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Role of PKCδ in Insulin Sensitivity and Skeletal Muscle Metabolism
Mengyao Li1, Sara G.Vienberg
1,2, Olivier Bezy
1,3, Brian T. O’Neill
1, C. Ronald
Kahn1
1Section of Integrative Physiology and Metabolism, Joslin Diabetes Center, Harvard
Medical School, Boston, Massachusetts 02215 USA
2Novo Nordisk Foundation Center for Basic Metabolic Research, Copenhagen University,
Copenhagen, Denmark
3Current Address: Cardiovascular and Metabolic Diseases Research Unit, Pfizer Inc.,
Cambridge, Massachusetts, USA
Address correspondence to:
C. Ronald Kahn, M.D.
Joslin Diabetes Center
One Joslin Place, Boston, MA 02215
Phone: (617) 309-2635.
Fax: (617) 309-2487.
Email: C.Ronald.Kahn@joslin.harvard.edu
Running Title:
PKCδ Regulates Glucose Tolerance and Muscle Insulin Signaling during Aging.
Keywords:
PKCδ, skeletal muscle, insulin signaling, aging
Page 1 of 49 Diabetes
Diabetes Publish Ahead of Print, published online August 25, 2015
2
ABSTRACT
Protein kinase C (PKC) δ has been shown to be increased in liver in obesity and plays an
important role in the development of hepatic insulin resistance in both mice and humans.
In the present study, we explored the role of PKCδ in skeletal muscle in the control of
insulin sensitivity and glucose metabolism by generating mice in which PKCδ was
deleted specifically in muscle using Cre-lox recombination. Deletion of PKCδ in muscle
improved insulin signaling in young mice, especially at low insulin doses, however, this
did not change glucose tolerance or insulin tolerance tests done with pharmacological
levels of insulin. Likewise, in young mice, muscle-specific deletion of PKCδ did not
rescue high-fat diet induced insulin resistance or glucose intolerance. However, with an
increase in age, PKCδ levels in muscle increased, and by 6- to 7-months of age, muscle-
specific deletion of PKCδ improved whole body insulin sensitivity and muscle insulin
resistance, and by 15 months of age, improved the age-related decline in whole body
glucose tolerance. At 15 months of age, M-PKCδKO mice also exhibited decreased
metabolic rate and lower levels of some proteins of the OXPHOS complex suggesting a
role for PKCδ in the regulation of mitochondrial mass at older age. These data indicate an
important role of PKCδ in the regulation of insulin sensitivity and mitochondrial
homeostasis in skeletal muscle with aging.
Page 2 of 49Diabetes
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Introduction
Protein kinase C (PKC) is a family of serine/threonine kinases that play important
roles in many cellular signaling events, including cell growth, differentiation, apoptosis
and hormonal responses. PKCs are classified into three major categories: conventional
PKCs (α, βI, βII, γ), novel PKCs (δ, ε, ν, θ) and atypical PKCs (ζ, ι, λ) (1-3). A number of
PKC isoforms have been implicated in both insulin action (4;5) and insulin resistance
(6;7). Activation of conventional and novel PKCs by insulin, hyperglycemia and lipids,
especially diacylglycerol (DAG), have been shown to lead to insulin resistance (7-9).
Protein kinase C δ (PKCδ) is a member of the novel family of PKC proteins and is
involved in many pathological conditions, including ischemic heart disease (10;11) and
cancer (12).
PKCδ has also been implicated in insulin action and insulin resistance (5;13-17).
While in vitro studies have suggested that PKCδ plays a positive role in insulin-
stimulated glucose uptake in muscle (4;5), animal studies, especially those focusing on
liver, have indicated that PKCδ is a major contributor to hepatic insulin resistance (18).
In previous studies using genome-wide scanning to compare the diabetes/obesity prone
C57BL/6J (B6) mice and diabetes/obesity resistant 129S6/Sv (129) mice, we identified
PKCδ as being strongly linked to the development of insulin resistance (15;16). Mice
with liver-specific reduction in PKCδ gene expression display increased hepatic insulin
sensitivity, improved glucose tolerance and reduced hepatic lipid accumulation, while
mice with liver-specific over-expression of PKCδ develop hepatic insulin resistance, fatty
liver and glucose intolerance (18). However, the contribution of muscle-derived PKCδ in
the development of insulin resistance in vivo has not been explored.
Page 3 of 49 Diabetes
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Skeletal muscle is the predominant site of insulin-stimulated glucose uptake in the
post-prandial state. Insulin resistance in muscle is one of the characteristic features of
type 2 diabetes and has been shown to exist in genetically susceptible individuals, years
prior to the onset of clinical diabetes (19). As noted above, in vitro studies have
suggested a positive role of PKCδ in insulin-stimulated glucose uptake performed in
skeletal muscle cells (2;13;20). Aging is associated with many metabolic changes
including lipid accumulation and the development of insulin resistance (21;22). These
changes, lead to increased prevalence of diabetes and metabolic syndrome beginning in
middle-aged individuals and increasing thereafter. In the current study, we explored the
role of PKCδ on muscle insulin sensitivity in relation to diet and age by generating mice
in which the PKCδ gene has been specifically deleted in skeletal muscle using Cre-lox
recombination. We found that while PKCδ does not appear to play a role in muscle
insulin resistance associated with diet-induced obesity, PKCδ does play a role in the
onset of insulin resistance in muscle as the mice enter middle-age. Thus, PKCδ levels in
muscle increase with age, and muscle-specific deletion of PKCδ improves whole body
insulin sensitivity, reverses whole body glucose intolerance and improves muscle insulin
signaling in middle-aged mice. Hence, PKCδ is a component of insulin resistance as mice
mature/age, suggesting its potential as a therapeutic target for type 2 diabetes.
Page 4 of 49Diabetes
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Methods
Animals. PKCδ-floxed (control) and MCK-Cre transgenic mice have been previously
described (18;23). All mice were housed in a 20-22°C temperature-controlled room on a
12 h-light/dark cycle, and were allowed ad libitum access to water and food. Animals
were maintained on a standard chow diet containing 22% calories from fat (Mouse Diet
9F 5020; PharmaServ), or given a high-fat diet (HFD) containing 60% calories from fat
(OpenSource Diet D 12492, Research Diet). All animal studies were performed
according to protocols approved by the Institutional Animal Care and Use Committee
(Joslin Diabetes Center, Boston).
Metabolic studies. For the glucose tolerance test, mice were fasted overnight and
injected with glucose intraperitoneally (2 g/kg body weight). Intraperitoneal insulin
tolerance tests were performed in the mice after 4 h fasting and insulin was given
intraperitoneally (1 mU/g body weight). Glucose levels were measured in blood collected
from the tail at the indicated times using Infinity Glucose Monitors and strips (US
Diagnostics). Serum insulin levels were assessed by ELISA according to the
manufacture’s recommendation (Crystal Chem).
Body composition and metabolic analysis. Body composition was measured using
DEXA scanning (Lunar PIXImus2 densitometer; GE Medical Systems) following
anesthesia using Avertin (tribromoethanol:tert-amyl alcohol, 0.015ml/g i.p.). Mice were
housed individually and assessed for their metabolic activities using an OPTO-M3 sensor
system (Comprehensive Laboratory Animal Monitoring System, CLAMS; Columbus
Instruments). Food intake, spontaneous activity, volume of oxygen consumption and
volume of CO2 production were measured over a 48 h light and dark cycle.
Page 5 of 49 Diabetes
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Mitochondrial isolation and oxygen consumption rate analysis. Mitochondria were
isolated and purified from hindlimb muscle using a protocol modified from one
previously described (24). Briefly, 0.5 g of hindlimb skeletal muscle was excised and
digested in a buffer containing dispase (1 mg/g muscle weight) and trypsin (10 mg/g
muscle weight) for 10 min at room temperature. The tissue pieces were homogenized
using a Potter-Elvehjem homogenizer and centrifuged at 600 g for 10 min at 4°C. The
supernatant containing the mitochondrial fraction was filtered through cheese cloth and
centrifuged at 14,000 g for 10 min at 4°C. The pellet was resuspended and washed two
more times in wash buffer (100 mM KCl, 50 mM Tris·HCl,1 mM MgCl2, 0.2 mM
EDTA, 0.2 mM ATP, pH 7.4) to get the final mitochondrial pellet. Isolated mitochondria
were diluted in the Mitochondrial Assay Solution (70 mM sucrose, 220 mM mannitol, 10
mM KH2PO4, 5 mM MgCl2, 2 mM HEPES, 1.0 mM EGTA and 0.2% fatty acid-free
BSA, pH 7.2) supplied with 10 mM pyruvate/2 mM malate or 40 µM palmitoyl-
carnitine/0.5 mM malate. Oxygen consumption rate (OCR) was measured using a
Seahorse Bioscience XF24 analyzer in the absence (basal) or presence (state 3) of 400
µM adenosine diphosphate. ADP independent respiration activity (state 4) was measured
in the presence of 2 µM oligomycin. Protein concentrations were measured using the
Bradford method (Bio-Rad).
Hyperinsulinemic-euglycemic clamp. Insulin sensitivity was measured in the mice at 6-
months of age by performing hyperinsulinemic-euglycemic clamp coupled with D-[3-
3H]-glucose and
14C-deoxyglucose infusions as previously described (25). Whole body
insulin sensitivity and muscle specific glucose uptake were measured using a continuous
insulin infusion dose of 5 mU/kg/min and a constant level of blood glucose at 100 mg/dl
Page 6 of 49Diabetes
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during the clamp. Whole body insulin sensitivity was expressed as glucose infusion rate
(GIR), which was determined by the level of exogenous glucose infusion to maintain
blood glucose levels as the initial levels during the clamp. Insulin-stimulated glucose
uptake into the muscle was assessed during the final 45 minutes of the hyperinsulinemic-
euglycemic clamp.
In vivo insulin signaling, western blot and ROS/RNS assay. After 6 hours of fasting,
mice were anesthetized using Avertin (240 mg/kg body weight) and injected with regular
human insulin (Novolin) at the doses of either 200 U/kg or 20 U/kg of body weight via
the inferior vena cava. Muscles were dissected 10 minutes after the insulin bolus and
frozen in liquid nitrogen. For western blot analysis, muscles were homogenized in RIPA
buffer (Thermo Scientific) supplemented with phosphatase inhibitor and protease
inhibitor cocktail (Sigma-Aldrich). Antibodies against phospho-Akt, phospho-ERK,
phospho-IRβ, Akt, ERK and VDAC were purchased from Cell Signaling. Antibodies
against PKCδ, IRβ and GAPDH were purchased from Santa Cruz. The phospho-IRS-1
antibody was purchased from Invitrogen, antibodies against IRS-1 were purchased from
BD Bioscience, and antibodies against OXOPHOS proteins (mitochondrial complex I, II
and IV) were from Abcam. For muscle fractionation, cytoplasmic and membrane protein
were collected from skeletal muscle using a plasma membrane protein extraction kit
(Abcam, ab65400). Muscle oxidative levels were determined by a spectrofluorimetric
method using the OxiSelect in vitro ROS/RNS Assay kit (Cell Biolabs, STA-347).
Ex vivo Glucose uptake. Muscle glucose uptake was measured as described previously
(26;27). Briefly, EDL and soleus muscles were dissected from 7-month old control and
M-PKCδKO mice and incubated in Krebs-Ringer bicarbonate buffer (KRB) that contains
Page 7 of 49 Diabetes
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2mM of pyruvate for 1 hour in the presence or absence of 2.5mU/ml of insulin. Muscles
were then transferred to transport solution in KRB buffer with 7mM 14
C-D-mannitol and
1mM 3H-2-deoxy-glucose (2-DG), with or without insulin. Accumulation of
3H-2-DG
was determined, and rates of uptake were calculated as described previously (26).
Exercise capacity and muscle grip strength tests. Maximal exercise tolerance was
measured in 7-month old mice using a treadmill running protocol, modified from the
previously described assay (28). In brief, mice were given 30 minutes to acclimate to the
treadmill (Columbus Instruments, Columbus, OH). They then exercised at 5 m/min at 00
incline for 5 min, and after each interval (5 min) the speed was increased by 5 m/min
until reaching 20 m/min with 00 incline. The slope was then increased 5 degrees every 5
min, while maintaining the speed at 20 m/min until the last mouse reached the exhaustion
point. Maximal exercise tolerance was determined by the cumulative amount of work
(kJ) that each mouse performed, calculated as body weight (kg) × vertical distance
covered (m) × 9.81. Muscle strength of the forelimbs of each mouse was measured using
a Grip Strength Meter (Columbus Instruments, Columbus, OH) as described previously
(29). In brief, each mouse was held from the tip of its tail and the front paws grasped the
grid. The grip was released when the mouse was pulled back gently. Hind limbs were
kept free during the test. Each animal was tested 3 times with a 5 minute break between
each measurement.
Muscle fiber purification. Mice were perfused with saline via the left ventricle. Single
tibialis anterior (TA) muscle fibers were isolated with fine forceps under a bright-field
microscope following perfusion. Isolated fibers were homogenized and processed as
described above, and PKCδ protein expression was measured by western blot.
Page 8 of 49Diabetes
9
Gene expression analysis. Total RNA was extracted from tissues using Trizol. 1 µg of
RNA was reverse-transcribed using a high-capacity complementary DNA reverse
transcription kit (Applied Biosystems) according to the manufacture’s instructions.
Quantitative realtime PCR was performed in 5 µl of the resulting cDNA after a 10-fold
dilution in the presence of the SYBR Green PCR Master Mix (Applied Biosystems) and
300 nM primers. PCR reactions were run in duplicate in the ABI Prism 7700 Sequence
Detection System, and Ct values were normalized to GAPDH gene levels. PKCδ primer
sequence: F: 5’-CAGCCTTTCTGTGCTGTGAA-3’; R: 5’-
CTGGATAACACGGCCTTCAT-3’. GAPDH primer sequence: F: 5’-
TGTCGTGGAGTCTACTGGTGTCTT -3’; R: 5’-
TCTCGTGGTTCACACCCATCACAA -3’.
Statistics. Data are expressed as mean ± SEM. Differences were analyzed using an
unpaired Student’s t test, One-way ANOVA or two-way ANOVA. One-way ANOVA
was followed by the Tukey's Multiple Comparison post-hoc test, and two-way ANOVA
was performed using Bonferroni’s Multiple Comparison post-test. Statistical calculations
were performed using the GraphPad Prism software (GraphPad, San Diego, CA). A
probability value of < 0.05 was considered significantly different.
Page 9 of 49 Diabetes
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Results
Muscle PKCδ expression increases in B6 mice as they age but not in response to
high-fat diet. Aging and obesity are two major causes of insulin resistance and diabetes
(30). We first assessed if PKCδ expression was regulated in mice by dietary insult or
during aging. PKCδ protein levels in muscle increased significantly in an age-dependent
manner, increasing to 165% and 170% of 10-week old level at 8-months and 15-months
of age respectively (Figure 1A). In contrast to our previous studies which showed
increased hepatic PKCδ expression in response to HFD (18), muscles from mice
challenged with a HFD for 10 weeks displayed a reduction in PKCδ protein levels by
50% (Figure 1B and C).
Mice with muscle-specific PKCδ deletion display improved insulin signaling at 10-
weeks of age. Previous in vitro studies suggested an important role for PKCδ in insulin
stimulated glucose uptake in muscle (4;5), while in vivo studies indicated a role for PKCδ
in liver leading to insulin resistance (15-17). To assess the true metabolic role of PKCδ in
muscle in vivo, we generated a mouse in which the PKCδ gene was deleted specifically in
muscle by breeding mice in which exon 2 of PKCδ was flanked with loxP sites with mice
carrying the Cre recombinase gene driven by the muscle creatine kinase (MCK) promoter
(18;31). The resultant M-PKCδKO mice exhibited remarkable decreases of PKCδ
mRNA levels in TA (74%), quadriceps (82.6%), gastrocnemius (79%), and extensor
digitorum longus (EDL) (65%) muscles (all P < 0.05) (Figure 2A). Likewise, a 70%
reduction of PKCδ protein levels was observed in TA muscle fibers purified from M-
PKCδKO mice (Figure 2B). At 10-weeks of age, M-PKCδKO mice did not show
differences in body weight or body composition (i.e., fat and lean mass) as assessed by
Page 10 of 49Diabetes
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DEXA when compared to control littermates (Figure S1A). Furthermore, at sacrifice, no
differences in body weight or muscle weights were observed (data not shown). Muscle
fiber size (data not shown) and gene expression of myosin - proteins which define fiber
type - remained unchanged (Figure S1B). Intraperitoneal glucose tolerance tests (GTT)
performed at 10-weeks of age revealed no difference between the M-PKCδKO and
control mice with the exception of a small, but significant, decrease in glucose levels at
60 minutes in M-PKCδKO mice (Figure 2C and Figure S1C). Likewise, intraperitoneal
insulin tolerance tests (ITT) demonstrated no difference in the insulin-mediated reduction
of glucose levels between knockout and control mice (Figure S1D), and plasma insulin
levels in both the fasted and fed states were unchanged (Figure S1E).
However, insulin signaling, especially at low insulin doses, was improved in the
M-PKCδKO mice. Thus, following injection of insulin (20 U/kg body weight) into the
vena cava of 10-week old mice, phosphorylation of IR in TA muscle was increased by
2.1-fold in M-PKCδKO mice compared to control (Figure 2D and E). Likewise, insulin
stimulated phosphorylation of AKT and ERK was increased by 1.5-fold and 1.4-fold in
M-PKCδKO mice when compared to controls. Interestingly, when insulin was injected at
a higher dose (200 U/kg of body weight), no difference was observed between the control
and M-PKCδKO mice, suggesting saturation of insulin receptor signaling (Figure S1F
and G) at the high dose.
High-fat diet-induced glucose intolerance and muscle insulin resistance are not
prevented by PKCδ deletion in muscle. Previous studies have shown that PKCδ is
increased in the liver of HFD-induced obese mice and that PKCδ plays an important role
in liver in HFD-induced insulin resistance (18). To assess the role of PKCδ in muscle on
Page 11 of 49 Diabetes
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HFD-induced insulin resistance, we placed 5-week old M-PKCδKO and control mice on
either a HFD (60% fat by calories) or chow diet (23% fat by calories) and followed them
for 10 weeks. During this time, control mice on chow diet gained 52% above their
starting body weight, and those on HFD gained almost twice as much (Figure 3A). Body
composition as assessed by DEXA at the end of the diet period revealed that the M-
PKCδKO mice on CD had a tendency to have a greater lean mass than controls (Figure
S2A), but this did not quite reach statistical significance. After 10 weeks of HFD, both
control and M-PKCδKO mice had a similar 2-fold increase in fat mass (Figure S2B).
To determine whether this difference in body weight and composition was
associated with a change in energy balance, we monitored the metabolic activities of the
M-PKCδKO and control mice using metabolic cages. M-PKCδKO mice displayed
similar food intake as control mice (Figure S2C), but showed a trend toward increased
spontaneous activity during the light phase (Figure S2D). The oxygen consumption rate
and respiratory exchange ratio (RER) were not different between M-PKCδKO and
control mice (Figure S2E and F). Despite the higher spontaneous activity, M-PKCδKO
mice showed no difference in the expression of genes that mark muscle fiber type
(myosin IIa, myosin IIx and myosin IIb) or differentiation (myoD1 and myf5) on both
CD and HFD (Figure S2G).
Fasting plasma glucose levels were unchanged between control and M-PKCδKO
mice on either CD or HFD (Figure S3A). Control mice fed a HFD for 10 weeks showed a
2-fold increase in plasma insulin levels, and insulin levels were further increased by
~40% in M-PKCδKO mice on both chow and HFD diet compared to control (Figure 3B).
Both control and M-PKCδKO mice showed impaired glucose tolerance on HFD (Figure
Page 12 of 49Diabetes
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S3B) and higher levels of glucose throughout the insulin tolerance test (Figure S3C) with
no differences between control and M-PKCδKO mice. As expected, HFD challenge in
control mice resulted in reduced muscle insulin signaling with decreased levels of IRS1
tyrosine phosphorylation and AKT serine phosphorylation (Figure 3C), even when
stimulated with an insulin at a dose of 200 U/kg of body weight. The reduction in insulin
signaling, however, was not affected by PKCδ deletion in muscle. Consistent with the
data shown in Figure 1, western blot analysis revealed that PKCδ expression in muscle
was reduced by HFD treatment by 50% in control mice. Since PKCδ activation is
associated with translocation to cell membranes (32), we determined the amount of PKCδ
protein associated with the membrane fraction in muscle of control and M-PKCδKO
mice on both diets. The ratio of membrane to cytosolic fraction of PKCδ was increased
by 54% on HFD in control mice, consistent with the increased activation of PKCδ by
HFD. Compared with control, M-PKCδKO mice demonstrated a 56% decrease in the
ratio of membrane to cytosolic fraction of PKCδ on CD, and a 70% decrease in the ratio
on HFD (Figure 3D). No differences were found in the levels of the reactive oxygen
species (ROS) and reactive nitrogen species (RNS) between M-PKCδKO and control
mice on CD or HFD (Figure S3D).
Insulin sensitivity and glucose tolerance are improved in M-PKCδKO mice at 6- to
7-months of age. This increase was associated with a trend towards increased uptake of
14C-deoxyglucose in the skeletal muscles (TA, EDL, soleus and quad) of the M-PKCδKO
mice (Figure 4B). When insulin-stimulated glucose uptake was assessed ex vivo in soleus
and EDL muscle strips, the M-PKCδKO mice also displayed a trend to increased glucose
uptake, with 44-47% increases of insulin-stimulated 3H-2-deoxy-glucose uptake into
Page 13 of 49 Diabetes
14
isolated EDL and soleus muscles (Figure S4A and Figure 4C). This was associated with
significant 21-25% reductions in glucose levels at 60 and 90 minutes during
intraperitoneal glucose tolerance testing (GTT) in 7-month old M-PKCδKO mice (Figure
4D and S4B). Insulin secretion at 0, 15 and 30 minutes during GTT was unchanged in M-
PKCδKO mice (Figure 4E), and intraperitoneal insulin tolerance tests (ITT) demonstrated
no difference in the insulin-mediated reduction of glucose levels between M-PKCδKO
and control mice (Figure S4C). Assessment of maximal exercise capacity and forelimb
grip strength revealed no differences between age-matched control and M-PKCδKO mice
(Figure S5A and B).
Glucose intolerance and insulin resistance are alleviated by muscle-specific PKCδ
deletion at 15-months of age. At 15-months of age, muscle-specific PKCδ deletion led
to a marked improvement of glucose tolerance with a 28% decrease in AUC (P < 0.05)
(Figure 5A). As in the younger animals, this improved glucose tolerance was not
associated with changes in serum insulin levels in the fasted or fed state (Figure S7A). To
determine whether the improvement of glucose tolerance was related to improved muscle
insulin sensitivity in old M-PKCδKO mice, we measured insulin signaling in the TA
muscle from older control and M-PKCδKO mice at 15-months of age. At this age,
following high dose (200 U/kg body weight) intravenous insulin injection,
phosphorylation of AKT on serine 473 was increased by 3.5-fold, with parallel changes
in ERK phosphorylation in M-PKCδKO muscle when compared to controls (Figure 5B
and C).
To determine whether deletion of PKCδ in muscle could prevent the age-related
decline in insulin signaling, we compared insulin-medicated AKT and ERK
Page 14 of 49Diabetes
15
phosphorylation in muscle from control and M-PKCδKO mice at different ages. When
compared to 10-week old mice, control mice at 15-months of age exhibited a 68%
reduction in insulin-stimulated AKT serine 473 phosphorylation, which was prevented in
M-PKCδKO mice (Figure 5D and E). This correlated with increased protein levels of
PKCδ in muscle as the mice aged (Figure S6). Thus, when compared to control mice, M-
PKCδKO mice exhibited age-dependent decreases in PKCδ protein levels, decreasing by
30% and 75% at 4- and 15-months of age respectively. The residual protein levels PKCδ
may represent an incomplete rearrangement of the floxed alleles in a small subset of
myofibers or, more likely, the presence of other cells types such as satellite cells,
adipocytes, endothelial cells and fibroblasts (23). Indeed, isolated endothelial cells
exhibited 15-fold higher levels of PKCδ mRNA expression compared to skeletal muscle
(Figure S6B and C) when normalized to GAPDH. This high level of PKCδ expression in
endothelial cells likely accounts for the residual expression of PKCδ observed in both the
whole tissue lysates and the isolated muscle fibers.
Impact of muscle-specific PKCδ deletion on body composition and energy
expenditure at 15-months of age. Metabolic assessment was performed in M-PKCδKO
and control mice at 15-months of age. While the total body and muscle weight did not
change (data not shown), DEXA analysis at this age revealed that M-PKCδKO mice had
10% less total fat and 9% less visceral fat when compared to control mice (Figure 6A and
B). Furthermore, M-PKCδKO mice had a 16.5% increase in the ratio of lean mass to fat
mass when compared to control mice (Figure 6C). Gene expression of muscle fiber type
markers and markers of differentiation indicated that there was no change in the
characteristics of muscle from older M-PKCδKO mice (Figure S7B). CLAMS metabolic
Page 15 of 49 Diabetes
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cage analysis revealed that while M-PKCδKO mice showed similar food intake and
activity as control mice (Figure S7C and D), both oxygen consumption and CO2
production normalized to lean body mass in older M-PKCδKO mice were reduced
significantly by 10% and 11% respectively during the light cycle. A similar trend was
also observed during the dark cycle (Figure 6D and E). RER was not different between
the two genotypes (Figure S7E).
To investigate the mechanism by which muscle-specific PKCδ deletion leads to
decreased oxygen consumption, we isolated mitochondria from 15-month old M-
PKCδKO and control mice, and measured their metabolic capacity using different
substrates in the absence (basal) or presence of ADP (state 3). In the presence of
pyruvate/malate (PDH and complex I substrate), the addition of ADP increased the
oxygen consumption rate (OCR) by 1.6-fold over basal in mitochondria isolated from
older control mice, and mitochondria of age-matched M-PKCδKO mice displayed a
similar change (Figure S7F). Likewise, in the presence of palmitoyl-carnitine/malate (β-
oxidation substrate), both control and M-PKCδKO mice displayed a 2.2-fold increase in
the OCR in state 3 when compared to basal state (Figure S7G). The respiratory control
ratio (RCR: state 3/state 4), an index of mitochondrial coupling, was also similar between
control and M-PKCδKO mice in the presence of either pyruvate/malate or palmitoyl-
carnitine/malate (Figure S7H). Protein levels of the voltage dependent anion channel
(VDAC) were decreased by 80% and levels of complex I subunit also trended to be
decreased in 15-month old M-PKCδKO mice when compared to control littermates
(Figure S8A and B). Compared to 10-week old mice, the ROS and RNS levels in muscle
were significantly increased by 19% in 15-month old control mice (Figure 6F). Together
Page 16 of 49Diabetes
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these data suggest a role for PKCδ in the regulation of mitochondrial homeostasis at older
age.
Page 17 of 49 Diabetes
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Discussion
We have previously demonstrated an important role of PKCδ in the regulation of
hepatic insulin sensitivity and hepatosteatosis in both mice and humans (18). Indeed,
PKCδ levels in liver are higher in strains of mice that are susceptible to diet induced
obesity and insulin resistance. Overexpression of PKCδ in liver of mice increases insulin
resistance, and liver-specific knockout of PKCδ in mice challenged with HFD improves
insulin sensitivity and glucose tolerance. On the other hand, previous studies with
myoblasts and myotubes in vitro indicate that PKCδ may play a different role in muscle
serving as a positive regulator of insulin signaling (5;13;33). Moreover, overexpression
of PKCδ in primary muscle cells has been shown to increase GLUT4 translocation and
glucose uptake in the basal state, and pharmacological inhibition of PKCδ caused down-
regulation of insulin-induced GLUT4 translocation and glucose uptake (5). In cultured
muscle cells, insulin stimulation has also been shown to lead to a direct association of
PKCδ with the insulin receptor, with enhanced insulin-induced tyrosine phosphorylation
and insulin receptor internalization (33). In Psammomys obesus, exercise also increases
PKCδ levels in muscle and insulin receptor phosphorylation (34;35).
In the present study, we investigated the role of PKCδ in skeletal muscle insulin
action and whole body insulin sensitivity by creating a muscle-specific PKCδ knockout
(M-PKCδKO) mouse. We found that at a young age, muscle insulin signaling was
improved in M-PKCδKO mice when stimulated with a low dose of insulin of 20 U/kg
body weight, but was not changed at a maximal doses (200 U/kg), consistent with
improved insulin sensitivity with no change in maximal response. Despite the increased
signaling in M-PKCδKO muscle at lower dose, decreased PKCδ level in muscle had no
Page 18 of 49Diabetes
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effect on whole body glucose tolerance in mice placed on either CD or HFD. However, at
6-months of age M-PKCδKO mice demonstrated an improvement in whole body insulin
sensitivity as measured during a hyperinsulinemic-euglycemic clamp. At 15-months of
age, muscle-specific deletion of PKCδ in mice improved muscle insulin signaling and
resulted in a significant improvement in glucose tolerance. The improvement in insulin
sensitivity and glucose tolerance was also accompanied by a shift towards a “leaner”
body composition in the older M-PKCδKO mice with less fat, especially visceral fat, and
a higher ratio of lean to fat mass. Since the accumulation of visceral fat is an important
contributor to the development of diabetes and insulin resistance (19;36), it is possible
that the decreased visceral fat observed in old M-PKCδKO mice plays a role in the
improved insulin sensitivity and glucose tolerance. These data also suggest a crosstalk
between muscle and fat, similar to what was previously observed when the insulin
receptor was deleted from muscle (MIRKO mouse) but in the opposite direction, i.e., in
MIRKO mice fat mass is increased (37). One reason for the different responses observed
in M-PKCδKO mice at different ages is the increase in PKCδ expression in muscle as the
mice progressed in age. Indeed, PKCδ protein levels in muscle increase by 70% as the
mice mature from 10-weeks to 15-months of age. Many previous in vitro studies have
shown that upon exposure to reactive oxygen species (ROS), PKCδ is up-regulated and
activated to initiate important cellular responses (38;39). Indeed, at 15-months of age
mice demonstrated a 19% increase of ROS/RNS levels in skeletal muscle when compared
to younger cohorts, suggesting the accumulation of ROS during the aging process may
contribute to the increased levels of PKCδ.
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Previous studies have shown that PKCδ plays important roles in the regulation of
mitochondrial mass and mitochondrial metabolism (40-42). Indeed, markers of
mitochondrial mass, such as VDAC protein and mitochondrial complex I protein, were
decreased in 15-month old M-PKCδKO mice. M-PKCδKO mice also displayed a 10%
reduction in oxygen consumption at the whole body level when compared to controls. To
see if loss of PKCδ impairs mitochondrial function, we determined oxygen consumption
rates in isolated mitochondria from M-PKCδKO and control mice at 15-months of age.
Surprisingly we observed no differences in basal or ADP-dependent mitochondrial
oxidative capacity, when using two different substrates and normalized to mitochondrial
protein. These data suggest PKCδ may play an important role in the maintenance of
mitochondrial mass in skeletal muscle, but does not have as great an impact on
mitochondrial function.
We previously showed that liver-specific reduction of PKCδ in HFD-treated
obese mice improved glucose tolerance and insulin sensitivity by decreasing IRS-1 serine
phosphorylation and increasing Akt phosphorylation (18). We hypothesized that similar
mechanisms might contribute to HFD-induced insulin resistance in muscle and that
muscle-specific PKCδ deletion may rescue HFD-induced insulin resistance. Indeed,
several previous studies have shown that diacylglycerol (DAG) accumulation in muscle
can activate novel PKCs, such as PKCθ, and impair muscle insulin action (43). However,
our results show that, in contrast to liver-specific PKCδ deletion, PKCδ deletion in
muscle is not sufficient to rescue the HFD-induced glucose intolerance and muscle
insulin resistance. These opposite observations may be explained by the opposing
expression patterns of PKCδ in the two tissues when exposed to HFD. Hepatic PKCδ
Page 20 of 49Diabetes
21
expression was increased by more than 50% in the HFD-treated B6 mice when compared
to CD group. However, muscle PKCδ expression was decreased by 50% in mice placed
on HFD when compared to those on CD, indicating differences in PKCδ regulation in
these tissues. While the mechanism of the HFD-induced differences in PKCδ expression
in liver and muscle are unclear, activation of other PKC isoforms in muscle, such as
PKCθ and ε, may contribute to the impairment of insulin signaling in muscle and
overcome the protective effect of PKCδ deletion in the muscle of diet-induced obese
mice (44-46).
In summary, the role of PKCδ in muscle is different depending on the etiology of
insulin resistance. While the role of PKCδ in elderly animals remains to be determined,
PKCδ levels are increased in skeletal muscle of middle-aged mice, and muscle-specific
deletion of PKCδ improved the middle-age-related decline in glucose tolerance and
whole body insulin resistance. It also improved the ratio of lean to fat mass. In HFD-
induced obesity, PKCδ levels were decreased in skeletal muscle, and deletion of PKCδ
did not rescue HFD-induced insulin resistance and glucose intolerance. The distinct
pattern of regulation in muscle and liver sheds new light on how PKCδ functions in
different insulin-responsive tissues and the therapeutic potential of PKCδ as a target for
the tissue-specific improvement in insulin resistance.
Page 21 of 49 Diabetes
22
Figure Legends
Figure 1. Effect of aging and HFD on PKCδ muscle expression in wild type B6 mice.
(A) PKCδ protein content in triceps muscle of 10-week, 8-month and 15-month old wild
type B6 mice as measured by western blot analysis (n = 8 per group). (B) PKCδ mRNA
expression and (C) protein content in tibialis anterior (TA) muscle of 3-month old wild-
type B6 mice fed either a CD or HFD for 10 weeks. A representative western blot is
shown in Figure 4E. Data are mean ±SEM. n = 8. * P < 0.05 vs. 10 weeks by one-way
ANOVA; ‡ P < 0.05 vs. CD by Student’s t-test.
Figure 2. Glucose tolerance and muscle insulin signaling are normal in muscle-
specific PKCδ knockout mice at 10-weeks of age. (A) PKCδ mRNA expression in
tibialis anterior (TA), quadriceps (Quad), gastrocnemius (Gastro), extensor digitorum
longus (EDL) and soleus muscles from Lox and M-PKCδKO mice. (B) Representative
western blots and densitometric quantification of PKCδ in the purified TA muscle fibers
of Lox and M-PKCδKO mice with quantification below. (C) Intraperitoneal glucose
tolerance tests were performed in Lox and M-PKCδKO mice. (D) Western blot analysis
of insulin signaling pathway in skeletal muscle of Lox and M-PKCδKO mice after
intravenous insulin injection (20 U/kg of body weight). (E) Densitometric analysis of IR,
IRS-1, AKT and EKR phosphorylation from panel D after insulin stimulation. White bars
represent control mice; black bars represent M-PKCδKO mice. Data are mean ±SEM.
n=6-8 per group in panels (A) – (C), and n = 4 per group for panels (D) and (E). ‡ P <
0.05 by Student’s t-test.
Page 22 of 49Diabetes
23
Figure 3. Muscle-specific knockout of PKCδ does not alleviate HFD-induced muscle
insulin resistance. (A) Body weight changes of Lox and M-PKCδKO mice fed with
either a CD or HFD from 5- to 16- weeks of age (n = 10 per group). (B) Insulin levels
were measured in 4-month old Lox and M-PKCδKO mice fed either a CD or HFD (n = 6-
8 per group). (C) Western blot analysis of insulin signaling in TA muscle of Lox and M-
PKCδKO after intravenous insulin injection (200 U/kg of body weight). (D) Western blot
of PKCδ protein in cytoplasmic (cyto) and membrane (MEM) fractions of gastrocnemius
muscle from Lox and M-PKCδKO mice fed either a CD or a HFD. White bars represent
control mice; black bars represent M-PKCδKO mice. Data are mean ±SEM. * P < 0.05
by two-way ANOVA in figure 3A (CD-treated control vs. HFD-treated control mice) and
3B.
Figure 4. Muscle-specific knockout of PKCδ improves whole body insulin sensitivity
and glucose tolerance at 6- to 7-months of age. (A) Glucose infusion rate and (B)
insulin stimulated 14
C-deoxyglucose uptake were measured during hyperinsulinemic-
euglycemic clamps in 6-month old M-PKCδKO mice (n=3-7). (C) Rates of 3H-2-deoxy-
glucose uptake into soleus muscles in the presence or absence of 2.5 mU/ml of insulin (n
= 4-5). (D) Intraperitoneal glucose tolerance tests (GTT), and (E) Insulin secretion during
GTT at 0, 15 and 30 min (n = 5). White bars represent Lox mice; black bars represent M-
PKCδKO mice. Data are mean ±SEM. ‡ P < 0.05 by Student’s t-test.
Figure 5. Glucose tolerance and muscle insulin signaling are improved in M-
PKCδKO mice at 15-months of age. (A) Intraperitoneal glucose tolerance tests were
performed in 15-month old Lox and M-PKCδKO mice (n=12 per group) and area under
the curve was measured. (B) Insulin signaling was analyzed in TA muscle of 15-month
Page 23 of 49 Diabetes
24
old Lox and M-PKCδKO mice by western blot after intravenous insulin injection (200
U/kg of body weight). (C) Insulin-stimulated phosphorylation of the insulin receptor,
AKT and ERK after insulin stimulation were quantified by densitometric analysis. (D)
Representative western blot of insulin signaling from TA muscles of Lox and M-
PKCδKO mice at 10 week, 4 months and 15-months of age. (E) Densitometric analysis
of insulin-stimulated AKT and ERK phosphorylation at 10-weeks and 15-months of age.
White bars represent Lox mice; black bars represent M-PKCδKO mice. Data are mean
±SEM. ‡ P < 0.05 by Student’s t-test; * P < 0.05 by two-way ANOVA.
Figure 6. Impact of muscle-specific deletion of PKCδ on body composition and
energy expenditure at 15-months of age. (A) Total fat and lean mass, (B) percentage of
visceral fat and subcutaneous fat per body weight and (C) ratio of lean mass to fat mass
were assessed by DEXA in 15-month old Lox and M-PKCδKO mice (n=12 per group).
(D) Oxygen consumption and (E) CO2 production measured in Lox and M-PKCδKO
mice over 48 h in CLAMS metabolic cages (n=12 per group). (F) Reactive oxygen and
reactive nitrogen species were measured in TA muscles of Lox and M-PKCδKO mice at
10 weeks and 15-months of age. White bars represent control mice; black bars represent
M-PKCδKO mice. Data are mean ±SEM. ‡ P < 0.05 by Student’s t-test. * P < 0.05 by
two-way ANOVA.
Page 24 of 49Diabetes
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Figure Supplement Legends
Figure S1. Insulin tolerance, gene expression of myogenesis markers and muscle
fiber type markers and muscle insulin signaling in 10-week old mice. (A) Body
weight, total fat and total lean mass of Lox and M-PKCδKO mice fed a chow diet were
measured by DEXA analysis at 10-weeks of age. (B) Gene expression of muscle markers
for muscle fiber type and myogenesis were assessed by qPCR in TA muscle of 10-week
old Lox and M-PKCδKO mice. (C) Area under the curve of intraperitoneal glucose
tolerance tests and (D) insulin tolerance tests were performed in Lox and M-PKCδKO
mice. (E) Fasted and fed serum insulin levels in Lox and M-PKCδKO mice. (F) Western
blot analysis of insulin signaling pathway in skeletal muscle of Lox and M-PKCδKO
mice after intravenous insulin injection (200 U/kg of body weight). (G) Densitometric
analysis of insulin receptor phosphorylation and AKT phosphorylation from panel F after
insulin stimulation. White bars represent control mice; black bars represent M-PKCδKO
mice. Data are mean ±SEM. n=6-8 per group in panels (A) – (E), and n = 3 per group in
panels (F) and (G).
Figure S2. Body composition and energy expenditure in M-PKCδKO mice on either
a chow diet or high-fat diet. (A) Total lean and (B) total fat mass were assessed by
DEXA in 4-month old Lox and M-PKCδKO mice fed with CD or HFD. (C) Food intake
was measured by CLAMS in a 48 hr cycle after 10 weeks treatment with CD or HFD. (D)
Spontaneous activity was measured in Lox and M-PKCδKO mice during 48 h cycle in
metabolic cages. (E) Oxygen consumption (VO2) and (F) respiratory exchange ratio
(RER) were analyzed by metabolic cage assessment. (G) Gene expression of muscle
markers for muscle fiber type and myogenesis were assessed by qPCR in TA muscle of
Page 25 of 49 Diabetes
26
Lox and M-PKCδKO mice fed either a CD or a HFD (n = 4 per group). n = 5 per group
in panels (A) - (F). White bars represent control mice; black bars represent M-PKCδKO
mice. Data are mean ±SEM.
Figure S3. Muscle-specific knockout of PKCδ does not alleviate HFD-induced
muscle insulin resistance. (A) Glucose levels were measured in 4-month old Lox and
M-PKCδKO mice fed either a CD or HFD. (B) Intraperitoneal glucose tolerance tests and
its area under the curve, and (C) insulin tolerance tests were performed in 4-month old
Lox and M-PKCδKO mice fed either a CD or HFD from 6-weeks of age. (D) Reactive
oxygen and reactive nitrogen species were measured in TA muscles of Lox and M-
PKCδKO mice fed either a CD or HFD. White bars represent control mice; black bars
represent M-PKCδKO mice. Data are mean ±SEM. n= 5 per group in panels (A) to (D).
Figure S4. Ex vivo glucose uptake in EDL and insulin tolerance of M-PKCδKO mice
at 6- to 7-months of age. (A) Rates of 3H-2-deoxy-glucose uptake into muscles in the
presence or absence of 2.5 mU/ml of insulin (n = 4-5). (B) Area under the curve of
intraperitoneal glucose tolerance tests (GTT), and (C) insulin tolerance tests performed in
Lox and M-PKCδKO mice at 7-months of age (n = 5). White bars represent Lox mice;
black bars represent M-PKCδKO mice. Data are mean ±SEM.
Figure S5. Exercise capacity and muscle grip strength of mice at 7-months of age.
(A) Maximal exercise capacity was measured in 7-month old Lox and M- PKCδKO mice
that were exercised on a treadmill until exhaustion. (B) Forelimb grip strength was
determined in Lox and M- PKCδKO mice. Data are mean ±SEM.
Page 26 of 49Diabetes
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Figure S6. PKCδ protein are increased in muscle of 15-month old mice. (A) PKCδ
protein content in TA muscle of 10-week, 4- and 15-month old Lox and M-PKCδKO
mice by western blot analysis. PKCδ (B) and CD31 (C) mRNA expression in
gastrocnemius (Gastro), tibialis anterior (TA) and liver endothelial cells (Endothelial)
from control mice. Data are mean ±SEM. * P < 0.05 by two-way ANOVA.
Figure S7. Plasma insulin and energy expenditure of mice at 15-months of age. (A)
Fast and fed insulin levels were measured in 15-month old control and M-PKCδKO mice
(n=6 per group). (B) mRNA levels of muscle fiber type markers were measured in TA
muscle of 15-month old Lox and M-PKCδKO mice by qPCR (n = 5 per group). (C) Food
intake in 15-month old Lox and M-PKCδKO mice (n=12 per group). (D) Spontaneous
activity was measured in Lox and M-PKCδKO mice over 48 h in CLAMS metabolic
cages (n=12 per group). (E) respiratory exchange ratio (RER) from the same analysis. (F)
Basal and ADP-dependent (State 3) respiratory rates of isolated mitochondria from
hindlimb muscle were measured using a Seahorse X24 Flux Analyzer in the presence of
pyruvate/malate (n=4 per group) as described in Materials and Methods. (G) Basal and
State 3 respiratory rates of isolated mitochondria from hindlimb muscle were measured
with palmitoyl-carnitine/malate (fatty acid) as substrate (n=4 per group). (H) Respiratory
control ratio (RCR) of isolated mitochondria was measured with pyruvate/malate (PM) or
palmitoyl-carnitine/malate (PC) as substrate (n=4 per group). OCR, oxygen consumption
rate. White bars represent control mice; black bars represent M-PKCδKO mice. Data are
mean ±SEM.
Figure S8. Impact of muscle-specific PKCδ deletion on mitochondrial protein
content at 10-week, 4-months and 15-months of age. (A) Protein levels of
Page 27 of 49 Diabetes
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mitochondrial marker (VDAC) and muscle oxidative phosphorylation complex subunits
(mitochondrial complex I, II and IV) were assessed by western blot in TA muscles of Lox
and M- PKCδKO mice at 10-weeks, 4-months and 15-months of age (n = 3 per group).
(B) Densitometric quantification of western blots. White bars represent Lox mice; black
bars represent M-PKCδKO mice. Data are mean ±SEM. * P < 0.05 by two-way
ANOVA;
Acknowledgments
We thank Graham Smyth for assistance with the animal care, Christian Doherty
and Mike Hirshman from Laurie J. Goodyear’s lab for assistance with the ex vivo 2-DG
glucose uptake assay, Hans P.M.M. Lauritzen for assistance with the muscle fiber
purification, and Erica P. Homan for assistance with the proof reading. This work was
supported by National Institutes of Health grants R01DK033201, and the Joslin DRC
grant DK036836. B.T.O. was also funded by a K08 Training award from the NIDDK of
the NIH (K08DK100543).
M.L. generated the data and wrote the manuscript. O.B. initiated the project,
reviewed and edited the manuscript. B.T.O. and S.G.V. reviewed the manuscript and
generated the data. C.R.K. oversaw the project, contributed to discussion, and helped
write the manuscript. C.R.K. is the guarantor of this work and, as such, had full access to
all the data in the study and takes responsibility for the integrity of the data and the
accuracy of the data analysis. There are no potential conflicts of interest relevant to this
article by any of the authors.
Page 28 of 49Diabetes
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Page 35 of 49 Diabetes
1 0 W 8 M 1 5 M
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Page 36 of 49Diabetes
Fig. 2
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Akt
P-ERK1/2
ERK1/2
0
5 0
1 0 0
1 5 0
2 0 0
2 5 0
P-I
R/I
R (
% o
f L
ox
aft
er i
ns
uli
n s
tim
ula
tio
n)
*
0
5 0
1 0 0
1 5 0
2 0 0
P-I
RS
1/I
RS
1 (
% o
f L
ox
aft
er i
ns
uli
n s
tim
ula
tio
n)
0
1 0 0
2 0 0
3 0 0
4 0 0
P-A
KT
/AK
T (
% o
f L
ox
aft
er i
ns
uli
n s
tim
ula
tio
n)
P = 0 .1
0
1 0 0
2 0 0
3 0 0
P-E
RK
/ER
K (
% o
f L
ox
aft
er i
ns
uli
n s
tim
ula
tio
n)
*
GAPDH
GAPDH
Page 37 of 49 Diabetes
C D H F D
0
1
2
3
4
5
Pla
sm
a i
ns
uli
n
(ng
/ml)
*
**
A. B.
Fig. 3
C.
D.
Lox M-PKCδKO
Cyto MEM Cyto MEM Cyto MEM Cyto MEM
Lox M-PKCδKO
CD HFD
PKCδ
PKCδ(lighter exposure)
P-IR (T1150/1151)
Insulin
P-Akt (S473)
Akt
P-ERK1/2
P-IRS1 (Y612)
IRS1
ERK1/2
IR
Lox
+ - - + + - - + + - - + + - - +
Lox -
CD HFD
GAPDH
PKCδ
M-PKCδKO M-PKCδKO
5 6 7 8 9 1 0 1 1 1 2 1 3 1 4
1 0
2 0
3 0
4 0
5 0
C D -fl/fl
C D -M -P K C K O
H F D -fl/fl
H F D -M -P K C K O
A g e (w k s )
Bo
dy
We
igh
t (g
)
**
**
Page 38 of 49Diabetes
Fig. 4
0 3 0 6 0 9 0 1 2 0
0
1 0 0
2 0 0
3 0 0
4 0 0
5 0 0
L o x
M -P K C K O
T im e (m in )
Glu
co
se
(m
g/d
l) ‡‡
0 1 0 2 0 3 0 4 0
0 .0
0 .5
1 .0
1 .5
T Im e (m in )
Ins
uli
n (
ng
/ml)
du
rin
g G
TT
L o x
M -P K C K O
D.
E.
C.
A.
Lo
x
M-P
KCK
O
0
5
1 0
1 5
2 0G
luc
so
e i
nfu
sio
n r
ate
(mg
/kg
/min
) ‡
TA
ED
L
So
leu
s
Qu
ad
0
2 0 0
4 0 0
6 0 0
Glu
co
se
up
tak
e
(nm
ol/
g/m
in)
B.
B a s a l In s u l in
0
2
4
6
8
Glu
co
se
Up
tak
e
(um
ol/
ml/
h)
Page 39 of 49 Diabetes
Fig. 5
D.
1 0 W 1 5 M
0
5 0
1 0 0
1 5 0
P-A
KT
/AK
T (
% o
f L
ox
)
aft
er i
ns
uli
n s
tim
ula
tio
n)
**
1 0 W 1 5 M
0
1 0 0
2 0 0
3 0 0
4 0 0
5 0 0
P-E
RK
/ER
K (
% o
f L
ox
)
aft
er i
ns
uli
n s
tim
ula
tio
n)
*
C.
0 3 0 6 0 9 0 1 2 0
0
1 0 0
2 0 0
3 0 0
4 0 0
5 0 0
L o x
M P K C K O
T im e (m in )
Glu
co
se
(m
g/d
l)
‡ ‡‡
A.
Lo
x
MP
KCK
O
0
2 0 0
4 0 0
6 0 0
8 0 0
*
Are
a u
nd
er t
he
cu
rv
e
(mg
/dl/
hr)
P-IR (T1150/1151)
P-Akt (S473)
Akt
P-ERK1/2
ERK1/2
IR
Insulin
Lox
+-
M-PKCδKO
- +-
GAPDH
+ +- +- - +- + +
B.
Lo
x
M-P
KCK
O
0
1 0 0
2 0 0
3 0 0
P-I
R/I
R (
% o
f L
ox
)
(aft
er i
ns
uli
n s
tim
ula
tio
n)
P = 0 .0 8
Lo
x
M-P
KCK
O
0
1 0 0
2 0 0
3 0 0
4 0 0
5 0 0
P-A
KT
/AK
T (
% o
f L
ox
)
(aft
er i
ns
uli
n s
tim
ula
tio
n)
‡
Lo
x
M-P
KCK
O
0
1 0 0
2 0 0
3 0 0
4 0 0
P-E
RK
/ER
K (
% o
f L
ox
)
(a
fte
r i
ns
uli
n s
tim
ula
tio
n)
‡
Insulin +- - - - - - - - - - - -+ + + + + + + + + + + +
Lox Lox LoxM-PKCδKO M-PKCδKO M-PKCδKO
10 W 4 M 15 M
P-Akt (S473)
P-ERK
ERK
Akt
GAPDH
E.
Page 40 of 49Diabetes
Fig. 6
A. B.
D. E.
F.
To
tal F
at
To
tal L
ean
0
1 0
2 0
3 0
4 0
Ma
ss
(g
ra
ms
)
‡
L ig h t D a rk
0
1 0 0 0
2 0 0 0
3 0 0 0
4 0 0 0
5 0 0 0
6 0 0 0
VO
2
(ml/
kg
of
lea
n m
as
s/h
r)
‡
V S F a t S Q F a t
0
2
4
6
8
1 0
1 2
1 4
Ma
ss
(% o
f b
od
y w
eig
ht) ‡ C.
Lo
x
M-P
KCK
O
0 .0
0 .5
1 .0
1 .5
Ra
tio
of
lea
n t
o f
at
ma
ss
‡
L ig h t D a rk
0
1 0 0 0
2 0 0 0
3 0 0 0
4 0 0 0
5 0 0 0
VC
O2
(ml/
kg
of
lea
n m
as
s/h
r)
‡
1 0 W 1 5 M
0
1 0 0 0
2 0 0 0
3 0 0 0 *
RO
S &
RN
S l
ev
els
(Arb
itra
ry
un
its
)
Page 41 of 49 Diabetes
Suppl. 1
A. B.
Myo
sin
2a
myo
sin
2x
Myo
sin
2b
Myo
D1
Myf5
0 .0
0 .2
0 .4
0 .6
0 .8
1 .0
1 .2
1 .4
1 .6
mR
NA
ex
pre
ss
ion
(fo
ld C
ha
ng
e)
C.
BW
To
tal F
at
To
tal L
ean
0
5
1 0
1 5
2 0
2 5M
as
s (
gra
ms
)
Lo
x
M-P
KCK
O
0
2 0 0
4 0 0
6 0 0
8 0 0
Are
a u
nd
er t
he
cu
rv
e
(mg
/dl/
hr)
0 3 0 6 0 9 0 1 2 0 1 5 0
0
5 0
1 0 0
1 5 0
2 0 0
2 5 0
L o x
M -P K C K O
T im e (m in )
Glu
co
se
(m
g/d
l)
D.
F a s t F e d
0 .0
0 .5
1 .0
1 .5
Pla
sm
a i
ns
uli
n (
ng
/ml)
E. F.
P-IR (Y1150/1151)
Lox M - PKCδ KO
- + + - - + + Insulin
P - Akt (S473)
Akt
P - ERK1/2
P-IRS1 (Y612)
GAPDH
ERK1/2
- +
IR
- + - -
PKCδ
IRS1
Lo
x
M-P
KCK
O
0
5 0
1 0 0
1 5 0
P-I
R/I
R (
% o
f L
ox
)
(aft
er i
ns
uli
n s
tim
ula
tio
n)
Lo
x
M-P
KCK
O
0
5 0
1 0 0
1 5 0
P-A
KT
/AK
T (
% o
f L
ox
)
(aft
er i
ns
uli
n s
tim
ula
tio
n)G.
Page 42 of 49Diabetes
A. B.
C D H F D
0
5
1 0
1 5
2 0
2 5
To
tal
lea
n m
as
s (
g)
F. G.
CD HFD0
5
10
15
20
To
tal
fat
ma
ss
(g
)
C.
Lig
ht
Dark
Lig
ht
Dark
0
5 0
1 0 0
1 5 0
2 0 0
2 5 0
C D H F D
Ac
tiv
ity
(X
-am
b C
ou
nts
)
Lig
ht
Dark
Lig
ht
Dark
0
2 0 0 0
4 0 0 0
6 0 0 0
8 0 0 0
C D H F D
VO
2
(ml/
kg
of
lea
n m
as
s/h
r)
Lig
ht
Dark
Lig
ht
Dark
0 .0
0 .2
0 .4
0 .6
0 .8
1 .0
C D H F D
RE
R
D. E.
C D H F D
0
2 0
4 0
6 0
Fo
od
in
tak
e (
KJ
/48
hrs
)
Suppl. 2
Myo
sin 2
a
myo
sin 2
x
Myo
sin 2
b
Myo
D1
Myf
50.0
0.5
1.0
1.5
2.0
2.5CD-fl/flCD-M-PKCKOHFD-fl/flHFD-M-PKCKO
mR
NA
ex
pre
ss
ion
(fo
ld C
ha
ng
e)
Page 43 of 49 Diabetes
C. D.
0 30 60 90 1200
200
400
600
800 CD-fl/fl
CD-M-PKCKO
HFD-fl/fl
HFD-M-PKCKO
Time (min)
Glu
co
se
(m
g/d
l)
C D H F D
0
5 0
1 0 0
1 5 0
2 0 0
Pla
sm
a g
luc
os
e
(mg
/dl)
A.
B.
0 30 60 90 1200
100
200
300CD-fl/fl
CD-M-PKCKO
HFD-fl/fl
HFD-M-PKCKO
Time (min)
Glu
co
se
(m
g/d
l)
C D H F D
0
2 0 0
4 0 0
6 0 0
8 0 0
1 0 0 0
Are
a u
nd
er t
he
cu
rv
e
(mg
/dl/
hr)
C D H F D
0
1 0 0 0
2 0 0 0
3 0 0 0
4 0 0 0
RO
S &
RN
S l
ev
els
(Arb
itra
ry
un
its
)
Suppl. 3 Page 44 of 49Diabetes
C.
A. B.
B a s a l In s u l in
0
1
2
3
4
5
Glu
co
se
Up
tak
e
(um
ol/
ml/
h)
Lo
x
M-P
KCK
O
0
2 0 0
4 0 0
6 0 0
8 0 0
1 0 0 0
Are
a u
nd
er t
he
cu
rv
e
(mg
/dl/
hr)
0 3 0 6 0 9 0 1 2 0
0
1 0 0
2 0 0
3 0 0
M -P K C d K O
L o x
T im e (m in )
Glu
co
se
(m
g/d
l)
Suppl. 4 Page 45 of 49 Diabetes
Lox M-PKCKO0
50
100
150T
ota
l W
ork
(J)
Lox M-PKCKO0.0
0.5
1.0
1.5
Gri
p s
tre
nth
(N
ew
ton
)
Suppl. 5
A.
B.
Page 46 of 49Diabetes
fl KO fl fl fl fl fl fl fl fl KO KO KO KO KO KO KO KO
10 W 4 M 15 M
PKCδ
GAPDH
1 0 W 4 M 1 5 M
0
5 0
1 0 0
1 5 0
PK
C
pro
tein
co
nte
nt
(Arb
itra
ry
Un
its
) ***
*
L o x
M -P K C K O
Suppl. 6
A.
B.
Gastr
oT
A
En
do
thelial
0
5
1 0
1 5
2 0
Re
lati
ve
mR
NA
ex
pre
ss
ion
of
PK
C
Gastr
oT
A
En
do
thelial
0
1 0
2 0
3 0
4 0
Re
lati
ve
mR
NA
ex
pre
ss
ion
of
CD
31
C.
Page 47 of 49 Diabetes
C. D.
L ig h t D a rk
0 .0
0 .2
0 .4
0 .6
0 .8
1 .0
RE
R
L ig h t D a rk
0
3 0
6 0
9 0
1 2 0
1 5 0
Ac
tiv
ity
(X
-am
b C
ou
nts
)
E.
B a s a l S ta te 3
0
1 0 0
2 0 0
3 0 0
4 0 0
5 0 0
py
ru
va
te/m
ala
te O
CR
(pM
ole
s/m
in/1
0
g)F.
B a s a l S ta te 3
0
1 0 0
2 0 0
3 0 0
4 0 0
pa
lmit
oy
l-c
arn
itin
e/m
ala
te
OC
R (
pM
ole
s/m
in/1
0
g)
G. H.
P M P C
0
1
2
3
4
RC
R (
Sta
te3
/Sta
te4
)
Lo
x
MP
KCK
O
0
1 0
2 0
3 0
4 0
Fo
od
in
tak
e
(KJ
/48
hr)
B.
Fast Fed0
2
4
6
8
10
Pla
sm
a i
ns
uli
n(n
g/m
l)
A.
Suppl. 7
Myo
sin 2
a
myo
sin 2
x
Myo
D1
0.0
0.5
1.0
1.5
2.0 Lox
M-PKCKO
mR
NA
ex
pre
ssio
n
(fo
ld C
ha
ng
e)
Page 48 of 49Diabetes
Suppl. 8
fl KO fl fl fl fl fl fl fl fl KO KO KO KO KO KO KO KO
CI- NDUFB8
CIV- mtCo1 CII-SDHB
10 W 4 M 15 M
GAPDH
VDAC
1 0 W 4 M 1 5 M
0
1 0 0
2 0 0
3 0 0
4 0 0
VD
AC
pro
tein
co
nte
nt
(Arb
itra
ry
Un
its
) *
1 0 W 4 M 1 5 M
0
5 0
1 0 0
1 5 0
Co
mp
lex
II
an
d I
V
pro
tein
co
nte
nt
(Arb
itra
ry
Un
its
)
1 0 W 4 M 1 5 M
0
5 0
1 0 0
1 5 0
Co
mp
lex
I p
ro
tein
co
nte
nt
(Arb
itra
ry
Un
its
)
A.
B.
Page 49 of 49 Diabetes
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