1 TFAM enhances fat oxidation and attenuates high fat diet induced insulin resistance in skeletal muscle Jin-Ho Koh 1 , Matthew L. Johnson 1 , Surendra Dasari 2 , Nathan K. LeBrasseur 3 , Ivan Vuckovic 4 , Gregory C. Henderson 1 , Shawna A. Cooper 1 , Shankarappa Manjunatha 1 , Gregory N. Ruegsegger 1 , Gerald I. Shulman 5 , Ian R. Lanza 1 and K. Sreekumaran Nair 1, * 1. Division of Endocrinology and Metabolism, Mayo Clinic, Rochester, MN 2. Department of Health Sciences Research, Mayo Clinic, Rochester, MN 3. Department of Physical Medicine & Rehabilitation, Mayo Clinic, Rochester, MN 4. Mayo Clinic Regional Comprehensive Metabolomics Core, Mayo Clinic, Rochester, MN 5. Department of Medicine and Cellular and Molecular Physiology, Yale University, New Haven, CT *Corresponding author K. Sreekumaran Nair, M.D., Ph.D. Professor of Medicine Division of Endocrinology and Metabolism Mayo Clinic College of Medicine 200 First St SW Rochester, MN 55905 Tel: 507-255-2415 Fax: 507-255-4828 Email: [email protected]Running Title: Muscle TFAM & insulin resistance Page 1 of 54 Diabetes Diabetes Publish Ahead of Print, published online May 14, 2019
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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
Page 1 of 54 Diabetes
Diabetes Publish Ahead of Print, published online May 14, 2019
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
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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.
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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
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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
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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
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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
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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.
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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.
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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
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
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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.
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Figure 1
193x204mm (300 x 300 DPI)
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Figure 2
172x166mm (300 x 300 DPI)
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210x128mm (300 x 300 DPI)
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193x125mm (300 x 300 DPI)
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Figure 5
179x200mm (300 x 300 DPI)
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Figure 6
176x195mm (300 x 300 DPI)
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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]
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,
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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
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(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):
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).
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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
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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.
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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).
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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).
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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.
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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.
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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.
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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.
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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.
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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.
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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.
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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.
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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.