive glucose transporter (GLUT4) gene expression in muscle cells by the transcriptional coactivator PGC-1. Proc Natl Acad Sci U S A 2001;98:3820-3825

38. Lin J, Wu H, Tarr PT, Zhang CY, Wu Z, Boss O, Michael LF, Puigserver P, Isotani E, Olson EN, Lowell BB, Bassel-Duby R, Spiegelman BM: Transcriptional co-activator PGC-1 alpha drives the formation of slow-twitch muscle fibres. Nature 2002;418:797-801

39. Randle PJ, Garland PB, Hales CN, Newsholme EA: The glucose fatty-acid cycle. Its role in insulin sensitivity and the metabolic disturbances of diabetes mellitus. Lancet (London, England) 1963;1:785-789

40. Pu J, Peng G, Li L, Na H, Liu Y, Liu P: Palmitic acid acutely stimulates glucose uptake via activation of Akt and ERK1/2 in skeletal muscle cells. J Lipid Res 2011;52:1319-1327

41. Giacco F, Brownlee M: Oxidative stress and diabetic complications. Circ Res 2010;107:1058-1070

42. Souto Padron de Figueiredo A, Salmon AB, Bruno F, Jimenez F, Martinez HG, Halade GV, Ahuja SS, Clark RA, DeFronzo RA, Abboud HE, El Jamali A: Nox2 mediates skeletal muscle insulin resistance induced by a high fat diet. The Journal of biological chemistry 2015;290:13427-13439

43. Lee HY, Lee JS, Alves T, Ladiges W, Rabinovitch PS, Jurczak MJ, Choi CS, Shulman GI, Samuel VT: Mitochondrial-Targeted Catalase Protects Against High-Fat Diet-Induced Muscle Insulin Resistance by Decreasing Intramuscular Lipid Accumulation. Diabetes 2017;66:2072-2081

of 54


Diabetes

25

44. Biswas G, Adebanjo OA, Freedman BD, Anandatheerthavarada HK, Vijayasarathy C, Zaidi M, Kotlikoff M, Avadhani NG: Retrograde Ca2+ signaling in C2C12 skeletal myocytes in response to mitochondrial genetic and metabolic stress: a novel mode of inter-organelle crosstalk. The EMBO journal 1999;18:522-533

45. Gan Z, Burkart-Hartman EM, Han DH, Finck B, Leone TC, Smith EY, Ayala JE, Holloszy J, Kelly DP: The nuclear receptor PPARbeta/delta programs muscle glucose metabolism in cooperation with AMPK and MEF2. Genes Dev 2011;25:2619-2630

46. Kleiner S, Nguyen-Tran V, Bare O, Huang X, Spiegelman B, Wu Z: PPAR{delta} agonism activates fatty acid oxidation via PGC-1{alpha} but does not increase mitochondrial gene expression and function. The Journal of biological chemistry 2009;284:18624-18633

47. Finck BN, Bernal-Mizrachi C, Han DH, Coleman T, Sambandam N, LaRiviere LL, Holloszy JO, Semenkovich CF, Kelly DP: A potential link between muscle peroxisome proliferator- activated receptor-alpha signaling and obesity-related diabetes. Cell Metab 2005;1:133-144

48. Koves TR, Li P, An J, Akimoto T, Slentz D, Ilkayeva O, Dohm GL, Yan Z, Newgard CB, Muoio DM: Peroxisome proliferator-activated receptor-gamma co-activator 1alpha-mediated metabolic remodeling of skeletal myocytes mimics exercise training and reverses lipid-induced mitochondrial inefficiency. The Journal of biological chemistry 2005;280:33588-33598

49. Nahle Z, Hsieh M, Pietka T, Coburn CT, Grimaldi PA, Zhang MQ, Das D, Abumrad NA: CD36-dependent regulation of muscle FoxO1 and PDK4 in the PPAR delta/beta-mediated adaptation to metabolic stress. The Journal of biological chemistry 2008;283:14317-14326

50. Wende AR, Huss JM, Schaeffer PJ, Giguere V, Kelly DP: PGC-1alpha coactivates PDK4 gene expression via the orphan nuclear receptor ERRalpha: a mechanism for transcriptional control of muscle glucose metabolism. Mol Cell Biol 2005;25:10684-10694

51. Hansford RG, Hogue BA, Mildaziene V: Dependence of H2O2 formation by rat heart mitochondria on substrate availability and donor age. J Bioenerg Biomembr 1997;29:89-95

52. Votyakova TV, Reynolds IJ: DeltaPsi(m)-Dependent and -independent production of reactive oxygen species by rat brain mitochondria. J Neurochem 2001;79:266-277

53. Skulachev VP: Membrane-linked systems preventing superoxide formation. Biosci Rep 1997;17:347-366

54. Yamaguchi S, Katahira H, Ozawa S, Nakamichi Y, Tanaka T, Shimoyama T, Takahashi K, Yoshimoto K, Imaizumi MO, Nagamatsu S, Ishida H: Activators of AMP-activated protein kinase enhance GLUT4 translocation and its glucose transport activity in 3T3-L1 adipocytes. Am J Physiol Endocrinol Metab 2005;289:E643-649

of 54


Diabetes

26

FIGURE LEGENDS

Figure 1. TFAM overexpression attenuates high fat diet induced fat accumulation and loss

of insulin sensitivity. (A-C) mtDNA, mRNA and protein expression of human TFAM in

transgenic (Tg) and wild type (Wt) mice was measured (n=6 per group). (D-E) Body composition

of Wt and Tg mice on chow or high fat diet (HFD; n=11-15 per group). (F) Tg mice had lower fat

accumulation with HFD when compared to Wt. (G-H) Metabolic rate, VO2 and VCO2 of Wt and

Tg mice on Chow or HFD were measured via an open cage calorimetry system (n=5-9 per group).

(I) Activity of Wt and Tg mice on HFD was measured via infrared beam interruption method

(n=3-5 per group). (J-M) Whole body glucose uptake, disposal capacity, and blood glucose in Tg

and Wt mice were measured with euglycemic clamp (n = 8-12 per group). All values were shown

as mean±SEM. Student’s t-test was utilized for all comparisons. *stands for p<0.05, ** stands for

p<0.01, *** stands for p<0.001, **** stands for p<0.0001.

Figure 2. TFAM increases skeletal muscle’s mitochondrial fatty acid oxidative capacity and

energy metabolism. (A-D) Mitochondrial were isolated from fresh quadriceps muscle. Oxygen

consumption and respiratory control ratio (RCR) of OxPhos were measured using various

substrates (n=10 per group). G=glucose, M=malate, PCM=palmitoyl carnitine and malate. (E-U)

Metabolites and proteins were extracted from gastrocnemius muscle (n=6 per group) and

analyzed appropriately. (E-H) ADP and AMP increased with TFAM regardless of diet. ATP

decreased with TFAM overexpression. NAD+ decreased with TFAM under HFD conditions. (I-

K) Creatine (Cr), creatine phosphate (CrP), and creatinine (creatine phosphate shuttle substrates)

significantly changed with TFAM overexpression. (L-M) Citrate synthase protein abundance and

mitochondrial citrate synthase (CS) activity increased with TFAM. (N-U) Citric acid cycle and

of 54


Diabetes

27

glycolytic intermediates, substrates, and enzyme activities increased with TFAM supporting

higher TCA cycle activity and glycolysis. (V) Carnosine, a suppressor of energy metabolism, also

decreased with TFAM overexpression. All values were shown as mean±SEM. Student’s t-test or

two-way ANOVA with Tukey’s correction was utilized for all statistical analyses. * stands for

p<0.05. ** stands for p<0.01, *** stands for p<0.001, **** stands for p<0.0001.

Figure 3. TFAM overexpression increases anti-oxidant defense and reduces reactive oxygen

species. (A-C) Mitochondrial reactive oxygen species (mtH2O2), 8-Oxo-2'-deoxyguanosine (8-

oxo-dG), glutathione (GSH), and glutathione disulfide (GSSG) were measured from freshly

isolated quadriceps muscle (n=6-11 per group). GSSG/GSH ratio indicates redox status where a

lower number indicates lower oxidative stress. (D-E) Superoxide dismutase (SOD2), catalase,

Cytochrome C [Cyt C], phospho-AMPK [pAMPK], total-AMPK [tAMPK] and PPAR protein

expression was measured from quadriceps muscle (n = 6 per group). Antioxidant enzyme

abundance increased in Tg mice. (F) TFAM overexpression in myotybes reduced palmitate (PM)

induced H2O2 emission (n = 6 per group). EV stands for empty vector. (G) TFAM overexpression

in myotubes increased expression of antioxidants (SOD2 and catalase) and GLUT4 with PM

treatment (n = 6 per group). (H) TFAM reduced palmitate induced cleavage of caspase 3 in

myotubes (n = 6 per group). (I) TFAM and EV myotubes were treated with sham or PM (n = 6

cells per group). Mitochondrial and cytosolic abundance of Cyt C was measured. TFAM

overexpression blocked PM mediated release of Cyt C into cytosol. (J) Myc-tagged constitutively

active AMPK (Myc-CA-AMPK) was overexpressed in myotubes, immunopreciptated and probed

for Cyt C binding. (K) TFAM overexpression reduces mtH2O2 by inducing antioxidant enzymes

of 54


Diabetes

28

SOD2 and catalase as well as blocking cytosolic release of Cyt C to trigger apoptosis. All values

were shown as mean±SEM. Student’s t-test or two-way ANOVA with Tukey’s correction was

utilized for all statistical analyses. * stands for p<0.05. ** stands for p<0.01, *** stand for

p<0.001.

Figure 4. TFAM increases lipid -oxidation and reduces insulin resistance associated

bioactive lipids. Metabolites were extracted from gastrocnemius muscle and proteins were

extracted from quadriceps muscle (n=6 per group). (A-H) Car stands for carnitine. (I) LCAD

stands for Acyl-CoA dehydrogenase, long chain. (J-O) Sphingosine (SPH), sphinganine (SPA),

sphingosine-1-phosphate (S1P) and various long chain ceramide species were measured in

gastrocnemius muscle (n=6 per group). Cer stands for ceramide. (P) TFAM increased oxidation

of fatty acids via -oxidation and reduced abundance of Cer18, a bioactive lipid associated with

skeletal muscle insulin resistance. All values were shown as mean±SEM. Student’s t-test with

Tukey’s correction was utilized for all statistical comparisons. * stands for p<0.05, ** stands for

p<0.01, *** stands for p<0.001, **** stands for p<0.0001.

Figure 5. TFAM overexpression asymmetrically changes electron transport chain enzymes

in a posttranslational manner and prevents fatty acid induced uncontrolled uncoupling.

Quadriceps muscle was utilized for both protein and mRNA analyses. (A) Protein expression of

key mitochondrial enzymes (ND dehydrogenase [ND], succinate ubiquinone oxidoreductase

[SUO], cytochrome c oxidoreductase core I and II subunit [Core1 and 2], cytochrome c oxidase

subunit I and IV [COX1 and COX4], ATP synthase α [ATPsyn], and uncoupling protein 3

[UCP3]) were measured in Wt and Tg mice under Chow or HFD conditions (n = 6 muscle per

of 54


Diabetes

29

group). (B) Proteins that decreased with TFAM overexpression had higher ubiquitination

(measured via ubiquitin immunoprecipitation). (C) mRNA expression of protein in panel A were

measured (n = 8 per group). ND gene [NDFAU9], SUO gene [SDHB], Core1 and 2 gene

[UQCRC1 and 2], COX1 gene [MT-CO1], COX4 gene [COX4], ATPsyn gene [ATP5A1], UCP3

gene (UCP3). Upper band is 18s and bottom band is mRNA of the gene. (D) TFAM overexpressed

or empty vector (EV) myotubes were preloaded with TMRM and incubated with either glucose (GLUC)

or palmitate (PALM). Cells were imaged every 10secs. 2M FCCP was added at 180secs (t180) and cells

were rested for 120secs (t300). 8M FCCP was added at t300 and cells were rested for 180secs (t480).

Differences in the TMRM dye intensity from baseline (t0) were computed at resting (t180-t0/t0), 2M FCCP

(t300-t0/t0) and 8M FCCP (t480-t0/t0). A total of 13 cells were used per group. (E) Images taken before t180,

t300 and t480. (F-G) Time resolved changes in the TMRM intensities. (H) TFAM decreased complex I, II

and IV and increased complex II and V, resulting in mild uncoupling and prevention of fatty acid induced

mitochondrial membrane potential (Δψm) depolarization. All values were shown as mean±SEM. A two-

way ANOVA with Tukey’s correction was used for statistical comparisons. * stands for p<0.05, ** stands

for p<0.01, *** stands for p<0.001, **** stands for p<0.0001.

Figure 6. TFAM protects muscle against HFD induced insulin resistance via AMPK and

PPARβ. (A) TFAM protected mice against HFD induced impairment of AKT phosphorylation

(pAKT; n = 6 per group). (B) Proteins were measured in quadriceps muscle (n=6 per group) (C)

TFAM preserved GLUT4 expression under HFD conditions by increasing its upstream regulators.

(D-E) Myotubes were incubated in either 2,4-Dinitrophenol [DNP] or DMSO [Ctrl] and proteins

involved in glucose transport and ETC complexes were measured.(F) Mechanistic view of TFAM

in preventing HFD-induced skeletal muscle insulin resistance. All values were shown as

mean±SEM. Statistical tests were performed either with Student’s t-test or two-way ANOVA

of 54


Diabetes

30

with Tukey’s correction wherever appropriate. * stands for p<0.05, ** stands for p<0.01, ***

stands for p<0.001, **** stands for p<0.0001.

of 54


Diabetes

Figure 1

193x204mm (300 x 300 DPI)

of 54


Diabetes

Figure 2

172x166mm (300 x 300 DPI)

of 54


Diabetes

210x128mm (300 x 300 DPI)

of 54


Diabetes

193x125mm (300 x 300 DPI)

of 54


Diabetes

Figure 5

179x200mm (300 x 300 DPI)

of 54


Diabetes

Figure 6

176x195mm (300 x 300 DPI)

of 54


Diabetes

TFAM enhances fat oxidation and attenuates high fat diet induced insulin resistance in skeletal muscle

Jin-Ho Koh1, Matthew L. Johnson1, Surendra Dasari2, Nathan K. LeBrasseur3, Ivan Vuckovic4, Gregory C. Henderson1, Shawna A. Cooper1, Shankarappa Manjunatha1, Gregory N. Ruegsegger1, Gerald I. Shulman5, Ian R. Lanza1 and K. Sreekumaran Nair1,*

1. Division of Endocrinology and Metabolism, Mayo Clinic, Rochester, MN2. Department of Health Sciences Research, Mayo Clinic, Rochester, MN3. Department of Physical Medicine & Rehabilitation, Mayo Clinic, Rochester, MN4. Mayo Clinic Regional Comprehensive Metabolomics Core, Mayo Clinic, Rochester, MN5. Department of Medicine and Cellular and Molecular Physiology, Yale University, New

Haven, CT

*Corresponding authorK. Sreekumaran Nair, M.D., Ph.D.Professor of MedicineDivision of Endocrinology and MetabolismMayo Clinic College of Medicine200 First St SWRochester, MN 55905Tel: 507-255-2415Fax: 507-255-4828Email: [email protected]

of 54


Diabetes

mailto:[email protected]

This document details all the experimental procedures used in this study.

Generation of MCK-Tfam transgenic mice (Tg). The Mayo Clinic Institutional Animal Care and Use Committee approved the study. To construct a single muscle specific human TFAM (TFAM) DNA plasmid, a bluescript vector containing muscle creatine kinase promoter (MCKp) and LPL (Gift from Dr. Yu in Department of Medicine, Columbia University, NY) and another pcDNA 3.1/Hygro (+) vector containing TFAM were individually transformed with DH5α. Transformed vectors were spread on the agar plate and incubated overnight at 37 °C. Each colony was collected and further incubated overnight with LB medium at 37 °C. Plasmids were extracted using miniprep Kit (Qiagen, Hilde, Germany). The bluescript-MCKp-LPL vector was digested using Hind III and Eco RV enzyme (NEB, Ipswich, WA) and purified using gel electrophoresis to extract bluescript-MCKp vector. The pcDNA3.1 TFAM-Hygro vector also was digested using Hind III and Eco RV enzyme (NEB, Ipswich, WA) and purified using gel electrophoresis to extract TFAM cDNA. The bluescript-MCKp vector and TFAM were conjugated using T4 ligase (NEB, Ipswich, WA). Poly-adenylation was added to the construct prior to inserting into TA-cloning vector. Resulting vector containing MCKp- TFMA was transformed using DH5α.Transformed vector was spread on the agar plate and each colony was collected and further incubated overnight with LB medium at 37°C. Plasmid was extracted using Miniprep Kit (Qiagen, Hilde, Germany).

To generate MCKp-TFAM Tg mice, TA-MCKp-TFAM was injected into pronucleus of single-celled FVB embryos prior to implantation into a pseudopregnant mother. Sexually mature F1 progeny were derived and their TFAM genotype was verified via PCR-based genotyping. Tissue distribution of TFAM expression was assessed for skeletal muscle, heart, liver, kidney, intestine, brain, white adipose tissue, and brown adipose tissue via RT-PCR and Northern Blot (Fig. 1A-C). A total of 6 lines were identified based on the tissue expression of TFAM and utilized for all further analyses.

Animals. Male and female TFAM Tg mice and Wt controls (3 months of age) were used for all studies. Mice were housed individually in a temperature-controlled facility with a 12:12-h light-dark cycle. Mice were randomly assigned to one of two groups for 12-weeks: normal chow diet (CHO: 30% kcal protein, 57% kcal carbohydrate, 13% kcal fat) or high fat diet (HFD: 20% kcal protein, 20% kcal carbohydrate, 60% kcal fat). Food was purchased from Dyets, Inc. (Bethlehem, PA). Food and water were provided ad libitum and food consumption was recorded daily by weighing food bowls and accounting for waste.

Animal body composition. Mice underwent body composition testing by EchoMRI (Echo Medical Systems, Houston, TX) and dual energy X-ray absorptiometry using a Lunar PIXImus™ densitometer (GE Medical-Lunar, Madison, WI). Mice were fasted overnight and weighed. Each mouse was placed into a clear plastic holder to secure it in the device during the one minute reading. Restraint device was made from two telescoping acrylic tubes with an ultra-high molecular weight (UHMW) plug at the end of both tubes. In preparation for DEXA scanning, mice were sedated using a combination of ketamine/xylazine adjusted for body mass. The goal of sedation was to minimize extremity movement, but breathing ability was maintained. Further doses of ketamine (maintenance dosage) were administered as needed intraperitoneally. Respiratory rate prior to the initial dose of ketamine/xylazine was measured. During the sedation,

of 54


Diabetes

if the respiratory rate dropped by more than 20% (regardless of the extent of mouse movement), the sedation was withheld.

Whole body metabolic outcomes and physical activity measurement. An 18-chamber open-circuit cage calorimeter system (CLAMS, Columbus Instruments, Columbus, OH) was used to measure oxygen consumption (VO2) and carbon dioxide production (VCO2) of individual mice, as previously described1. Indirect calorimetry was performed in a constant light (12:12h light:dark phases) and temperature (23°C) controlled room. Oxygen and carbon dioxide analyzers were calibrated with certified primary standard calibration gas (20.49% oxygen, 5,031 ppm carbon dioxide, and nitrogen balanced). Mice were then acclimated to the chambers for 12h followed by the start of the 48h experimental period at 6 AM. Measurements were conducted for 24h each of fasted and fed (ad libitum) conditions. The respiratory exchange ratio (RER) was calculated from VO2 and VCO2 values. Energy expenditure was calculated using the equation [(3.815 1.232 RER) VO2]. Food intake was measured as part of the cage calorimetry experiments using an integrated balance accounting for spillage of food. The ambulatory activity levels of mice were measured using an infrared photocell beam interruption method.

Insulin tolerance tests. Insulin tolerance tests were performed two days following the physical activity assessment. Following a 6h fast, the tail was cut to obtain a drop of blood for a 0 time point glucose measurement with a glucometer. 0.75 U/kg body weight of insulin was injected intrapertonially. Additional blood glucose measurements were collected at 15, 30, 60, and 120 minutes following the insulin injection. If glucose levels fell below 20 mg/dl, the mouse was injected intrapertonially with 50 mg of glucose and given food to prevent hypoglycemic shock induced convulsions or coma.

Insulin sensitivity measurement. Five days before the start of the experiment, catheter was placed in the right jugular vein (silastic 0.025 OD) for infusions of insulin, and 20% dextrose. Mice were fasted for 6h prior to surgery. Animals were placed under tribromoethanol (Avertin, 125-250 mg/kg IP) anesthesia throughout the duration of the surgical procedure. Mice received 100-300 mg/kg acetaminophen added to the water bottles starting 48h prior to surgery. The mice were prepped for surgery by clipping the fur on the ventral neck from the thoracic inlet to the chin, and the dorsum extending from the caudal skull to the mid thorax. Ophthalmic ointment was placed in the eyes and the areas shaved previously were disinfected with surgical iodine and 70% ethanol. The mice were placed in dorsal recumbency and a 7-9 mm skin incision was made lateral of midline over the right jugular vein. A 5-7mm section of the vein was isolated from surrounding tissue by blunt dissection, and small transverse incision was made in vein. The catheter then was inserted into the lumen and secured in place with two 6-0 silk ligatures. A grooved director was used to tunnel subcutaneously from the incision site to the intrascapular area. Using an 18g 1 ½" needle, an incision was made and the free end of the catheter was guided along the grooved director through the incision and secured using two wound clips. The catheter was flushed daily with ~50uL saline containing 200units/mL heparin and 5mg/mL ampicillin. Mice were weighed daily and any mouse not within 10% of their pre-surgery weight by day 5 was removed from the study. On study day, mice were fasted for 5h and Micro-Renathane tubing (0.033 OD) was connected to the catheter leads and infusion machines 1h before the start of the experiment. At -5 minutes blood was collected (~50uL in heparinized microhematocrit tubes) from the tip of the tail to check basal glucose and insulin levels. The insulin clamp began at t=0 minutes with a primed continuous infusion of human insulin

of 54


Diabetes

(16mU/kg bolus followed by 2.5mU/kg/min Humulin R) continued until t=120 minutes. Every 10 minutes starting at t=0 a blood sample was collected (~0.3uL) from the cut tail for blood glucose measurements using the AlphaTrak glucometer (AlphaTrak; Abbott Laboratories, Abbott Park, IL). Euglycemia (120-130 mg/dl) was maintained during the clamp by infusing 20% dextrose as necessary over the course of the experiment. At t=100 and t=120 minutes a 50uL blood sample was collected to determine insulin levels. During clamp mice was allowed to run freely in a cage and was observed throughout the entire 120 minute duration of the clamp. After clamp is over mice was euthanized by CO2.

Blood and tissue collection. After the DEXA scan, animals were given a non-recovery dose of pentobarbital (100 mg/kg) via an intraperitoneal injection. Blood was obtained by cardiac puncture and tissues (including skeletal muscle) were harvested for biochemical studies. At this point, the mice were roughly 4 months of age.

Hydrogen peroxide emission in mice muscle. H2O2 was determined using an Amplex Red oxidation as described previously2. The reactive oxygen species (ROS) emitting potential of isolated mitochondria was evaluated under state 2 conditions. A Fluorolog 3 (Horiba Jobin Yvon, Edison, NJ) spectrofluorometer with temperature control and continuous stirring was used to monitor Amplex Red (Invitrogen, Carlsbad, CA) oxidation in a 50µL aliquot of the freshly isolated mitochondrial suspension. Isolated mitochondria were placed in a quartz cuvette with 2 ml of buffer z containing (in mM) 110 K-MES, 35 KCl, 1 EGTA, 5 K2HPO4, 3 MgCl2-6H2O, and 5 mg/ml BSA (pH 7.4, 295 mOsm). Amplex Red oxidation was measured in the presence of glutamate (10 mM), malate (2 mM), and succinate (10 mM). The fluorescent signal was corrected for background autooxidation and calibrated to a standard curve. H2O2 production rates were expressed per tissue wet weight and per protein concentration of the mitochondrial suspension.

Mitochondrial Function. High-resolution respirometry was performed with freshly isolated muscle mitochondria as previously described3. Approximately 50 mg of muscle tissue was homogenized and mitochondria were separated using differential centrifugation. Mitochondria were added to a 2 ml chamber (Oxygraph-2K, Oroboros) and allowed to equilibrate. Glutamate (10 mM) and malate (2 mM) were added to stimulate State 2 respiration specific to Complex I followed by adding ADP at saturating concentrations (2.5 mM) to induce State 3 respiration of Complex I. Cytochrome C (Cyt C) was added to verify mitochondrial membrane integrity. Succinate (10 mM) was added to stimulate State 3 respiration through Complex I+II. State 4 respiration (Leak) was induced by addition of oligomycin (2µg/µl) and the proton gradient was dissipated by sequential titration of 0.05 mM carbonylcyanide-4-(trifluoromethoxy)-phenylhydrazone (FCCP) to induce uncoupled respiration. Protein content of isolated mitochondrial was determined using a commercially available kit (DC Protein Assay, Bio-Rad). Mitochondrial respiration was normalized to tissue wet weight (reflective of mitochondrial content) and mitochondrial protein (reflective of mitochondrial protein quality).

Determination of mRNA. mRNA was determined using semi-quantitative RT-PCR as described previously4. Following primers were obtained from Integrated DNA Technology (Coralville, Iowa):

of 54


Diabetes

1. PGC-1α: 5′-GTGCAGCCAAGACTCTGTATGG-3′ (forward), 5′-GTCCAGGTCATTCACATCAAGTTC-3′ (reverse)

2. CS: 5’-TGGTCCCAGGATACGGTCAT-3’ (forward), 5’-ACTGTGAGCGTCTACGTTGG-3’ (reverse)

3. NDFAU9: 5’-TTCGGGTTGTTAGAGCGCTT-3’ (forward), 5’- TGTGACCCCATTCGTCCAAG-3’ (reverse)

4. SDHB: 5’-CAGGAGGGCAAGCAACAGTA-3’ (forward), 5’-CAGGCGTTCCTCTGTGAAGT-3’ (reverse)

5. UQRCR1: 5’-ACATTGGTCGCAGTCTCCTG-3’ (forward), 5’-CGGATCCGGTTGTAGTCTGG-3’ (reverse)

6. UQRCR2: 5’-TCAGAGGGCTTCCTGAGTGA-3’ (forward), 5’-AACTCGTCGAGAAAAGGCGT-3’ (reverse)

7. CYCS: 5’-GCTCTTGAGTCCTCTCCAGC-3’ (forward), 5’-AGAAATCCACCTGCCTCTGC-3’ (reverse)

8. MT-CO1: 5’-AGAAGCAGGAGCAGGAACAG-3’ (forward), 5’-CTGTGTTATGGCTGGGGGTT-3’ (reverse)

9. COX4/1: 5’-CAGAAGGCGCTGAAGGAGAA (forward), 5’-ATCAGAACGAGCGCAGTGAA-3’ (reverse)

10. ATP5A: 5’-TCCGAGAAGCTGCAAGGATG-3’ (forward), 5’-TCAACAGACGTGTCAGCTCC-3’ (reverse)

11. UCP3: 5’-TGCTGAAGATGGTGGCTCAG-3’ (forward), 5’-CGCTTTGTTCTGTTCCAGGC-3’ (reverse).

Digital images of mRNA were captured using a C-DiGit blot scanner (Li-COR Bioscience, Lincoln, NE). Transcript intensity was normalized using 18S (Ambion, Austin, TX).

Western blot analysis. We utilized previously described methods5. Frozen tissues were powdered and homogenized in a 15:1 (v/w) ratio of ice-cold buffer containing 50 mM Tris-HCl (pH 7.4), 150 mM NaCl, 1 mM EDTA, 10% Nonidet P-40, 0.5 M NaF, 10 mM Na pyrophosphate, 0.25% sodium deoxycholate, and a protease inhibitor (Thermo Fisher Scientific, Houston, TX). Homogenates were frozen and thawed 3 times to disrupt the mitochondria and re-homogenized. Homogenates were centrifuged at 1000g for 15 min at 4°C and supernatant was collected. Protein concentration was measured by a previously described method6 and sample volumes were adjusted to give the same protein concentration. Aliquots of supernatant were solubilized in Laemli buffer, proteins were separated by SDS-PAGE and transferred to nitrocellulose membranes as described previously7. lots were probed with antibodies against PPARβ, ATP synthase subunit α (ATP synth α), succinate-ubiquinone oxidoreductase (SUO), NADH ubiquinone oxidoreductase (NADH), ubiquinone-cytochrome c oxidoreductase core subunit 1 (Core1) and 2 (Core2), cytochrome c oxidase subunit (COX) I and IV (Thermo Fisher Scientific, Houston, TX), PGC-1α and UCP3 (EMD Millipore, Billerica, MA), c-Myc-Tag, phosphor S473-AKT, phosphor T308-AKT, AKT (pan), AMPK and phospho AMPK (Cell Signaling Technologies, Danvers, MA), GLUT4 (a gift from Mike Mueckler), PPARα, SOD2, Catalase, NRF-1 and MEF-2A (Santa Cruz Biotechnology, Santa Cruz, CA), CaMKKβ (Bethyl, Montgomery, TX), citrate synthase (CS; Alpha Diagnostics, San Antonio, TX), cytochrome c (Cyt c; BD Biosciences, San Jose, CA). Blots were incubated with an appropriate horseradish-conjugated secondary antibody (Jackson Immunoresearch Laboratories, West Grove, PA).

of 54


Diabetes

Antibody bound protein was detected by ECL. Digital images of protein were captured using a C-DiGit blot scanner (Li-COR Bioscience, Lincoln, NE).

Oxidative damage. DNA oxidative damage was measured using the 8-oxo-2’deoxyguanosine (8-oxo-dG) adduct biomarker in the muscle homogenate as previously described8. Protein oxidative damage was measured as the content of protein carbonyls in muscle homogenate using the Oxi-select Protein Carbonyl Immunoblot kit (Cell Biolabs, San Diego, CA) following the manufacturer’s guidelines. Proteins were separated by SDS-PAGE and transferred to polyvinylidene fluoride membrane (PVDF). After dinitrophenylhydrazine (DNPH) derivatization of oxidized protein on PVDF, membrane was incubated with Anti-DNP and secondary antibody. Digital images of protein carbonyl content were captured using a C-DiGit blot scanner (Li-COR Bioscience, Lincoln, NE).

Fatty acid preparations. Palmitate was conjugated with the fatty acid free BSA9. Briefly, 200 mM sodium palmitate was dissolved in 50% ethanol by heating at 55 °C and vortexing until dissolved. Which 200 mM sodium palmitate were diluted 25-tiems in a 10% fatty acid free low-endotoxin BSA solution (Sigma, St. Louis, MO) to achieve a final molar ration of 5:1. This medium was incubated at 40°C for 2h. Conjugated BSA-palmitate were diluted 40-fold in cell culture media to reach a final concentration of 0.2 mM palmitate. Control BSA was prepared by adding the same amount of 50% ethanol into a 10% BSA solution. All preparations were aliquoted and frozen at -20°C.

TFAM myotube cultures. C2C12 myoblasts were grown in DMEM (4.5g glucose/L, Sigma, St. Louis, MO) containing 10% fetal bovine serum, 100 μU/m/penicillin and 100 µU/ml streptomycin, GlutaMAX (Thermo Fisher Scientific, Houston, TX), and differentiation was initiated by switching to medium containing 2% heat inactivated horse serum when the myoblasts were 90% confluent. Adenovirus bearing human TFAM (AD- TFAM), which was purchased from Applied Biological Materials Inc. (British Columbia, Canada), with or without 0.2mM palmitate were infected into myotube. Myotubes were harvested after 5 days of differentiation.

Constitutively activate AMPK (CA-AMPK) mytotube cultures. Differentiation was induced in the C2C12 myoblasts as described above. Adenovirus bearing myc tagged constitutively active AMPK (pAd-Track/myc-AMPK-CAα, myc-AMPK-CA, a gift from John O Holloszy’s Lab) was overexpressed for 7 days in myotubes and extracts of the myotubes were prepared for western blotting.

Muscle electroporation. Constitutively activate (CA)-AMPK (pcDNA3.1-V5-His-AMPK-CAα, CA-AMPK) plasmid DNA was gift from John O Holloszy’s Lab (Washington University School of Medicine). Plasmic DNA was transfected into mouse tibialis anterior muscle using a previously described electric pulse mediated gene transfer technique (Koh, Hancock, Terada, Higashida, Holloszy, & Han, 2017b). For this, mice were anesthetized with isoflurane gas. A tibialis anterior muscle was injected with 50 µg of plasmid DNA containing either myc-CA-AMPK, or empty vector (EV) in 50 µL with GFP saline using a 27gauge needle at a rate of 20µl/min. After injection, an electric field was applied to the muscles (Koh, Hancock, Terada, Higashida, Holloszy, & Han, 2017b). Twenty one days after the transfection, animals were anesthetized with an intraperitoneal injection of sodium pentobarbital and muscles were

of 54


Diabetes

dissected out, frozen in liquid nitrogen and kept at -80°C until analyzed. Anesthetized mice were euthanized by exsanguination.

Immunoprecipitation studies. Anti-Myc (Sigma, St. Louis, MO) or anti-ubiquitin (Thermo Fisher Scientific, Houston, TX) antibody with magnetic beads (EMD Millipore, Billerica, MA) were mixed at RT for 1.5hr. Aliquots 250 mg of protein from each group were added in mixture of antibody and beads, and then rotated at 4°C overnight. Following morning, magnetic beads were washed and protein was eluted from the beads with 5X lane marker reducing sample buffer (Thermo Fisher Scientific, Houston, TX), which was boiled for 5 min. Western blotting was used to measure Cyt C (BD Biosciences, San Jose, CA), NADH (Thermo Fisher Scientific, Houston, TX), Core1 (Thermo Fisher Scientific, Houston, TX) or COX1 (Thermo Fisher Scientific, Houston, TX) with antibody. Digital images of protein were captured using a C-DiGit blot scanner (Li-COR Bioscience, Lincoln, NE).

Hydrogen peroxide emission in myotubes. Palmitate or saline treated TFAM or EV myotybes in 75t flask were washed with DPBS and suspended with 0.25% trypsin-EDTA medium. Myotubes were centrifuged and re-suspend with respiration medium [105 mM K-MES, 30 mM KCL, 10 mM KH2PO4, 5 mM MgCl2-6H2O, 5mg/ml BSA] pH 7.4 supplemented with 1 mM EGTA (pH 7.3) including 10 mM Amplex Red, Horseradish peroxidase 1U/ul, Superoxide dismutase 10U/ul. Five mg/ml digitonin (~50% TLC, Sigma, St. Louis, MO) was added to permeabilize the membranes. Two mmol/L Glutamate (5mM) and 1 mmol/L malate were added to stimulate H2O2 production under state 4 conditions. A fluorolog 3 (Horiba Jobin Yvon, Edison, NJ) spectrofluorometer was used to monitor Amplex Red (Invitrogen, Carlsbad, CA) oxidation in myotubes.

Muscle acylcarnitines. Acylcarnitines (C0, C2, C3, C4, C5, C8, C12, C14, C16, C18, and C18:1) were measured by LC-MS as described previously 10. Analyte concentrations were expressed relative to total protein content of the tissue.

Muscle ceramides. Tissue ceramides (cer14, Cer16, Cer18, Cer24, Cer24:1), sphinganine (SPA), sphingosine (SPH) and sphingosine-1-phosphate (S1P) were measured using LC-MS as previously described11. Analyte concentrations were expressed relative to total protein content of the tissue.

Muscle diacylglcerols. Saturated and unsaturated diacylglycerols (16:0/16:0, 18:0/18:2, 18:0/18:1, 18:1/18:1, 18:2/18:2, 16:0/18:1) were extracted from the gastrocnemius muscle and analyzed using LC-MS as previously decribed12. Analyte concentrations were expressed relative to total protein content of the tissue.

Muscle energy metabolism NMR analysis. ~30 mg of muscle tissue was pulverized and 100µl of 6% HClO4 was added. Tissue was ground for 30s with a hand homogenizer, 200 µl 6% HClO4 was added and the mixture was shaken on vortex and frozen in liquid nitrogen. The sample was thawed and spun down at 10,000xg for 15 minutes. Supernatant was collected and the pellet was re-extracted with 100µl of 6% HClO4. Combined extracts were neutralized with 140µl of 2M KHCO3. The mixture was spun down at 10,000xg for 15 minutes and supernatant (500ul) was collected. 100l of phosphate buffer (pH 7.4) and 50l of TSP-d4 solution in D2O (1mM) were added and sample was transferred to a 5mm NMR tube.

of 54


Diabetes

NMR spectra were acquired on a Bruker 600MHz Avance IIIHD spectrometer equipped with a BBI room temperature probehead and SampleJet auto sampler (Bruker Biospin, Rheinstetten, Germany). 1H NMR spectra were recorded using 1D noesy pulse sequence (noesygppr1d), with calibrated 90° pulse (~11 ms) and following conditions: NS=128; TD=64k; SW=14 ppm; T=298.2K; AQ=3.9s; D1=5s. Spectra were phase and baseline corrected using TopSpin 3.5 software (Bruker, Rheinstetten, Germany). Metabolites were identified and quantified using Chenomx NMR suite 8.3 software (Chenomx, Alberta, Canada) by fitting spectral lines of library compounds into the NMR spectra of tissue extract. Quantification was based on peak area of TSP-d4 signal. Metabolite concentrations were exported as µM in the NMR sample and recalculated as µmol per g of tissue.

Mitochondrial membrane potential (Δψm) measurement using TMRM. To measure Δψm by fluorescent microscopy (Nicon), C2C12 cells were seeded in either 8-well Lab-Tek with No.1 borosilicate glass or 6 well glass bottomed plates (Thermo Fisher, Waltham, MA). Ad-TFAM or empty vector (EV) was introduced to cells after changing differentiation medium. After 5 days, cells were pre-loaded with 100 nM TMRM (tetramethylrhodamine methyl ester, Invitrogen, Carlsbad, CA), and 500 ng/mL Hoechst 33342 (Invitrogen) and incubated for 20min at 37C. Cells were washed three times with DPBS and were maintained in an imaging buffer (Invitrogen) that contained either 1g/L glucose or 0.2mM palmitate. After 2 hour incubation, cells were imaged by Nikon 100X oil immersion lens. To access mitochondrial membrane potential, 2µM and 8 µM (final concentration) FCCP was added by stepwise directly to the cells. Fluorescence intensity of thirteen to eighteen cells in each group was evaluated every 10 seconds using Nikon instruments software (NIS). TMRM intensity was calculate as Δt=(tX-t0)/t0 X 100, where tx=fluorescence intensity at any time point, t0 = baseline fluorescence.

DNP treatment in C2C12After 5 days of cell differentiation, cells were incubated for 24 hour with 3uM (final concentration) DNP or DMSO as control in differentiation medium.

References1. Izumiya, Y., et al. Fast/Glycolytic muscle fiber growth reduces fat mass and improves

metabolic parameters in obese mice. Cell Metab 7, 159-172 (2008).2. Lanza, I.R., et al. Influence of fish oil on skeletal muscle mitochondrial energetics and

lipid metabolites during high-fat diet. Am J Physiol Endocrinol Metab 304, E1391-1403 (2013).

3. Lanza, I.R. & Nair, K.S. Functional assessment of isolated mitochondria in vitro. Methods in enzymology 457, 349-372 (2009).

4. Terada, S., Wicke, S., Holloszy, J.O. & Han, D.H. PPARdelta activator GW-501516 has no acute effect on glucose transport in skeletal muscle. Am J Physiol Endocrinol Metab 290, E607-611 (2006).

5. Koh, J.H., et al. PPARbeta Is Essential for Maintaining Normal Levels of PGC-1alpha and Mitochondria and for the Increase in Muscle Mitochondria Induced by Exercise. Cell Metab 25, 1176-1185 e1175 (2017).

6. Lowry, O.H., Rosebrough, N.J., Farr, A.L. & Randall, R.J. Protein measurement with the Folin phenol reagent. J Biol Chem 193, 265-275 (1951).

7. Baar, K., et al. Skeletal muscle overexpression of nuclear respiratory factor 1 increases glucose transport capacity. FASEB J 17, 1666-1673 (2003).

of 54


Diabetes

8. Lanza, I.R., et al. Chronic caloric restriction preserves mitochondrial function in senescence without increasing mitochondrial biogenesis. Cell Metab 16, 777-788 (2012).

9. Pillon, N.J., Arane, K., Bilan, P.J., Chiu, T.T. & Klip, A. Muscle cells challenged with saturated fatty acids mount an autonomous inflammatory response that activates macrophages. Cell Commun Signal 10, 30 (2012).

10. Chace, D.H., et al. Electrospray tandem mass spectrometry for analysis of acylcarnitines in dried postmortem blood specimens collected at autopsy from infants with unexplained cause of death. Clinical chemistry 47, 1166-1182 (2001).

11. Blachnio-Zabielska, A.U., Persson, X.M., Koutsari, C., Zabielski, P. & Jensen, M.D. A liquid chromatography/tandem mass spectrometry method for measuring the in vivo incorporation of plasma free fatty acids into intramyocellular ceramides in humans. Rapid communications in mass spectrometry : RCM 26, 1134-1140 (2012).

12. Blachnio-Zabielska, A.U., Zabielski, P. & Jensen, M.D. Intramyocellular diacylglycerol concentrations and [U-(1)(3)C]palmitate isotopic enrichment measured by LC/MS/MS. J Lipid Res 54, 1705-1711 (2013).

of 54


Diabetes

Supplemental Figures

Supplemental Figure 1. TFAM overexpression increases mitochondrial-encoded mRNA expression. mRNA expression was assessed in the gastrocnemius (n=6 animals per group). All values were shown as mean±SEM. Student’s t-test was used for statistical comparisons. * stands for p<0.05.

of 54


Diabetes

Supplemental Figure 2. Caloric intake of TFAM and Wt mice on chow or high fat diet. (A-B) Total and average weekly caloric intake was computed for Wt and Tg mice on either chow or high fat diet (n=10-15 animals per group. All values were shown as mean±SEM. Student’s t-test was used for statistical comparisons. * stands for p<0.05, ** stands for p<0.01, *** stands for p<0.001, **** stands for p<0.0001. # symbol was used for asterisk when comparing Tg chow vs. Tg HFD animals.

of 54


Diabetes

Supplemental Figure 3. Respiratory exchange ratio (RER) is unaltered by TFAM overexpression. (A-B) Average RER was computed during the day and night during fed and fasted state for Wt and Tg mice on either chow (A) or high fat diet (B) (n=10-15 animals per group. All values were shown as mean±SEM. Student’s t-test was used for statistical comparisons.

of 54


Diabetes

Supplemental Figure 4. Tg mice on high fat diet had higher nightly activity compared to other groups. Total activity was counted using an infrared photo beam interruption method (n=4-6 animals per group). All values were shown as mean±SEM. Student’s t-test was used for statistical comparisons. * stands for p<0.05.

of 54


Diabetes

Supplemental Figure 5. Insulin dynamics in Tg mice. (A) Wt and Tg mice on either Chow or HFD were fasted overnight and their insulin levels were measured (n=8-12 per group). No differences in fasting insulin levels were observed. (B) Mice were put on euglycemic clamp and insulin levels were measured (n=8-12 per group). Tg mice on HFD had lower levels of insulin on when on clamp when compared to Wt mice on HFD. (C-D) Basal and clamped endogenous glucose production (EGP) was measured in Tg and Wt mice on either Chow or HFD (n=12 animals per group). Tg mice on HFD had higher levels of EGP when compared to Wt mice on HFD. No EGP differences were observed between the animals during the clamped state. All values were shown as mean±SEM. Student’s t-test was used for statistical comparisons. * stands for p<0.05.

of 54


Diabetes

Supplemental Figure 6. Insulin tolerance test. Following a 6hour fast, insulin tolerance test was conducted in Wt and Tg mice on either Chow or HFD (n=5-7 per group). Wt mice on HFD had lower insulin sensitivity (i.e. higher blood glucose) than rest of the groups. Tg mice had same blood glucose levels a1s Wt Chow mice regardless of the diet. All values were shown as mean±SEM. Student’s t-test was used for statistical comparisons. ** stands for p<0.01.

of 54


Diabetes

Supplemental Figure 7. Mitochondrial respiration normalized to tissue weight. Mitochondrial were isolated from fresh quadriceps muscle. Oxygen consumption was measured using carbohydrate (GM) substrates (n=10 per group). Oxygen consumption was normalized to tissue weight. (A-B) State 3 complex I respiration decreased in Tg mice when compared to Wt mice regardless of diet. All values were shown as mean±SEM. Student’s t-test was used for statistical comparisons. ** stands for p<0.01.

of 54


Diabetes

Supplemental Figure 8. TFAM increases glycogen synthase phosphorylation under chow but not HFD conditions. Tissue homogenates from the quadriceps muscle were probed for p-Ser641 glycogen synthase (GS) which was normalized to total GS protein content (n=6 per group). All values were shown as mean±SEM. Student’s t-test was used for statistical comparisons. * stands for p<0.05.

of 54


Diabetes

Supplemental Figure 9. Diacylglycerol levels decreased in muscle with TFAM. Saturated (16:0/16:0) and unsaturated (18:0/18:2, 18:0/18:1, 18:1/18:1, 18:2/18:2, 16:0/18:1) diacylglycerols were extracted from gastrocnemius muscle (n=6 per group) and analyzed using quantitative mass spectrometry. All values were shown as mean±SEM. Student’s t-test was used for statistical comparisons. ** stands for p<0.01.

of 54


Diabetes

TFAM enhances fat oxidation and attenuates high fat diet ......2019/05/03 · 1 TFAM enhances fat...

Documents

Transcript of TFAM enhances fat oxidation and attenuates high fat diet ......2019/05/03 · 1 TFAM enhances fat...