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The mitochondrial energy – redox axis in aging and caloric restriction: role of nicotinamide nucleotide transhydrogenase
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Content
THE MITOCHONDRIAL ENERGY – REDOX AXIS
IN AGING AND CALORIC RESTRICTION:
ROLE OF NICOTINAMIDE NUCLEOTIDE TRANSHYDROGENASE
by
Fei Yin
A Dissertation Presented to the
FACULTY OF THE USC GRADUATE SCHOOL
UNIVERSITY OF SOUTHERN CALIFORNIA
In Partial Fulfillment of the
Requirements for the Degree
DOCTOR OF PHILOSOPHY
(PHARMACEUTICAL SCIENCES)
May 2012
Copyright 2012 Fei Yin
ii
DEDICATION
To my parents for their love and support throughout my life
To my wife for her understanding and encouragement
To our sweet daughter whose birth has brought wonderful fun, great motivation,
and bright inspiration into our life
iii
ACKNOWLEDGEMENTS
First and foremost I offer my sincerest gratitude to my mentor, Professor Enrique
Cadenas. I am so grateful for his unconditional support, patient guidance and invaluable
suggestions throughout my study and writing of this thesis. In addition to the
extraordinary laboratory environment that he provided me, he also granted me the
freedom to develop my own idea, and set the highest standards of science and ethics for
me to reach for as a graduate student.
My sincere thanks go to my Ph.D. dissertation committee members, Dr. Ronald
Alkana, and Dr. Kelvin Davies, for their encouragement and insightful comments. I also
would like to thank my guidance committee members, Dr. Wei-Chiang Shen, Dr. Curtis
Okamoto and Dr. Michel Baudry for their help on the initiation and progression of my
study.
My warmest thanks are due to all of my current and former laboratory colleagues
including, Dr. Allen Chang, Dr. Jerome Garcia, Dr. Ryan Hamilton, Dr. Derick Han, Dr.
Juliana Hwang-Levin, Dr. Philip Lam, Dr. Li-Peng Yap, Dr. Qiongqiong Zhou, Amit
Agarwal, Joanne Lee, Chen Li, Tianyi Jiang and Harsh Sancheti. I also wish to extend
my thanks to Dr. Jia Yao and Dr. Gennady Ermak for their help and support to my study.
iv
TABLE OF CONTENTS
DEDICATION .................................................................................................................... ii
ACKNOWLEDGEMENTS ............................................................................................... iii
TABLE OF CONTENTS ................................................................................................... iv
LIST OF FIGURES ........................................................................................................... vi
ABBREVIATIONS ......................................................................................................... viii
ABSTRACT .........................................................................................................................x
CHAPTER 1 .......................................................................................................................1
OVERVIEW OF THE MITOCHONDRIAL ENERGY
METABOLISM AND REDOX SIGNALING IN BRAIN AGING
AND NEURODEGENERATION
1.1. Introduction .......................................................................................................... 1
1.2. The Mitochondrial Energy-Redox Axis ............................................................... 4
1.3. The Energy-Redox Axis with Cytosolic Signaling ............................................ 17
1.4. Significance ........................................................................................................ 23
1.5. Hypothesis and Specific Aims ........................................................................... 25
CHAPTER 2 .....................................................................................................................27
SHORT-TERM CALORIC RESTRICTION ATTENUATES
AGE-DEPENDENT IMPAIRMENT OF THE MITOCHONDRIAL
ENERGY-REDOX AXIS IN BRAIN
2.1. Abstract .............................................................................................................. 27
2.2. Introduction ........................................................................................................ 29
2.3. Materials and Methods ....................................................................................... 32
2.4. Results ................................................................................................................ 39
2.5. Discussion .......................................................................................................... 49
v
CHAPTER 3 .....................................................................................................................54
SILENCING OF NICOTINAMIDE NUCLEOTIDE
TRANSHYDROGENASE IMPAIRS CELLULAR REDOX
HOMEOSTASIS AND ENERGY METABOLISM IN PC12 CELLS
3.1. Abstract .............................................................................................................. 54
3.2. Introduction ........................................................................................................ 56
3.3. Materials and Methods ....................................................................................... 58
3.4. Results ................................................................................................................ 63
Table 3-1. GSH and GSSG Levels and the Redox Potential of the Cell .................. 68
3.5. Discussion .......................................................................................................... 80
3.6. Conclusion .......................................................................................................... 83
CHAPTER 4 ....................................................................................................................84
LOSS OF NICOTINAMIDE NUCLEOTIDE
TRANSHYDROGENASE IN C57BL/6J MICE INDUCES
DISTINCT RESPONSES IN BRAIN AND IN LIVER
4.1. Abstract .............................................................................................................. 84
4.2. Introduction ........................................................................................................ 85
4.3. Materials and Methods ....................................................................................... 89
4.4. Results ................................................................................................................ 93
4.5. Discussion ........................................................................................................ 111
CHAPTER 5 ...................................................................................................................117
CONCLUSIONS
BIBLIOGRAPHY ..........................................................................................................120
vi
LIST OF FIGURES
Figure 1-1. Role of the mitochondrial energy-redox axis in maintenance
of cellular H
2
O
2
levels and redox signaling 5
Figure 1-2. Metabolism of pyruvate and ketone bodies by brain mitochondria 8
Figure 1-3. NNT catalyzed generation of NADPH 16
Figure 1-4. Overview of specific aims 1-3 26
Figure 2-1. Body weight of rats during short-term CR 33
Figure 2-2. Aging and short-term CR alter the regulation of substrate
entry to TCA cycle 41
Figure 2-3. Mitochondrial respiration in AL and CR rat brain of
different age groups 43
Figure 2-4. Mitochondrial H
2
O
2
generation and S-glutathionylation
in aging and CR. 45
Figure 2-5. Assessment of the mitochondrial Trx2-Prx3 system
in aging and CR 47
Figure 2-6. Modulation of NNT activity and pyridine dinucleotide
levels by aging and CR 49
Figure 3-1. siRNA knockdown of the NNT gene in PC12 cells 64
Figure 3-2. Effect of NNT knockdown on NADPH production
and H
2
O
2
release 66
Figure 3-3. Cellular energy metabolism in siNNT-transfected PC12 cells 71
Figure 3-4. Effect of NNT knockdown on mitochondrial
bioenergetic machinery 73
vii
Figure 3-5. Alteration of redox status after NNT knockdown
precedes the impairment of energy metabolism 76
Figure 3-6. PDH and SCOT activities in siNNT-transfected PC12 cells 77
Figure 3-7. Activation of JNK and initiation of apoptosis
in siNNT-transfected PC12 cells 79
Figure 4-1. Loss of NNT function moderately affects
mitochondrial NADP(H) pool in brain 94
Figure 4-2. Loss of NNT function does not affect mitochondrial
NAD(H) pool in brain 96
Figure 4-3. Loss of NNT function elicits substantially impacts
on mitochondrial NADP(H) pool in liver 98
Figure 4-4. Mitochondrial NAD(H) pool in liver is perturbed
by loss of NNT function 100
Figure 4-5. Less effect of NNT dysfunction on mitochondrial redox
homeostasis is associated with upregulation of IDH2 in brain 102
Figure 4-6. Loss of NNT function in hepatocytes affects mitochondrial
NADP(H) and NAD(H) pools but not on expression of other
redox enzymes 104
Figure 4-7. Cellular energy metabolism in primary hepatocytes of
WT and NNT mutant mice 107
Figure 4-8. Loss of NNT function induces the modulation of
cytosolic redox-sensitive signaling 110
viii
ABBREVIATIONS
Ad libitum AL
Protein kinase B Akt
5' adenosine monophosphate-activated protein kinase AMPK
Caloric restriction CR
Extracellular acidification rate ECAR
Glutathione peroxidase GPx
Glutaredoxin Grx
Glutathione GSH
Glutathione disulfide GSSG
High-performance liquid chromatography HPLC
Isocitrate dehydrogenase IDH
Insulin-like growth factor-1 IGF-1
Insulin/IGF-1 signaling IIS
c-Jun N-terminal kinase JNK
α-ketoglutarate dehydrogenase α-KGDH
Mitogen-activated protein kinases MAPK
Nicotinamide nucleotide transhydrogenase NNT
Oxygen consumption rate OCR
Oxidative phosphorylation OXPHOS
ix
Pyruvate dehydrogenase PDH
Potential of hydrogen PH
Phosphatidylinositol 3-kinase PI3K
Peroxiredoxin Prx
Respiratory control ratio RCR
Sodium dodecyl polyacrylamide gel electrophoresis SDS-PAGE
Succinyl-CoA:3-oxoacid Co-A SCOT
Tricarboxylic acid TCA
Thioredoxin Trx
Thioredoxin reductase TrxR
Voltage dependent anion channel VDAC
x
ABSTRACT
The mitochondrial energy-transducing capacity is essential for the maintenance of
neuronal function: impairment of energy metabolism and redox homeostasis –integrated
in the mitochondrial energy-redox axis– is a hallmark of brain aging and is accentuated in
the early stages of neurodegenerative diseases. The energy component of the axis entails
the formation of reducing equivalents (NADH) and their flow through the respiratory
chain with consequent electron leak to generate O
2
.–
and H
2
O
2
. The redox component of
the axis entails the removal of H
2
O
2
by NADPH-dependent, thiol-based systems.
Mitochondrial NADPH generation is largely dependent on nicotinamide nucleotide
transhydrogenase (NNT) that catalyzes the reduction of NADP
+
to NADPH utilizing the
proton gradient as the driving force and NADH as the electron donor, thereby linking the
energy- and redox components of the axis.
The hypothesis to be tested is that NNT activity is critical for the maintenance of the
mitochondrial energy-redox axis and is compromised during aging leading to
mitochondrial dysfunction and loss of cellular redox homeostasis. This hypothesis was
tested by three specific aims that encompassed three different experimental models.
I have shown that NNT dysfunction impairs cellular redox homeostasis and energy
metabolism in PC12 cells. Knockdown of NNT results in decreased cellular NADPH
supply, increased H
2
O
2
levels, and increased redox potential. Altered redox status further
leads to the impairment of mitochondrial bioenergetic function through the activation of
xi
redox-sensitive c-Jun N-terminal kinase (JNK) signaling and concomitant inhibition of
pyruvate dehydrogenase. Active JNK also initiates mitochondrion-dependent intrinsic
apoptosis after NNT suppression.
The role of NNT in linking the mitochondrial energy status to the redox environment
was further investigated in brain and liver of C57BL/6J mice –essentially NNT knockout
mice– that show impaired glucose tolerance independent of obesity. The results showed
that NNT regulates mitochondrial NADPH levels in an energy-sensitive manner in both
tissues but with the NADP pool in the liver more affected. In primary hepatocytes of
C57BL/6J mice, loss of NNT also leads to impaired mitochondrial energy-transducing
capacity, a process related by the co-regulation of redox-sensitive signaling pathways.
The diminished impact of NNT in brain was ascribed to the upregulation of another
mitochondrial NADPH-generating enzyme, isocitrate dehydrogenase-2.
The functional changes of the mitochondrial energy-redox axis in brain were further
characterized in an aging model and the effects of short-term caloric restriction (CR)
were assessed. It was shown that the substrate supply (pyruvate and ketone bodies) and
mitochondrial energy-transducing capacity are enhanced in 26 month-old rats following
short-term CR; likewise, short-term CR increases the expression and activity of
mitochondrial redox systems which is associated with a more reduced redox environment
in old animals. Short-term CR also rescued the age-dependent decline of NNT activity
and the NADPH/NADP
+
pool. However, none of these effect elicited by short-term CR
xii
were observed in young rats, indicating a different response to short-term CR in animals
with different ages.
Taken together, these studies validated the hypothesis above and confirmed that age-
dependent impairment of cellular bioenergetics and redox homeostasis should be viewed
as an interdependent energy-redox axis integrated by the activity of NNT. I have shown
that NNT dysfunction disrupts the electron flux from fuel substrates to redox
components, and induces not only mitochondrial dysfunction but also leads to
impairment of redox sensitive signaling that, in turn, exacerbates further mitochondrial
dysfunction. Hence, dysregulation of mitochondria-cytosol communication is a critical
event in the progression of aging and, likely age-related neurodegeneration.
1
CHAPTER 1
OVERVIEW OF THE MITOCHONDRIAL ENERGY METABOLISM AND
REDOX SIGNALING IN BRAIN AGING AND NEURODEGENERATION
1.1. INTRODUCTION
Brain, like most organs, undergoes a gradual decline in energy metabolism during
aging (Boveris and Navarro, 2008; Drew and Leeuwenburgh, 2004; Navarro and Boveris,
2007b; Swerdlow, 2011). Because neurons require large amounts of energy for firing
action potentials, neurotransmission, and other processes, the age-related decline in
metabolism contributes to the cognitive declines associated with aging (Biessels and
Kappelle, 2005; Boveris and Navarro, 2008). Clinically, age-dependent reduction of
glucose utilization was observed in most human brain regions (Petit-Taboue et al., 1998).
Similarly, an age-dependent decrease in O
2
uptake was observed in rodent brain (Navarro
and Boveris, 2008). Aging is also a risk factor for age-associated diseases such as
neurodegenerative disorders. These diseases may occur when neurons fail to respond
adaptively to age-related decline in basal metabolic rates and in energy-driven tasks, such
as neuromuscular coordination, cognitive performance, and environmental awareness
(Swerdlow, 2007). In humans, cerebral glucose hypometabolism is an early and
2
consistent event in the progression of Alzheimer's disease, Parkinson’s disease,
Huntington’s disease, and mild cognitive impairment, before the onset of pathologies in
the brain (Atamna and Frey, 2007; Dagher, 2001; Feigin et al., 2001) and decreased
frontal cortex O
2
uptake has been reported in Parkinson’s disease and in dementia with
Lewy bodies (Navarro et al., 2009).
The energy-transducing capacity of mitochondria meets the cellular energy demands,
thus supporting metabolic, osmotic, and mechanical functions; they are sources of H
2
O
2
,
and play a pivotal role as mediators of the intrinsic apoptotic pathway. The most
prominent metabolic process carried out by mitochondria is oxidative phosphorylation
(OXPHOS) to generate ATP, the universal energy currency. On the other hand, high
levels of H
2
O
2
have been associated with mitochondrial redox changes and
macromolecule oxidation during aging and are believed to mediate the detrimental effects
associated with mitochondrial dysfunction in brain aging (Balaban et al., 2005) and
neurodegenerative disorders (Barnham et al., 2004; Beal, 1995; Simonian and Coyle,
1996). The cellular composition of brain consists mainly of terminally differentiated
neurons, and its regenerative capacity is relatively reduced as compared to other organs
such as liver. Thus, the brain is highly susceptible to neuronal loss due to hypometabolic
states and impairment of redox homeostasis. Age-related changes in energy production
and redox status cannot be viewed as independent variables, but rather as an
interdependent relationship reflected in the mitochondrial energy-redox axis that
3
represents a dual pronged approach to assess the changes in mitochondrial function as a
function of age and disease (Yap et al., 2009).
Mitochondrion-generated signaling molecules, such as H
2
O
2
, that report the
mitochondrial energy charge to the cytosol (Valdez et al., 2006) are implicated in the
regulation of the cellular redox status, thus transducing redox signals into a variety of
responses, such as proliferation, adaptation, and cellular death pathways (Moran et al.,
2001). Low to intermediate levels of H
2
O
2
are involved in the regulation of redox-
sensitive signaling and transcription whereas high levels are involved in oxidative
damage to cell constituents. The release of oxidants from mitochondria as a function of
the mitochondrial metabolic and redox states serves as a coordinated response between
these seemingly autonomous organelles and the rest of the cell through the modulation of
redox-sensitive signaling and transcription pathways. Conversely, mitochondria are the
recipients of cytosolic signaling, such as mitogen-activated protein kinases (MAPKs) and
the Phosphatidylinositol 3-kinase/Protein kinase B (PI3K/Akt) pathway of insulin
signaling, that elicit profound changes in the mitochondrial energy-transducing capacity.
4
1.2. THE MITOCHONDRIAL ENERGY-REDOX AXIS
Mitochondria provide most of the energy needed for cellular functions by the
metabolism of fuel molecules into ATP through oxidative phosphorylation. The
generation of ATP entails the oxidation of acetyl-CoA in the tricarboxylic acid (TCA)
cycle with concomitant generation of reducing equivalents (NADH, FADH
2
) that flow
through the respiratory chain generating a proton motive force (Mathews et al., 2000);
electron leakage leads to the generation of O
2
.–
, which further disproportionates to H
2
O
2
,
either catalyzed by the matrix Mn-superoxide dismutase (SOD2) or, secondarily, through
spontaneous dismutation (Melov, 2000). Steady-state levels of mitochondrial H
2
O
2
are
determined by both energy metabolism and the redox systems (Fig. 1-1). A decrease in
the mitochondrial energy-transducing capacity is a common feature of brain aging and
neurodegeneration and is associated with a progressive increase of H
2
O
2
steady-state
concentrations that can shift the cell from a reduced state to an oxidized state. Thus,
maintenance of mitochondrial redox homeostasis becomes crucial for cell function.
Mitochondrial energy metabolism – The effects of aging on mitochondrial energy
metabolism are tissue-specific and are more prominent in tissues that contain mostly
post-mitotic cells such as heart, skeletal muscle, and brain. Partial loss of the energy-
transducing capacity, attributed to changes in protein expression, has been documented in
mitochondria isolated from old animals. Glucose is the primary fuel for brain, whereas
5
Figure 1-1. Role of the mitochondrial energy-redox axis in regulation of cellular H
2
O
2
levels and redox signaling. Reducing equivalents from the tricarboxylic acid (TCA) cycle
flow through the respiratory chain (RC); electron leak accounts for 2-3% of O
2
consumed in
the form of O
2
.–
and H
2
O
2
. Reduction of H
2
O
2
is supported by thiol-based systems, for which
the ultimate reductant is NADPH. Sources of mitochondrial NADPH: nicotinamide nucleotide
transhydrogenase (NNT), isocitrate dehydrogenase-2 (IDH
2
), and malic enzyme. Domain-
specific signaling entails regulation of redox-sensitive JNK- and insulin/IGF1 (IIS) pathways.
6
metabolism of ketone bodies represents an alternative fuel source during glucose
deprivation (Grinblat et al., 1986) (Fig. 1-2). Pyruvate, generated from glycolysis,
undergoes oxidative decarboxylation by the pyruvate dehydrogenase complex (PDH) to
acetyl-CoA that feeds into the TCA cycle. PDH activity in brain was found to decrease
with age (Zhou et al., 2008; Zhou et al., 2009). Additionally, there is an age dependent
decrease in succinyl-CoA:3-oxoacid Co-A (SCOT) transferase activity (Lam et al.,
2009), a key mitochondrial matrix enzyme that metabolizes ketone bodies to acetyl-CoA;
the decrease SCOT activity as a function of age was due to irreversible protein post-
translational modifications (Lam et al., 2009). In a triple transgenic mouse model of
Alzheimer’s disease, ketone body metabolism is a temporary mechanism that prevents
the further decline of brain mitochondrial bioenergetic capacity (Yao et al., 2010) that is
associated with decreased activities of PDH and cytochrome oxidase. 2-Deoxy-D-glucose
treatment induced ketogenesis in the same mouse model and this resulted in increased
ketone body metabolism in the brain and a significant reduction of both amyloid
precursor protein and amyloid-β(Yao et al., 2010). The activities of TCA enzymes, such
as aconitase and α-ketoglutarate dehydrogenase (α-KGDH) also decline as a function of
age (Yarian et al., 2006) and the activities of PDH, α-KGDH, and isocitrate
dehydrogenase (IDH) are also lower in Alzheimer’s disease (Bubber et al., 2005; Yao et
al., 2010). It may be surmised that alterations in activities of TCA cycle enzymes and of
enzymes controlling the entry of acetyl-CoA into the TCA cycle, such as PDH and
7
SCOT, affect NADH levels and contribute significantly to the decline in mitochondrial
bioenergetics during aging and neurodegeneration.
The exergonic electron transfer through the complexes I to IV of the respiratory chain
is the driving force for the vectorial H
+
release into the inter-membrane space and H
+
re-
entry to the matrix through F
0
of complex V with ATP synthesis by F
1
-ATP synthase.
Electron transfer in mitochondria decreases in aged brain (Beckman and Ames, 1998;
Navarro and Boveris, 2007b), with more marked changes in complexes I, III, and IV
(Kwong and Sohal, 2000; Navarro and Boveris, 2004, 2007a; Navarro et al., 2008). The
inhibition of complex I activity upon aging occurs with a decrease in NAD
+
levels
(Boveris and Cadenas, 2000) that leads to impairment of the turnover efficiency of the
TCA cycle, irrespective of the presence of acetyl-CoA. Moreover, reduced electron
transfer can also lead to decreased mitochondrial inner membrane potential which is
observed in aged rat brain (LaFrance et al., 2005; Sastre et al., 1998). The F
1
-ATPase
activity of complex V also decreases with age due to nitration of Tyr
269
close to the Mg
++
binding site of the F1β subunit (Lam et al., 2009).
8
Figure 1-2. Metabolism of pyruvate and ketone bodies by brain mitochondria. Glucose is
the primary fuel for brain and ketone bodies the secondary fuel; metabolism of pyruvate (from
glucose) is regulated by the pyruvate dehydrogenase complex (PDH); metabolism of ketone
bodies requires the activity of succinyl-CoA transferase (SCOT) (which is expressed in brain
mitochondria). Acetyl-CoA, generated by PDH or SCOT activities, is further oxidized in the
tricarboxylic acid cycle with formation of NADH. The arrows indicate protein post-
translational modifications found in brain as a function of age: (a) phosphorylation
(inactivation) of PDH upon translocation of JNK to the outer mitochondrial membrane; (b)
and (c) nitration of SCOT and F
1
-ATPase, respectively upon diffusion of
.
NO to mitochondria
due to increased expression and activity of nNOS as a function of age. GLUT, glucose
transporter; MCT, monocarboxylate transporters.
GLUT
glucose
ketone
bodies
glucose
pyruvate
acetyl-CoA
succinyl-CoA
succinate
acetoacetate
pyruvate acetoacetate
MCT
NADH
PDH
SCOT
CO
2
+ NADH
HSCoA + NAD
TCA
lactate
acetoacetyl-CoA
O
2
H
2
O H
+
e
–
III
I
IV
H
+
ATP
ADP
e
–
V
a
b
c
9
The operational concepts of mitochondrial metabolic states and respiratory control are
defined as state 4 (resting or proton motive force controlled-respiration), with the
availability of respiratory substrates but not ADP, and state 3 (active respiration) with
ample respiratory substrate and ADP availability (Chance and Williams, 1955).
Mitochondrial respiration decreases in aging in terms of marked decline in state 3
respiration and the related respiratory control ratio (RCR) and membrane potential, as
well as an increase in state 4 respiration (Boveris et al., 1999; Lam et al., 2009; Navarro
et al., 2008), all of which indicate a lower energy-transducing efficiency.
Mitochondrial redox homeostasis – Following to the initial reports in intact heart and
liver mitochondria as an active source of H
2
O
2
by Chance and Boveris (Boveris and
Chance, 1973; Boveris et al., 1972), further work established that superoxide anion (O
2
.–
)
was the stoichiometric precursor of mitochondrial H
2
O
2
and that it was primarily
generated during ubisemiquinone auto-oxidation (Boveris and Cadenas, 1975; Boveris et
al., 1976; Cadenas et al., 1977) and, secondarily, by reverse electron transfer at the
NADH-dehydrogenase segment (Turrens and Boveris, 1980). Components of complex I
and complex III were reported to generate O
2
.–
(Cadenas, 2004; Turrens, 2003). As the
activities of complexes I, III, and IV decrease during aging, higher oxidant production is
observed: the rates of O
2
.–
and H
2
O
2
formation increase with age and are higher in
mitochondria from tissues of ad libitum-fed mice than in those on caloric-restricted diets
(Lass et al., 1998; Sohal et al., 1994). O
2
.–
, formed upon oxidation of the outer membrane
10
UQ pool (UQ
O
) can be vectorially released into the cytosol, in part, through a voltage-
dependent anion channel (VDAC) (Han et al., 2003). Thus, cytosolic levels of H
2
O
2
reflect the mitochondrial energy status, because mitochondrial H
2
O
2
generation in state 4
respiration is about 4-5 times higher than that during effective oxidative phosphorylation
(state 3 respiration) (Boveris et al., 1999). A comprehensive review by Murphy
recognizes four main determining factors that regulate O
2
.–
generation in mitochondria:
the ratios UQH
2
/UQ and NADH/NAD
+
, the Δψ of the inner membrane, and the local
mitochondrial [O
2
] (Murphy, 2009).
In brain mitochondria, H
2
O
2
is eliminated mainly by GSH- or thioredoxin (Trx)-
driven catalysts that depend on NADPH as ultimate electron donors.
Glutathione-based systems – Glutathione (GSH), synthesized in the cytosol from
glycine, glutamate, and cysteine in a two-step process by the enzymes γ-glutamylcysteine
synthetase and GSH synthase (Griffith, 1999), is imported into the mitochondria through
the dicarboxylate- and oxoglutarate carriers on the inner mitochondrial membrane
(Griffith and Meister, 1985; Zhong et al., 2008). The role of mitochondrial thiols in redox
signaling (Murphy, 2012) and cell death pathways (Yin et al., 2012a) has been recently
reviewed. The mitochondrial redox potential, calculated from the GSH/GSSG or
Trx2
red
/Trx2
ox
levels is approximately –300 mV and –340 mV (Jones, 2002; Kemp et al.,
2008), respectively. The mitochondrial GSH pool functions independently from the
cytosolic pool in response to the production of oxidants in the mitochondrial environment
11
(Hurd et al., 2005). Mouse brain showed a decreased GSH/GSSG ratio as a function of
age (Rebrin et al., 2007; Rebrin et al., 2003).
Mitochondrial GSH protects against oxidative stress largely as a cofactor for
glutathione peroxidases, glutathione-S-transferases, sulfiredoxins, and glutaredoxins
(Marí et al., 2009; Rhee and Woo, 2011). Glutathione peroxidase-1 (GPx1) localizes
mainly in the mitochondrial matrix, whereas glutathione peroxidase-4 (GPx 4; also
referred as phospholipid hydroperoxide glutathione peroxidase) (Schuckelt et al., 1991;
Ursini et al., 1997) occurs in the inter-membrane space; the latter detoxifies mainly
phospholipid hydroperoxides and its significance is underscored by the embryonic
lethality that follows systemic ablation of GPx4, explained in part by studies on the
expression of GPx4 in embryonic brain and its role in organogenesis (Borchert et al.,
2006).
Protein thiols are sensitive to changes in the redox environment (Cumming et al.,
2004): the reversible formation of mixed disulfides entails a reaction between protein
cysteine sulfhydryls and GSH in a process termed S-glutathionylation. Mixed-disulfide
formation affects the activity of enzymes, transcription factors, and transporters, thus
enabling them to respond to the redox environment by reversible activation/inactivation
(Fratelli et al., 2002; Thomas et al., 1995; Zheng et al., 1998). Thus, S-glutathionylation
reflects the redox status of the mitochondria (Schafer and Buettner, 2001) and is viewed
as a regulatory device for proteins involved in energy metabolism, redox signaling, and
cell function (Dalle-Donne et al., 2011; Dalle-Donne et al., 2008; Hill et al., 2010; Klatt
12
and Lamas, 2000). A number of proteins have been identified to be S-glutathionylated
during oxidative conditions, including components of the energy metabolism: (a) SCOT
and the E
2
subunit of PDH (Garcia et al., 2010; Hurd et al., 2005); (b)TCA cycle enzyme
such as aconitase (Han et al., 2005), α-KGDH (Nulton-Persson et al., 2003), and IDH
(Kil and Park, 2005), and (c) complexes I (Taylor et al., 2003), II (Chen et al., 2007), and
V (Garcia et al., 2010; West et al., 2006). S-glutathionylation of SCOT and ATP synthase
(F
1
complex, -subunit) in brain mitochondria results in a decrease of activity and a
substantially lower reduction potential (–171 mV); energized mitochondria with complex
I or complex II respiratory substrates increased NADH and NADPH levels, led to
reduction of GSSG thus increasing GSH levels, elicited deglutathionylation of
mitochondrial proteins, and resulted in a more reducing mitochondrial environment (–291
mV) (Garcia et al., 2010). Complex I is persistently glutathionylated under conditions of
oxidative stress and this resulted in increased generation of O
2
.–
and decreased
mitochondrial function (Taylor et al., 2003). Conversely, S-glutathionylation of adenine
nucleotide translocase (ANT) protects against mitochondrial membrane permeabilization
and apoptosis (Queiroga et al., 2010). These data provide evidence of mitochondrial
redox changes that modulate energy metabolism through protein thiol modifications.
The reversible formation of protein-GSH mixed disulfides has been suggested as a
protective mechanism that masks critical sulfhydryls from irreversible oxidation; the
reversibility of this process acquires further significance because of its involvement in the
redox regulation of signal transduction (Hill and Darley-Usmar, 2008; Klatt and Lamas,
13
2000). Protein mixed disulfides are specifically reduced by glutaredoxins (Grxs; Grx2 is
the mitochondrial isoform) through a monothiol mechanism (Holmgren and Aslund,
1995). Oxidized Grx2 is reduced by GSH, which is regenerated from GSSG by NADPH-
supported glutathione reductase (GR). Grx2 is constitutively expressed in neurons and
glia in mouse and human brain (Karunakaran et al., 2007). Knockdown of cytosolic Grx1
is associated with a loss of mitochondrial membrane potential (Saeed et al., 2008), and
Grx1 is essential for maintaining the functional integrity of brain mitochondrial complex
I (Kenchappa and Ravindranath, 2003). Grx2 protects cells against oxidative damage
(Enoksson et al., 2005) involving Akt signaling and the redox-sensitive transcription
factor NF- B and anti-apoptotic Bcl-2 (Nagy et al., 2008). Additionally, Grx2 has been
characterized as part of an iron-sulfur cluster that senses redox changes and controls Grx2
activity (Holmgren et al., 2005), thus expanding the interaction between oxidants,
mitochondrial redox status, and protein glutathionylation.
Thioredoxin-based systems – The reducing power for peroxiredoxins (Prx) is
transmitted through thiols of the Trx system: NADPH → TrxR → Trx → Prx
(Zhang et
al., 2007). A comprehensive study on immunohistochemical mapping of all six Prx
subtypes in mouse brain revealed that astrocytes and microglia were reactive to Prx6 and
Prx1, respectively; immunoreactivity for Prx1 and Prx4 in the nuclei of oligodendrocytes;
in neurons, Prx3 and Prx5 were found in the stratum lucidum of the hippocampus and
Prx2 in the habenular nuclei (Jin et al., 2005). Of these Prxs, Prx2 is critical for the
14
maintenance of hippocampal synaptic plasticity against age-associated oxidative damage
(Kim et al., 2011) by a mechanism entailing the oxidant- and age-dependent
mitochondrial decay of hippocampal neurons; also the expression of Prx2 in hippocampal
neurons increased as a function of age.
Mitochondrial Prx3 and Prx5 are involved in the enzymatic degradation of H
2
O
2
;
Prx5 can also reduce ONOO
–
(Dubuisson et al., 2004; Peng et al., 2004). Trx2 is highly
efficient at reducing disulfides in proteins (Miranda-Vizuete et al., 2000), thus impacting
cellular functions such as antioxidant defenses and redox control of transcription and
signal transduction (Aslund and Beckwith, 1999; Holmgren, 1989). Using a
polarographic method for real-time detection of H
2
O
2
, it was concluded that removal of
H
2
O
2
by energized brain mitochondria was largely dependent on the Trx/Prx system with
a modest contribution by the GSH/GPx system (Drechsel and Patel, 2010). Prx3 and Prx5
are expressed constitutively in human neuroblastoma Sh-SY5Y cells and their silencing
by small hairpin RNAs renders the cells more susceptible to oxidative damage and
apoptosis (De Simoni et al., 2008). Prx3 protects hippocampal neurons against
excitotoxicity and its upregulation prevented or reduced gliosis (Hattori et al., 2003).
Overexpression of Prx3 reduces H
2
O
2
production and lipid peroxidation and protects
cells from hypoxia-, TNFα-, cadmium-, and oxidant-induced cell death (Chang et al.,
2004; Chen et al., 2008; Nonn et al., 2003; Wonsey et al., 2002) and a neuroprotective
effect is observed when it is administered into ischemic brains (Hwang et al., 2010). The
increased Prx3 (along with TrxR2 and Trx2) immunoreactivity in the hippocampus of
15
aged dogs as compared with adult dogs leads to a reduction of neuronal damage (against
oxidative stress) during aging (Ahn et al., 2011). Prx3 levels are found significantly lower
in the brains of Alzheimer’s disease patients (Kim et al., 2001) and deficiency in Prx3 is
also associated with multiple neurodegenerative disorders such as Parkinson’s disease,
amyotrophic lateral sclerosis, and Down syndrome (Krapfenbauer et al., 2003; Wood-
Allum et al., 2006). Trx2
+/-
mice show reduced ATP production and electron-transport
chain rates (Perez et al., 2008); this notion is further supported by the increased apoptosis
in early embryos of Trx2
-/-
mice (embryonic lethal) with mitochondria maturation.
Systemic ablation of mitochondrial thioredoxin reductase-2 also yielded embryonic lethal
phenotypes (Conrad, 2009). The levels of TrxR-2 decrease with age in muscle and is
accompanied by enhanced susceptibility to apoptosis (Rohrbach et al., 2006 ). The
significance of these redox couples and redox catalysts for brain aging and age-related
neurodegeneration is underscored by the highly oxidized mitochondrial and cellular
redox environment in these processes (Lin and Beal, 2006; Navarro and Boveris, 2004;
Yin et al., 2012a).
Interdependence of energy- and redox components – role of NNT – The ultimate
reductant of mitochondrial redox systems is NADPH that supports the activities of
thioredoxin reductase and glutathione reductase. The NADPH/NADP
+
ratio decreases in
kidney mitochondria as a function of age (Yarian et al., 2006), thus indicating that
NADPH deficiencies are another factor in mediating mitochondrion-dependent aging.
16
Mitochondrial sources of NADPH include NADP
+
-dependent IDH2, malic enzyme, and
nicotinamide nucleotide transhydrogenase (NNT). It is believed that NNT contributes to
more than 50% of the mitochondrial NADPH production (Rydstrom, 2006). NNT is
encoded in nuclear and targeted to the inner mitochondrial membrane (Hoek and
Rydström, 1988 ), and it catalyzes the reversible reduction of NADP
+
to NADPH coupled
to the oxidation of NADH to NAD
+
and proton pumping from the inter membrane space
to the matrix (Fig. 1-3). The forward reaction, i.e., the generation of NADPH is strongly
favored due to the existence of proton motive force across the inner membrane (Ying,
2008).
Figure 1-3. NNT catalyzed generation of NADPH. The reversible reaction catalyzed by
NNT is strongly toward the generation of NADPH by consumption of the proton gradient
across the mitochondrial inner-membrane.
17
1.3. THE ENERGY-REDOX AXIS WITH CYTOSOLIC SIGNALING
The energy-transducing and redox-regulation capacities of mitochondria are highly
affected in aging and age-related neurodegeneration. Mitochondria generate second
messengers (redox: H
2
O
2
; energy: ATP) that are involved in the regulation of
redox/energy sensitive cell signaling pathways, thus coordinating functional responses
between mitochondria and other cellular processes. Conversely, mitochondria are the
recipients of cytosolic signaling molecules, which translocate to mitochondria under
specific conditions and elicit profound metabolic or redox effects in the organelles. The
communication between mitochondria and other components of the cell establishes a
regulatory devise that controls cellular energy levels and the redox environment;
impairment of this regulatory devise may be the basis for the mechanisms inherent in
aging and age-related degenerative disorders.
Mitochondrial regulation of cytosolic signaling – Mitochondrial-generated oxidants
regulate important signaling pathways such as the insulin/IGF-1 signaling (IIS) and the
MAPK (e.g., JNK) pathways. The PI3K/Akt route of insulin/IGF-1 signaling in the brain
is implicated in neuronal survival and synaptic plasticity via, among other effects,
maintenance of the metabolic function of mitochondria. Aging is associated with
decreases in the levels of both insulin/IGF-1 and their receptor (Frolich et al., 1998).
Mitochondrial H
2
O
2
is involved in the regulation of insulin signaling, which is not
18
surprising given the large quantity of redox-sensitive cysteine residues on the insulin and
IGF1 receptors and insulin receptor substrates (IRS): oxidation of specific cysteine
residues promote tyrosine autophosphorylation of the insulin receptor (Storozhevykh et
al., 2007) and inhibition of phosphatases (PTEN and PTP1B) involved in the insulin
receptor substrate (IRS) node of insulin signaling upon oxidation of critical cysteines to
disulfides (Mahadev et al., 2001). Aged cells are more vulnerable to H
2
O
2
-induced
apoptosis, which is accompanied with reduced activation of Akt and CR can prevent this
loss of Akt activation (Ikeyama et al., 2002). Akt also inhibits GSK3β upon
phosphorylation at Ser
9
and thereby protects cell against apoptosis, because activated
GSK3β stimulates phosphorylation of the anti-apoptotic member of the Bcl-2 family,
Mcl-1, thus leading to its degradation and the ensuing cytochrome c release and apoptosis
(Maurer et al., 2006; Pap and Cooper, 1998).
The MAPK JNK is also redox sensitive and oxidative stress conditions, entailing also
enhanced generation of mitochondrial H
2
O
2
, result in its activation (Nemoto et al., 2000;
Zhou et al., 2008); H
2
O
2
may act at multiple levels to activate JNK (and p38):
dissociation of thioredoxin from the ASK-1 complex (Saitoh et al., 1998), disruption of
the glutathione transferase (GT)-JNK complex (Adler et al., 1999), or inhibition of
MAPK phosphatase activity (Foley et al., 2004). Basal JNK activity, but not its protein
levels, is increased in mouse brain and liver, rat kidney and splenic lymphocytes, and
human skeletal muscle during aging (Hsieh et al., 2003; Kim et al., 2002; Li et al., 2000;
Suh, 2001; Williamson et al., 2003). Basal activities of ERK and p38 kinase, but not their
19
protein levels, are reported to decrease in brain cortex during aging, a phenomenon that
prevented by caloric restriction (Zhen et al., 1999); conversely, basal p38 and ERK
activities were increased in mouse liver, rat kidney, and human skeletal muscle (Hsieh
and Papaconstantinou, 2002; Kim et al., 2002; Williamson et al., 2003). It is unclear
whether or not these discrepancies are due to tissue specificity of p38 and ERK responses
to the aging process. Nevertheless, activation of ERK in response to epithelial growth
factor is decreased in cortical brain slices and hepatocytes from old rats (Liu et al., 1996;
Zhen et al., 1999), indicating reduced sensitivity to stimuli in these tissues during aging.
In addition, JNK also plays a central role in the progression of insulin resistance; a
likely mechanism entails the phosphorylation of the insulin receptor substrate-1 (IRS-1)
at Ser
307
, leading to inhibition of the insulin-promoted tyrosine phosphorylation of IRS
(Aguirre et al., 2000). Conversely, the MLK3-mediated JNK activation is inhibited by
Akt upon phosphorylation of MLK3 both in vitro and in vivo (Barthwal et al., 2003).
Because of the distinct downstream signaling between PI3K/Akt and JNK (survival
versus apoptosis; growth versus differentiation), the counterbalance of the IIS and JNK
pathways induced by different concentrations of H
2
O
2
is expected to determine the
coordinated response of the cell. These disparate effects of mitochondrial H
2
O
2
are
important, for they refer to a healthy aging or accelerated aging, and they need be
assessed in terms of the cellular peroxide tone, i.e., a quantitative assessment of
mitochondrial steady-state concentration of H
2
O
2
in connection with domain specific
20
signaling. Hence, the mitochondrial energy-redox axis is one of the factors that regulate
the peroxide tone of the cell in a domain-specific signaling fashion.
Cytosolic regulation of mitochondrial function – Mitochondria are also important
targets of cytosolic signaling molecules. It was shown that the expression and activation
of JNK1 increases in brain as a function of age and that active (bisphosphorylated) JNK
translocates to the mitochondrion where it triggers a phosphorylation cascade that results
in phosphorylation (inhibition) of pyruvate dehydrogenase (PDH), a key mitochondrial
enzyme complex that bridges anaerobic and aerobic brain energy metabolism. PDK2 is
an essential intermediate in this phosphorylation cascade. The outcome is a bioenergetic
crisis translated in decreased cellular ATP and increased lactate levels (anaerobic
glycolysis as a compensatory mechanism) (Zhou et al., 2008; Zhou et al., 2009).
The PI3K/Akt pathway promotes neuronal survival and synaptic plasticity (van der
Heide et al., 2006) by mechanisms entailing phosphorylation of proapoptotic Bcl-2
family members (Bad), of GSK3 (thus inhibiting tau hyperphosphorylation), and of FoxO
factors (that drives nuclear FoxO factors to the cytosol, thus inhibiting the transcription of
some apoptotic genes and those involved in heme degradation) (Cheng et al., 2010). In
NIH/3T3 cell lines mitochondrial H
2
O
2
modulates the entrance of cytosolic Akt
(phosphorylated at Ser
473
) to mitochondria and induces the further phosphorylation at
Thr
308
that is required for nuclear translocation (Antico Arciuch et al., 2009). Akt
translocates to the mitochondrion in human neuroblastoma cells and its phosphorylation
21
targets are a constitutive form of GSK3β in the mitochondrion and the β-subunit of
ATPase (Bijur and Jope, 2003).
ATP is the universal energy currency in the cell and mitochondrial produced ATP is
transported to the cytosol by ANT in exchange with ADP. The cytosolic ADP/ATP ratio
is an important parameter of not only the cellular consumption of ATP but also of ATP
synthesis as a reflection of mitochondria bioenergetic profile. The ubiquitous expressed
adenylate kinases in all cell types, which catalyze the inter-conversion of adenine
nucleotides (2ADP↔ATP+AMP), make AMP/ATP another important indicator of
energy status. 5' adenosine monophosphate-activated protein kinase (AMPK) is a kinase
with its activity controlled by intracellular AMP/ATP ratio and therefore rendered a
sensor of cellular energy status. In fact, recent studies have suggested that AMPK activity
can also be regulated by ADP (Oakhill et al., 2011; Xiao et al., 2011). AMPK is activated
upon various stress conditions including glucose deprivation, oxidative stress, ischemia
and hypoxia, and is a key player involved in the cellular response to exercise (Steinberg
and Kemp, 2009). Upon activation, AMPK induces multiple responses including
enhanced glucose metabolism and fatty acid oxidation to switch cellular metabolism from
anabolism to catabolism by: (a) increasing glucose uptake by stimulating glucose
transporters (GLUT) expression (Zheng et al., 2001) and its translocation to the plasma
membrane (Barnes et al., 2002; Kurth-Kraczek et al., 1999); (b) enhancing glycolysis by
activating 6-phosphofructo-2-kinase (PFK2) (Marsin et al., 2002); and (c) simultaneously
inhibiting fatty acid synthesis and enhancing β-oxidation by blocking acetyl-CoA
22
carboxylase (ACC1/2) activity (Hardie and Pan, 2002). AMPK is regarded a key
regulator of pathways implicated in aging and a potential longevity regulator in worms
(Greer et al., 2007) and is also involved in the beneficial effects of CR (Kahn et al.,
2005). Mixed results were reported regarding change of AMPK activity during aging in
different tissues, but growing evidence suggests that decreased AMPK activity and/or
decreased responsiveness to AMPK activity is associated with declined mitochondrial
function as a function of age (Finley and Haigis, 2009), thus indicating an inter-
relationship between mitochondrial energy status and AMPK activity.
It may be surmised that the cross-talk between IIS, JNK, and AMPK signaling in the
brain, their modulation by mitochondrial signaling molecules, and how these signaling
impinge on mitochondrial function is of importance to understand the process of aging
and their relevance to some neurodegenerative disorders. Since active JNK can also
initiate mitochondrion-dependent apoptosis, impairment of the communication between
mitochondrion-supported redox signaling and cytosolic signaling pathways may be the
basis for the mechanisms inherent in cell death pathways and the loss of cell function
associated with aging and age-related degenerative disorders.
23
1.4. SIGNIFICANCE
Aging is a major risk factor for most neurodegenerative diseases. Cellular and
molecular changes that occur during normal ageing render the brain vulnerable to
degeneration. Alterations in mitochondrial energy metabolism and redox homeostasis are
hallmarks of brain aging, and impinge on all aspects of cell function and are involved in
the development of age-related pathologies. Studies presented here have several novel
features that should contribute to the elucidation of the processes involved in the
metabolic network that controls cellular energy levels, redox environment and cell
signaling during aging and neurodegeneration.
(a) These studies provide new insights in the aging process beyond the accumulation
of oxidative damage as purported by the ‘mitochondrial theory of aging’, which has been
challenged by several studies (Jang and Remmen, 2009). Actually, work presented here
emphasizes the bioenergetic component in intimate relation with the mitochondrial redox
component; hence, it helps to identify the molecular basis for the connection of
metabolism (energy and redox) and lifespan.
(b) The pathways examined in these studies are part of an intricate signaling network
that has evolved around cellular energetics, mitochondrial metabolism, and generation of
H
2
O
2
, further supporting the links between the mitochondrial formation of signaling
molecules, the rate of aging, and the course of age-related diseases. The characterization
of the consequences of impaired mitochondrial energy-redox status on the overall cell
24
function will contribute to the understanding of the basic regulation of cell growth,
survival, and death in aging.
(c) The understanding of the function of NNT in the collective impairment of the
mitochondrial energy-redox axis during aging may provide further information and
potential therapeutic targets of the mitochondrial pathophysiology inherent in brain aging
and age related neurodegenerative disorders. Characterization of the signaling events
originating in the mitochondria and integrating on mitochondria might unravel the
molecular links between strategies aimed at restoring the mitochondrial energy-redox axis
and the aging process.
(d) This work helps to elucidate the mechanistic basis for the beneficial effects of CR
especially a more feasible short-term CR against aging, and will facilitate and guide the
application of CR or CR mimetics onto human subjects.
25
1.5. HYPOTHESIS AND SPECIFIC AIMS
Based on the information described in the previous sections, we hypothesize that NNT
is a critical component of the mitochondrial energy-redox axis that is compromised
during aging, and perturbation of the axis elicits not only mitochondrial dysfunction but
also cellular disorders through redox-sensitive signaling.
In the current study, we set up three specific aims, each of which serves to accomplish
a specific goal as shown in Fig.I-4.
Specific Aim 1: Characterize functional changes of the mitochondrial energy-redox
axis as a function of age and its modulation by caloric restriction
Specific Aim 2: Determine the overall role of NNT in regulating mitochondrial
function and cytosolic signaling pathways modulated by metabolic or oxidative signals.
Specific Aim 3: Mechanistically investigate the tissue-specific function of NNT in
linking the mitochondrial energy status to the redox environment.
26
Figure 1-4. Overview of specific aims 1-3. Focused on the NNT-linked mitochondrial
energy-redox axis, each specific aim addresses a specific question. Specific aim 1
investigates the age-dependent changes of the mitochondrial energy-redox axis and its
modulation by short-term CR; Specific aim 2 focuses on NNT regulation of mitochondrial
and cellular function and in a cell model; Specific aim III aims to mechanistically
investigate the underlying mechanism of NNT in linking the mitochondrial energy status to
the redox environment.
27
CHAPTER 2
SHORT-TERM CALORIC RESTRICTION ATTENUATES
AGE-DEPENDENT IMPAIRMENT OF
THE MITOCHONDRIAL ENERGY-REDOX AXIS IN BRAIN
2.1. ABSTRACT
Earlier studies have suggested that mitochondrial dysfunction is associated with
brain aging. Short-term caloric restriction (CR) has shown some beneficial effects
that are similar to life-long CR in tissues such as liver and skeletal muscle. In this
study, we characterized the functional changes of the mitochondrial energy-redox
axis in brain, entailing components involved in mitochondrial bioenergetics and
redox status maintenance, in the aging process and their modulation by short-term
CR. Our data show that 2-month of 40% CR leads to distinct responses for Fischer
344 rats at different ages regarding brain mitochondrial function. Mitochondrial
respiration and activities of enzymes that control the entry of substrates to TCA cycle,
including pyruvate dehydrogenase (PDH) and succinyl CoA transferase (SCOT), are
significantly higher after short-term CR for rats of 26-month, but not for rats of 6-
28
month. Increased activity of SCOT in brain is also associated with higher β-
hydroxybutyrate levels in the peripheral serum, suggesting the increased demand of
alternative energy source during short-term CR. On the other hand, mitochondrial
thioredoxin-2 (Trx2) system shows an age-dependent decrease of activities; but after
short-term CR, the activities or expression of redox enzymes, such as peroxiredoxin-
3 (Prx3) and Trx2 are upregulated in old animals. Short-term CR significantly
attenuated age-dependent increase in H
2
O
2
generation and the amount of
glutathionylated protein in mitochondria for animals at 26-month, but these were not
observed in 6-month old animals. Short-term CR also rescues the age-dependent
decline of nicotinamide nucleotide transhydrogenase (NNT) activity, in association
with a higher NADPH/NADP
+
ratio in brain homogenate. In summary, these data
show that declined mitochondrial energy metabolism and impaired mitochondrial
redox homeostasis by aging are functionally inter-dependent, and can be attenuated
by 2-month short-term CR in old rats, but not in young rats.
29
2.2. INTRODUCTION
Brain, as the center for thought, emotion, and memory, is affected by aging as other
organs, in terms of perturbed energy homeostasis, accumulation of damaged proteins and
lipids and lesions in their nucleic acids (Mattson and Magnus, 2006). Brain aging is a risk
factor for age-associated diseases such as neurodegenerative disorders. These diseases
may occur when neuronal cells fail to respond adaptively to age-related decline in basal
metabolic rates and physiological performances in energy requiring tasks such as
neuromuscular coordination, cognitive performance, and environmental awareness
(Swerdlow, 2007), which renders the brain particularly susceptible to an energy crisis.
Also, cellular composition of brain consists mainly of terminally differentiated neurons.
Hence the regenerative capacity of the brain in forms of cell regeneration is relatively
reduced as compared to other organs, which makes the brain more vulnerable to
oxidation-induced cell damage and cell death (Andersen, 2004).
In the central nervous system, mitochondria meet the energy demands of the cell that
support metabolic, osmotic, and mechanical functions; they are sources of H
2
O
2
, and play
a pivotal role as mediators of the intrinsic apoptotic pathway. Accordingly, the role of
mitochondria in aging is centered on two key events, (a) decrease in mitochondrial ability
to generate ATP through OXPHOS and (b) accumulation of oxidative damage (Navarro
and Boveris, 2007b). Moreover, mitochondria integrate distinct cytosolic signaling
pathways and generate second messengers, such as H
2
O
2
, that are implicated in the
30
modulation of redox-sensitive signaling pathways, thus transducing redox signals into a
wide variety of responses, such as proliferation, differentiation, and cellular death
pathways (Moran et al., 2001; Yin et al., 2012a). Hence, oxidants such as H
2
O
2
have a
dual function: on the one hand, H
2
O
2
is involved in the fine tuning of signaling and
transcription through modulation of redox-sensitive pathways and, on the other hand,
higher levels of H
2
O
2
–as expected with a diminished energy-conservation capacity of
mitochondria– are involved in oxidative damage to cell constituents (including proteins,
nucleic acids, and lipids), a well-documented phenomenon under the term of oxidative
stress.
Mitochondrial redox state is a critical mediator of metabolic-, signaling-, and cell
death-related processes by shuffling between oxidized and reduced states (Dalle-Donne et
al., 2008). The mitochondrial redox status cannot be viewed independent of its energy-
transducing capacity but is integrated in a mitochondrial energy-redox axis: the energy
component of this axis is encompassed by the generation of reducing equivalents (NADH
and FADH
2
) by the TCA and their flow through the respiratory chain with concomitant
generation of O
2
.–
and H
2
O
2
. The redox component is the domain of H
2
O
2
removal
systems –mainly glutathione peroxidases (GPx1 and GPx4) and peroxiredoxin-3– that
use GSH and thioredoxin-2 as electron donors, respectively. The ultimate reductant of
these systems is NADPH (supporting the activities of glutathione reductase and
thioredoxin reductase). Mitochondrial NADPH is mainly formed through three pathways:
NADP
+
-dependent IDH2, malic enzyme, and nicotinamide nucleotide transhydrogenase
31
(NNT). NNT, a nuclear encoded mitochondrial 110 kDa protein located on the inner
mitochondrial membrane (Hoek and Rydström, 1988 ), catalyzes the reduction of NADP
+
to NADPH using the membrane proton gradient as driving force and NADH as electron
donor, and accounts for more than 50% of the mitochondrial NADPH pool (Rydstrom,
2006). Hence, mitochondrial function and their involvement in domain-specific signaling
via second messengers are largely determined by maintenance of the mitochondrial
energy-redox axis entailing the energy metabolism, NNT, and redox homeostasis.
CR extends the mean and maximal life spans of many organisms such as yeast,
worms, rodents, and monkeys by up to 50% (Koubova and Guarente, 2003; Lane, 2000;
Weindruch and Walford, 1988), and delays the progression of multiple age-related
diseases such as diabetes, cancers, and cardiovascular disease (Colman et al., 2009;
Speakman and Mitchell, 2011). In the brain, CR delays age-related functional deficits and
may reduce the risk of neurodegenerative diseases including Alzheimer’s disease,
Parkinson’s disease, and Huntington disease (Maalouf et al., 2009; Mattson, 2000). CR
also suppresses changes in a variety of genes encoding mitochondrial proteins and
involved in energy metabolism and oxidative stress (Antico Arciuch et al., 2009; Lee et
al., 2000). It has been shown that CR induces mitochondrial biogenesis, improves
metabolic function, and reduces oxidative stress in rodent brain (Mattson et al., 2002;
Speakman and Mitchell, 2011); hence, it seems that mitochondria are in the central stage
of the mechanisms for the CR-induced beneficial effects (Guarente, 2008). The length of
time on a CR diet required for the exhibition of its effects is not well characterized,
32
especially in brain. Recently, it was shown that short-term caloric CR elicits beneficial
effects that are similar to life-long CR in tissues such as liver, heart, and skeletal muscle
(Cao et al., 2001; Judge et al., 2004; Mitchell et al., 2010; Rohrbach et al., 2006 ).
In this study, we characterized the functional changes of the mitochondrial energy-
redox axis in brain, including components involved in mitochondrial bioenergetics and in
control of redox homeostasis in the aging process and their modulation by short-term CR.
2.3. MATERIALS AND METHODS
2.3.1. Short-term CR regiment
Fisher 344 rats purchased from the National Institute of Aging (NIA) (age 4, 12 and 24
months) were allowed to acclimatize and started on a short term CR restriction regiment.
The short-term caloric restriction was initiated at 15% for 1 week and increased to 25%
for another week and then maintained at 40% for another 8 weeks. Ad libitum (AL) fed
rats were kept on NIH-31 diet and CR rats were on a NIH31/NIA fortified diet with 60%
calories of control diet but increased amount of nutrients so that the AL and CR group
would receive the same amount of vitamins and other nutrients to ensure normal body
functions. Their body weight was monitored and measured weekly, as shown in Fig. 2-1.
33
For young and middle age rats, 2-month CR reverse the growth of body weight, and for
old rats at 26-month, the decline of body weight was accelerated.
2.3.2. Isolation of brain mitochondria
Brain mitochondrial were isolated based on methodology previously described
(Anderson and Sims, 2000). Brain tissue are excised, chopped into fine pieces, washed
with and homogenized in isolation buffer containing 210 mM mannitol, 70 mM sucrose
and 2 mM HEPES, pH 7.4, plus 0.05% (w/v) BSA. The homogenate was centrifuged at
800 g for 8 min, the pellet was removed, and the centrifugation process was repeated. The
Figure 2-1 Body weight of rats during short-term CR. Fischer 344 rats at 4-, 12m- and
24-month were fed with either AL or CR diet for 9 weeks, and their body weights were
monitored weekly till the day of sacrifice. Data shown were the average body weight of 3
animals per diet per age group. A. Weekly body weight of rats fed with AL diet at different
age; B. Weekly body weight of rats fed CR diet at different age.
350
400
450
500
550
1 2 3 4 5 6 7 8 9
Body weight (g)
Week
4m-6m
12m-14m
24m-26m
250
300
350
400
450
500
1 2 3 4 5 6 7 8 9
Body weight (g)
Week
4m-6m
12m-14m
24m-26m
A B
34
supernatant was centrifuged at 8000 g for 10 min, the pellet was washed with the
isolation buffer, and the centrifugation was repeated. The pellet containing a mixture of
organelles was further fractionated by centrifugation at 8500 g for 10 min in a Percoll
gradient (consisting of three layers of 18, 30 and 60% (w/v) Percoll in sucrose/Tris buffer
(0.25 M sucrose, 1 mM EDTA and 50 mM Tris/HCl), pH 7.4). Mitochondria were
collected from the interface of 30% and 60% Percoll and washed with the sucrose/Tris
buffer. Mitochondrial protein concentration was determined using protein assay reagent
(Biorad).
2.3.3. Western blotting
Cell or mitochondria lysate was solubilized in SDS sample buffer, separated by
SDS/PAGE, and transferred onto PVDF membranes. Using appropriate antibodies, the
immunoreactive bands were visualized with an enhanced chemiluminescence reagent.
2.3.4. Pyridine dinucleotides levels
NAD
+
, NADH, NADP
+
, and NADPH levels were measured by High-performance
liquid chromatography (HPLC) (Klaidman et al., 1995). Briefly, isolated mitochondria or
cell pellet were homogenized in buffer (0.06 M KOH, 0.2 M KCN, and 1 mM
bathophenanthroline disulfonic acid) followed by chloroform extraction. Chloroform
extraction was carried out by centrifugation at 14,000 rpm in a microcentrifuge at 4 °C;
the resulting aqueous supernatant with soluble pyridine nucleotides was collected and
35
extracted thrice to remove lipids and proteins. Finally, it was filtered with a 0.45-μm
positively charged filter (Pall Life Sciences) to remove RNA and DNA in
microcentrifuge at 4 °C. The mobile phase consisted of 0.2 M ammonium acetate (buffer
A) at pH 5.5 and HPLC grade methanol (buffer B). A gradient program with initial
conditions as 100% buffer A and 0% buffer B was set. From 0 to 4 min, we used 0 to 3%
B, and from 4 to 23 min, we used 3 to 6.8% B, followed by washing the column with 50%
A and 50% B, and re-equilibrated to initial conditions for next run. Quantitation of
pyridine nucleotides was performed by integrating the peaks and adding the cyanide
adducts as detected by the fluorescence spectrophotometer (λ
exc
= 330nm; λ
em
= 460 nm).
2.3.5. Mitochondrial respiration measurement
MitoXpress™ Fluorescent method – Mitochondrial respiration was measured using
MitoXpress™ (Luxcel Biosciences Ltd, Cork, Ireland) fluorescent dye from isolated
mitochondria following a previously established protocol (Will et al., 2006). Briefly, 50
μg of isolated mitochondria were diluted to 1 μg/μL with respiratory buffer (250 mM
sucrose, 15 mM KCl, 1 mM EGTA (ethylene glycol tetraacetic acid), 5 mM MgCl
2
, 30
mM KH
2
PO
4
, pH 7.4) and added to test well. MitoXpress™ probe was reconstituted into
1 μM stock solution and further diluted 1:10 in respiration buffer. State 4 respiration was
stimulated with the addition of glutamate (5 mM) and malate (5 mM) as substrates. State
3 respiration was stimulated by the addition of glutamate (5 mM) and malate (5 mM) plus
ADP (410 μM). The rate of oxygen consumption was calculated based on the slope of the
36
response of isolated mitochondria to the successive administration of substrates. 100 μL
mineral oil was added to each well promptly after the addition of MitoXpress working
solution into the well. MitoXpress signal was measured at 1-minute intervals for 60
minutes using excitation and emission wavelength of 380 nm and 650 nm respectively.
To determine the rate of respiration, fluorescence-time profiles were linearized using the
following coordinate scale: abscissa, Y: I(t
0
)/(I(t)-I(t
0
)), where I(t
0
) and I(t) represent
fluorescence intensity signals at the start and at time (t) of monitoring, respectively;
ordinate, X: 1/t, min
-1
; exclude 0 time points and regions of signal saturation, i.e., long
monitoring times. Linear regression analysis was applied to the transformed profiles to
determine the slope and correlation coefficient for each of the transformed profiles. State
4 and State 3 respiration rate were calculated as the reciprocal ratio of the above 2
calculated slopes, respectively. The RCR was defined by dividing the rate of oxygen
consumption/min for State 3 (presence of ADP) by the rate of oxygen consumption/min
for State 4 respiration.
Clark-type electrode method – Aliquots of mitochondria (100 μg/mL) were used in
measurements of respiratory activity using a Clark-type oxygen electrode (Hansatech
Oxygraph) as previously described (Boveris and Cadenas, 1975). Oxygen electrode
buffer (130 mM KCl, 2 mM KH
2
PO
4
, 3 mM HEPES, 2 mM MgCl
2
, 1 mM EGTA) was
incubated for 1 min in a magnetically stirred chamber at 30 °C. The respiratory substrates,
glutamate (5 mM) and malate (5 mM) were added followed by the isolated mitochondria
(100 μg). State 4 respiration was first measured in the absence of ADP. Subsequently,
37
State 3 respiration was measured in the presence of ADP (410 μM) to determine the
maximal rate of coupled ATP synthesis. Respiratory control ratio is calculated as the
State 3 to State 4 ratio.
2.3.6. Measurement of SCOT activity
SCOT activity was measured as previously described (Williamson et al., 1971). Briefly,
mitochondria lysates were disrupted by sonication and then centrifuged at 10,000 g for 20
min. The supernatants were incubated with 50 mM Tris-HCl, 5 mM MgCl
2
, 4 mM
iodoacetamide, 0.2 mM acetoacetate, and 0.1 mM succinyl-CoA, pH 8.0. Catalytic
activity was measured spectrophotometrically at 313 nm.
2.3.7. Serum ketone body assay
Serum ketone body level was measured using the Liquid Color β-hydroxybutyrate
assay kit (Stanbio, Boerne, Tx) following the manufacturer's instruction.
2.3.8. H
2
O
2
release
H
2
O
2
generation from freshly isolated mitochondria was determined by the Amplex Red
Hydrogen Peroxide/Peroxidase Assay kit (Invitrogen) following the manufacturer’s
instructions, with the presence of 5mM of glutamate, 5mM of malate and no ADP.
38
2.3.9. Redox components activity assays
Trx-2 activity – Thioredoxin activity of isolated mitochondria was measured at 30°C.
The assay mixture contains 2 mM EDTA, 400 μM NADPH, 7 μg/ml thioredoxin
reductase from E. coli, and 83 mM insulin as substrate in 100mM K
3
PO
4
buffer, pH 7.0.
The consumption of NADPH was followed at 340 nm, and a value of 6200 was used for
the molar absorption coefficient of NADPH.
TrxR2 activity – Thioredoxin reductase activity of isolated mitochondria was measured
at 30°C. The assay mixture contains 50 mM KCl, 10mM EDTA, 240 μM NADPH, 0.2
mg/ml BSA, and 2.5 mM of DTNB containing a disulfide bond as substrate in 50mM-
K
3
PO
4
buffer, pH7.0. The reduction of DTNB by TrxR was followed at 412 nm, and a
value of 25200 was used for the molar absorption coefficient of reduced DTNB.
2.3.10. NNT activity
NNT activity from NADPH to AcPyAD
+
(3-acetylpyridine adenine dinucleotide, an
NAD
+
analogue) was assayed in the absence of an energy source at 37°C as
described(Chen and Guillory, 1979). The assay mixture contained 200 μM NADPH, 300
μM-AcPyAD
+
and 2 mM of NaN
3
in 50mM-K
3
PO
4
buffer, pH 7.0. The reduction of
AcPyAD
+
by NADPH will be followed at 375 nm, and a value of 5100 will be used for
the molar absorption coefficient of reduced 3-acetylpyridine adenine dinucleotide.
39
2.3.11. Statistical analyses of experimental data
The general analytic approach utilized a comparison of means across experimental
conditions using analysis of variance (ANOVA) procedures (which reduce to a t-test in
the case of 2 experimental conditions). In most cases, the comparison used a one-factor
ANOVA (factor = experimental condition). For the experiments involving CR feedings
evaluated among 3 age groups, a two-factor (feeding by age group) ANOVA was used.
For each outcome variable, the following will be tested: (a) are there mean differences
between the AL and CR conditions, (b) are there mean differences across the three age
groups, and (3) do the age effects differ in animals fed AL or CR diet (test of interaction).
2.4. RESULTS
2.4.1. Aging and short-term CR alter the entry of substrates into TCA cycle in brain
Previous studies have demonstrated that activities of key enzymes that regulate energy
metabolism in mitochondria such as PDH and SCOT decrease as a function of age due to
post-translational modifications, which may partially account for the compromised
mitochondrial function in aging (Lam et al., 2009; Zhou et al., 2009). Mitochondria
isolated from brain of AL and CR rats at 6-, 14-, and 26-months, show decreased PDH
expression as a function of age; this effect was reversed by short-term CR at 26-month
40
(Fig. 2-2A). The age-dependent increase in PDH phosphorylation (and its inhibition) was
also alleviated by short-term CR at 26-month (Fig. 2-2B). SCOT is the key mitochondrial
matrix enzyme involved in the metabolism of ketone bodies, which are the only
alternative energy substrates for brain during glucose deficiency. SCOT levels are
upregulated by short-term CR for rats at 26-month (Fig. 2-2C), and increased activity of
SCOT in 26-month CR group was also associated with higher β-hydroxybutyrate levels
in the peripheral serum (Fig. 2-2D), suggesting the increased demand of alternative
energy source after short-term CR. However, these changes induced by short-term CR in
26-month rats were not observed in 6- and 14-month-old rats.
41
Figure 2-2 Aging and short-term CR alter the regulation of substrate entry to TCA
cycle. A. Whole brain mitochondria of rats of 6-, 14, and 26-month after 2-month of either
AL or CR diet feedings were isolated and the levels of phosphorylated form and total
expression of PDH-E1α were determined by Western blot, and VDAC was the loading
control; B. Levels of phospho- PDH-E1αto PDH-E1α were quantified and the relative ratio
of p-PDH/PDH was calculated and then normalized to the 6-month AL group, p<0.05 in AL
group with age; C. SCOT activity in isolated mitochondria from AL and CR rats at 26-month
was determined; D. Concentrations of β-hydroxybutyrate in peripheral serum of rats at above
mentioned settings were measured, p<0.05 in AL group with age. Bars represent mean values
± SEM of 4-5 animals per diet per group (* p < 0.05).
PDH E1α
VDAC
0
0.25
0.5
Concentration (mM)
Age (months)
14 26 6
AL CR AL CR AL CR
Age (months)
14 26 6
AL CR AL CR AL CR
p-PDH
A
D
0
50
100
150
200
250
Relative Intensity
Age (months)
14 26 6
AL CR AL CR AL CR
B
0
50
100
Activity (nmol min-1 mg-1)
*
26m-CR 26m-AL
C
42
2.4.2. Short-term CR rescued age-dependent decline of mitochondrial respiration
Brain represents 2% of the body weight, but accounts for 20% of total O
2
consumption
(Moreira et al., 2007). Correspondingly, brain relies heavily on mitochondrial respiration
for energy production. Mitochondrial respiration is an overall parameter that reflects
mitochondria function. Mitochondrial respiration decreases in aging in term of marked
decline in state 3 respiration (the active state, i.e., the respiratory activity that sustains
ATP formation) and the related RCR, as well as an increase in state 4 respiration (the
inactive state, with ADP depleted), all of which indicate a lower energy-transducing
efficiency. Therefore, mitochondrial respiration of AL and CR rat brain was measured
using Complex I-driven substrates malate/ glutamate. As shown in Fig. 2-3, the age-
associated decrease of state 3 respiration was not observed in CR rats (Fig. 2-3A); and the
age-associated increase of state 4 respiration was also attenuated in CR rats (Fig. 2-3B).
The changes of state 3 and state 4 respiration led to a substantial decreased of RCR
during aging, but was attenuated by 2-month CR in 26-month old rats using the florescent
dye method (Fig. 2-3C) and the classic Clark-type electrode (Fig. 2-3D). The
enhancement of mitochondrial respiratory capacity by short-term CR was not observed in
young animals (6-month), and conversely, there was a decrease of the RCR value in these
rats after CR.
43
Figure 2-3 Mitochondrial respiration in AL and CR rat brain of different age groups.
Oxygen consumption was measured in freshly isolated brain mitochondria were measured at
State 3 and State 4, and the respiratory control ratio (RCR, State 3 / State 4) was calculated
and normalized to 6-month AL group. State 4 respiration was measured in the presence of 5
mM L-malate and 5 mM L-glutamate; State 3 respiration was measured with 5 mM L-malate,
5 mM L-glutamate and 410 μM ADP. A. State 3 respiration of mitochondria from AL and
CR rat at different ages was measured by using the Luxcel MitoXpress (Luxcel Biosciences
Ltd, Cork, Ireland) Oxygen-sensitive Fluorescent assay; B. State 3 respiration was measured
by the same MitoXpress method; C. RCR was calculated based on State 3 and 4 data in A
and B, p<0.05 in AL group with age; D. Mitochondrial State 3 and 4 respiration was
measured by the Clark-type electrode and RCR was calculated using the same settings of
animals as above, p<0.05 in AL group with age. Bars represent mean values ± SEM of 4-5
animals per diet per group (* p < 0.05).
0
50
100
150
% of 6m
AL
CR
0
50
100
150
200
250
% of 6m
AL
CR
0
50
100
RCR (% of 6m-AL)
Age (months)
14 26 6
AL CR AL CR AL CR
Age (months)
14 26 6
Age (months)
14 26 6
Age (months)
14 26 6
AL CR AL CR AL CR
A
D C
B
0
40
80
120
RCR (% of 6m-AL)
44
2.4.3. Mitochondrial H
2
O
2
generation and redox status in aging and short-term CR
Steady-state levels of mitochondrial H
2
O
2
are regulated by both energy metabolism
(electron leak associated with oxidative phosphorylation) and the redox systems
(pathways associated with H
2
O
2
reduction), and thus reports the condition of the energy-
redox axis in the mitochondria. H
2
O
2
generation rate increased with age in isolated brain
mitochondria, and was reduced after CR in 26-month group; again, this effect by CR was
not seen in 6-month group, and there was an increase in H
2
O
2
generation rate in these
young animals (Fig. 2-4A). Protein thiols are sensitive to changes in the redox
environment: the reversible formation of mixed disulfides entails a reaction between
protein cysteine sulfhydryls and GSH in a process termed S-glutathionylation. S-
glutathionylation reflects the redox status of the mitochondria (Schafer and Buettner,
2001) and is viewed as a regulatory device for proteins involved in energy metabolism,
redox signaling, and cell function (Dalle-Donne et al., 2008). Our data show that protein
glutathionylation levels increased with age in mitochondria (Fig. 2-4B), which is
consistent with the decreased GSH/GSSG ratio with age in brain mitochondria (Rebrin et
al., 2003). Short-term CR significantly reduced the amount of glutathionylated protein in
mitochondria for animals at 26-month, suggesting that a more reducing environment is
induced by short-term CR at that age. However, the decrease in S-glutathionylation was
not observed in young animals.
45
2.4.4. Modulation of mitochondrial Trx system by aging and short-term CR
Maintenance of mitochondrial H
2
O
2
homeostasis is the domain of glutathione
peroxidases and peroxiredoxins: the latter belong to a family of thiol peroxidases
involved in peroxide reduction. Mitochondrial Prx3 is the target of up to 90% of H
2
O
2
generated in the mitochondrial matrix with a high reaction rate (2×10
7
M
-1
·s
−1
) especially
at low levels of H
2
O
2
(Cox et al., 2010; Rhee et al., 2005). The reducing power for Prx is
transmitted through thiols of the Trx system: NADPH → TrxR → Trx → Prx
(Hofmann
et al., 2002). To characterize the effect of aging and short-term CR on the capacity of
Figure 2-4. Mitochondrial H
2
O
2
generation and S-glutathionylation in aging and CR. A.
H
2
O
2
release from freshly isolated brain mitochondria was monitored for 30 min using
Amplex Red fluorescence dye in the presence of 5 mM L-malate and 5 mM L-glutamate, and
the release rate was calculated. Bars represent mean values ± SEM of 4-5 animals per diet per
group, p<0.05 in AL group with age. (* p < 0.05). B. Isolated brain mitochondria were
immunoblotted with glutathione antibody for levels of glutathionylated proteins.
COX-IV
Glutathionylated
proteins
35 KD
55 KD
Age (months)
14 26 6
AL CR AL CR AL CR
100
200
300
H
2
O
2
generation
(pmol min
-1
mg
-1
)
Age (months)
14 26 6
AL CR AL CR AL CR
A
B
46
removing H
2
O
2
by brain mitochondria, the expression levels of Prx3 and Trx2, as well as
their enzymatic activities were measured in mitochondria isolated from AL and CR rat
brain of different ages. As shown in Fig. 2-5A, Trx2 levels decreased with age and were
upregulated after 2-month CR for 26-month group, which was consistent with the trend
of its activity (Fig. 2-5B), but no significant differences in the expression and its activity
were found between the AL and CR groups at the age of 6- or 14-months. The activity of
TrxR2 also decreased with age but short-term CR did not enhance its activity (Fig. 2-5C).
Conversely, the expression of Prx3 in brain mitochondria (Fig. 2-5A) increased with age
and was further upregulated by CR as quantified in Fig. 2-5D. The discrepancy of the
alteration of the components of the redox systems indicates that the source of reducing
equivalents might account more for the increased [H
2
O
2
]
ss
and oxidized redox
environment in mitochondria that occurs with aging.
47
Figure 2-5 Assessment of the mitochondrial Trx2-Prx3 system in aging and CR. A.
Isolated brain mitochondria were immunoblotted with Trx2 and Prx3 antibody for their
expression levels, and VDAC is the loading control; B. Trx2 activity was measured in
isolated mitochondria of all the groups, p<0.05 in AL group with age; C. TrxR2 activity was
measured in isolated mitochondria of all the groups, p<0.05 in AL group with age; D. Prx3
expression levels examined in A was quantified and normalized to its levels in 6-month AL
group, p<0.05 in AL and CR group with age.. Bars represent mean values ± SEM of 4-5
animals per diet per group (* p < 0.05).
0
10
20
30
40
Activity (nmol min-1 mg-1)
Age (months)
14 26 6
AL CR AL CR AL CR
Trx-2
VDAC
Age (months)
14 26 6
AL CR AL CR AL CR
Prx-3
0
300
600
Relative expression to 6m-AL
Age (months)
14 26 6
AL CR AL CR AL CR
A
D C
B
Age (months)
14 26 6
AL CR AL CR AL CR
0
250
500
Activity (nmol min-1 mg-1)
48
2.4.5. NADPH availability and NNT activity in aging and short-term CR
NADPH is the ultimate electron donor for mitochondrial redox systems including Trx2
and GSH systems. Decreased NADPH supply could account for impaired mitochondrial
redox homeostasis in aging. NADPH and NADP
+
levels in brain homogenate of AL and
CR rats were measured, and the NADPH/ NADP
+
ratio decreased with age, and was
increased by short-term CR, but only at 26-month (Fig. 2-6A). As the primary NADPH
generator in mitochondria, the loss of function of NNT with respect to aging also
occurred in aging (Fig. 2-6B). This finding indicates that NNT could be highly involved
in the age-related mitochondria dysfunction. It is also shown that short-term CR resulted
in increased NNT activity in old rats (Fig. 2-6B), indicating that the modulation of NNT
activity could be one of the effects of CR on regulating mitochondrial function.
49
2.5. DISCUSSION
2.5.1. Effects of short-term CR on young and old animals
CR exhibits beneficial effects including not only life span extension, but also the delay
of progression of many pathophysiologies, especially in the central nervous system. In
studies in humans, CR improves biomarkers associated with longevity and decreases the
incidence of neurodegenerative diseases (Heilbronn et al., 2006; Luchsinger et al., 2002;
Redman et al., 2008). However, the adherence of human subjects to long-term CR could
Figure 2-6. Modulation of NNT activity and pyridine dinucleotide levels by aging and
CR. A. Fresh brain mitochondria were measured for NNT activity by absorbance change at
375 nm emitted from 3-acetylpyridine adenine dinucleotide (AcPyAD
+
), a NAD
+
analog,
p<0.05 in AL group with age; B. NADPH and NADP
+
levels were determined by HPLC
method and the ratio of NADPH and NADP
+
was calculated, p<0.05 in AL group with age.
Bars represent mean values ± SEM of 4-5 animals per diet per group (* p < 0.05).
0
1
2
NADPH/NADP Ratio
Age (months)
14 26 6
AL CR AL CR AL CR
Age (months)
14 26 6
AL CR AL CR AL CR
*
0
5
10
15
Activity (nmol min-1 mg-1)
A
B
50
be a potential obstacle for its application to clinical settings. It was shown that in liver,
short-term CR substantially shifted the genomic profile of old mice toward the profile
associated with long-term CR, and these genomic changes occur as early as two weeks
after initiation of CR (Cao et al., 2001). And 2-4 weeks of 30% CR improved survival
rate and kidney function following renal ischemia reperfusion injury in mice, correlated
with increased insulin sensitivity and increased expression of markers of antioxidant
defense (Mitchell et al., 2010). It was also suggested that increased mitochondrial
function in CR is due to induction of mitochondrial biogenesis and recycling of damaged
mitochondria (Guarente, 2008). Therefore, we would like to test the effect of a short-term
CR (2-month) on declined brain mitochondrial function during aging. Our results
suggested that 2-month of 40% CR leads to distinct responses for Fischer 344 rats at
different ages regarding brain mitochondrial function. Most of the improvement of
mitochondrial energy metabolism and redox homeostasis were observed only in the 26-
month old rats, but not in the 6- or 14-month-old ones, and there was even some adverse
effects for young animals after short-term CR, including: (a) PDH inhibition (Fig. 2-2B);
(b) blood levels of ketone bodies (Fig. 2-2D); (c) mitochondrial respiration efficiency
(Fig. 2-3); (d) mitochondrial H
2
O
2
generation (Fig. 2-4A); (e) protein oxidative
modification (Fig. 2-4B); (f) mitochondrial Trx2 activity (Fig. 2-5); and (g) NNT activity
and NADPH availability (Fig. 2-6). All of these indicate that 2-month of 40% CR is
enough to induce functional changes of mitochondria of old rats, but not in young rats.
Whether or not a longer time of CR is required for young animals to exhibit beneficial
51
effects is still to be determined. Because the standard protocol for long-term CR in
rodents starts at the same age as the young group of this study (4-month) and throughout
their life, it is possible that at a later time point the positive effects of CR will be
observed on young animals, although the exact point of transition remains to be explored.
2.5.2. Substrate availability and utilization during aging and CR
Glucose is the primary fuel for brain, whereas metabolism of ketone bodies represents
an alternative fuel source during glucose deprivation (Grinblat et al., 1986). It was shown
that activities of enzymes that control the entry of substrates to TCA cycle, such as PDH
and SCOT, which are key enzymes in pyruvate and ketone body metabolism, respectively,
decrease with age in brain. Age-dependent increase of blood levels of β-hydroxybutyrate
indicates the demand for alternative energy source when glucose metabolism is impaired
during aging. Both PDH and SCOT activity were increased by short-term CR for 26-
month rats, and increased SCOT activity was accompanied by further increased of β-
hydroxybutyrate in this study. During CR, when energy uptake is limited and the energy
transduced from glucose is not sufficient, the cells tend to utilize alternative energy
sources such as ketone body (generated by lipid metabolism in liver) to fulfill the demand
for ATP. SCOT activity is also upregulated to enhance ketone body metabolism. Ketone
body metabolism has been shown as a temporary mechanism that prevents the further
decline of brain mitochondrial bioenergetic capacity associated with decreased activities
of PDH activity in a triple transgenic mouse model of Alzheimer’s disease (Yao et al.,
52
2010). 2-Deoxy-D-glucose (a CR mimetic) treatment induced ketogenesis in the same
mouse model and this resulted in increased ketone body metabolism in the brain and a
significant reduction of both amyloid precursor protein and amyloid-β (Yao et al., 2011).
2.5.3. Mitochondrial energy-redox axis and H
2
O
2
regulation
As the electron flows through the mitochondrial complexes, the electron leakage leads
to the generation of O
2
.–
, which is further converted to H
2
O
2
, either through enzymatic
catalysis by superoxide dismutase (SOD) or spontaneous dismutation. In this study,
electron transfer in mitochondria decreased in aged brain, and short-term CR on old rats
exhibited similar effects on mitochondrial respiration as seen in long-term CR mice
(Nisoli et al., 2005). The rate of mitochondrial generation of H
2
O
2
(Fig. 2-4A) reflects the
mitochondrial energy status (Fig. 2-3), for mitochondrial H
2
O
2
generation in state 4
respiration is about 4-5 times higher than that during effective OXPHOS (state 3
respiration) (Boveris et al., 1999). Mitochondrial steady state levels of H
2
O
2
are also
determined by the redox systems where it is reduced to H
2
O. Our data show that there
was an age-dependent decrease of activities of Trx2/TrxR2 system, and the activity of
Trx2 and Prx3 was upregulated upon CR (Fig. 2-5). The increased expression of Prx3
with age could be a compensatory effect at the transcription level, or the enhanced
immune-blotting signal may be a reflection of the accumulation of overoxidized (inactive)
form of Prx3, as seen in liver (Musicco et al., 2009). A further analysis of the activity of
Prx3 or the quantification of its active form is thus necessary. Given that overexpression
53
of Trx2 in some cell lines did not protect them against oxidative stress (Patenaude et al.,
2004), and decreased ratio of NADPH/NADP
+
was observed in aging (Fig. 2-6A) (Yarian
et al., 2006), the availability of reducing equivalents, i.e., alterations in NADPH/NADP
+
levels, may limit the electron flow through the redox systems, thus affecting the
efficiency of Prx3-catalyzed degradation of H
2
O
2
and the Grx2-catalyzed protein de-
glutathionylation, and hence contribute to the increased oxidative status observed in
aging (Fig. 2-4). All of these indicate the importance of the enzymes that produces
NADPH in mitochondria (primarily NNT, and IDH2), as well as the function of the
energy components, which provides reducing equivalents and driving force for these
enzymes (NADH and proton gradient for NNT; isocitrate for IDH2). This is supported by
the similar patterns of mitochondrial bioenergetic function and NNT activity that were
shown in aging and short-term CR (Fig. 2-2, 2-3, and 2-6). The disruption of electron
flow from fuel molecules to redox regulators due to decreased NNT activity in aging
could exaggerate the impairment of mitochondrial function through the redox-mediated
post-translational modifications of critical enzymes involved in the energy metabolism
(Yap et al., 2009). Thus, NNT makes the mitochondrial energy metabolism and redox
status an integrated regulatory device rather than independent events, and further affects
cellular function through the redox-sensitive signaling pathways (Yin et al., 2012b).
54
CHAPTER 3
SILENCING OF NICOTINAMIDE NUCLEOTIDE TRANSHYDROGENASE
IMPAIRS CELLULAR REDOX HOMEOSTASIS
AND ENERGY METABOLISM IN PC12 CELLS
1
3.1. ABSTRACT
Mitochondrial NADPH generation is largely dependent on the inner-membrane
Nicotinamide Nucleotide Transhydrogenase (NNT), which catalyzes the reduction of
NADP
+
to NADPH utilizing the proton gradient as the driving force and NADH as the
electron donor. Small interfering RNA (siRNA) silencing of NNT in PC12 cells results in
decreased cellular NADPH levels, altered redox status of the cell in terms of decreased
GSH/GSSG ratios and increased H
2
O
2
levels, thus leading to an increased redox potential
1
Yin, F., Sancheti, H., and Cadenas, E. (2012b). Silencing of nicotinamide nucleotide transhydrogenase
impairs cellular redox homeostasis and energy metabolism in PC12 cells. Biochim Biophys Acta 1817,
401-409.
55
(a more oxidized redox state). NNT knockdown results in a decrease of oxidative
phosphorylation while anaerobic glycolysis levels remain unchanged. Decreased
oxidative phosphorylation was associated with a) inhibition of mitochondrial pyruvate
dehydrogenase (PDH) and succinyl-CoA:3-oxoacid CoA transferase (SCOT) activity; b)
reduction of NADH availability, c) decline of mitochondrial membrane potential, and d)
decrease of ATP levels. Moreover, the alteration of redox status actually precedes the
impairment of mitochondrial bioenergetics. A possible mechanism could be that the
activation of the redox-sensitive c-Jun N-terminal kinase (JNK) and its translocation to
the mitochondrion leads to the inhibition of PDH (upon phosphorylation) and induction
of intrinsic apoptosis, resulting in decreased cell viability. This study supports the notion
that oxidized cellular redox state and decline in cellular bioenergetics –as a consequence
of NNT knockdown– cannot be viewed as independent events, but rather as an
interdependent relationship coordinated by the mitochondrial energy – redox axis.
Disruption of electron flux from fuel substrates to redox components due to NNT
suppression induces not only mitochondrial dysfunction but also cellular disorders
through redox-sensitive signaling.
56
3.2. INTRODUCTION
Mitochondria integrate distinct cytosolic signaling pathways and generate second
messengers (e.g., H
2
O
2
, NAD
+
/NADH, ATP) that are involved in the regulation of redox-
sensitive cell signaling. Mitochondrial generation of H
2
O
2
is a function of the
mitochondrial energy – redox axis: the energy component of this axis is encompassed by
the generation of reducing equivalents (NADH) by the TCA cycle and their flow through
the respiratory chain with concomitant generation of O
2
.–
and H
2
O
2
. The redox
component is the domain of H
2
O
2
removal systems –mainly glutathione peroxidase and
peroxiredoxin 3 – that use GSH and thioredoxin-2 as electron donors; the ultimate
reductant for these systems is NADPH (supporting the activities of glutathione reductase
and thioredoxin reductase). Hence, the steady-state levels of mitochondrion-generated
H
2
O
2
in cytosol are largely determined by maintenance of the mitochondrial energy –
redox axis.
Mitochondrial NADPH is formed mainly through three pathways (a) NADP
+
-
dependent IDH2, (b) malic enzyme, and (c) nicotinamide nucleotide transhydrogenase
(NNT). Of these pathways, ~50% of the mitochondrial NADPH pool is uncoupler
sensitive, thus suggesting that NNT-catalyzed reduction of NADP
+
accounts for more
than 50% of mitochondrial NADPH pool (Rydstrom, 2006). NNT –a nuclear encoded
mitochondrial 110 kDa protein located on mitochondrial inner membrane (Hoek and
Rydström, 1988 ) – catalyzes the reversible reduction of NADP
+
to NADPH and the
57
conversion of NADH to NAD
+
(equation 1). Under physiological conditions, the proton
gradient across the mitochondrial inner membrane strongly
NADH + NADP
+
+ H
+
intermembrane
↔ NAD
+
+ NADPH + H
+
matrix
[1]
stimulates the forward reaction, i.e., the generation of NADPH. Under anaerobic and
energy-deficient conditions, the reverse reaction catalyzed by NNT, i.e., the generation of
NADH, also has transient effects on maintaining mitochondrial membrane potential
through NADPH hydrolysis and H
+
pumping, but the contribution of the backward
reaction to the proton gradient is probably of little significance under physiological
conditions (Ying, 2008).
The production of NADPH by NNT requires NADH as electron donor and the proton
gradient as driving force. NNT could therefore provide a critical link between the
mitochondrial metabolic function and redox homeostasis by coupling NADPH generation
to the TCA cycle, active respiration, and O
2
.–
/H
2
O
2
production by electron transfer chain.
The crucial role of NNT in the maintenance of the redox environment is revealed by the
fact that ablation of NNT renders C. elegans more susceptible to oxidative stress and
decreases the cellular GSH/GSSG ratio (Arkblad et al., 2005). In mammalians, mice
deficient in SOD2 die much earlier if they also lack functional NNT (Huang et al., 2006).
Moreover, glucose causes a dramatic increase in oxidant levels in β-cells from mice
carrying loss-of-function mutants of NNT (Freeman et al., 2006).
Thus, NNT could play a significant role in the maintenance of the mitochondrial
energy–redox axis and determine the levels of mitochondrion-generated H
2
O
2
in cytosol
58
and its subsequent involvement in domain-specific regulation of redox-sensitive signaling
pathways: p38 MAPK, JNK, and other serine kinases are redox-sensitive, although the
extent to which H
2
O
2
from sources other than mitochondria contribute to their regulation
is not clear. The involvement of mitochondrial H
2
O
2
in the regulation of JNK, however,
has been established as well as the translocation of JNK to the outer mitochondrial
membrane in primary cortical neurons and its effects on energy metabolism by triggering
pathways that involve inhibition of pyruvate dehydrogenase (PDH) upon phosphorylation
of the E
1α
subunit (Zhou et al., 2008).
This study was aimed at assessing (a) the function of NNT in regulating redox status
of the cell; (b) the effect of NNT-regulated redox change on cellular energy metabolism,
and (c) at identifying and validating potential mitochondrial-cytosol signaling pathway(s)
that are modulated by metabolic or oxidative signals in NNT knockdown cells and that
impact the fate of the cell. These aims were performed in PC12 cells transfected with
either non-sense siRNA or siRNA against NNT.
3.3. MATERIALS AND METHODS
3.3.1. Cell culture – Experiments in this study were performed on rat pheochromocytoma
cells (PC12), a dopaminergic cell model with well-defined steps in response to metabolic,
59
oxidative stress-, or apoptotic signals. This cell line enables to investigate the effects of
NNT knockdown on cellular redox status, metabolic function and apoptosis, as well as its
implications in neurodegeneration. PC12 cells, obtained from American Type Culture
Collection, were maintained in RPMI medium 1640 supplemented with 10% horse serum,
5% fetal bovine serum, and 1% penicillin-streptomycin (Murphy, 2012). Cells
differentiation was done in differentiation medium (RPMI medium 1640 + 1% donor
horse serum + 100 ng/ml nerve growth factor + 50 ng/ml cyclic AMP) for 5 days.
3.3.2. siRNA transfection – The siRNA sequences against NNT were 5´-
ggcggaaacuuugaaacgadTdT-3´ and 5´-ucguuucaaaguuuccgccdGdG- 3´ (Ambion). The
control siRNA was Silencer Negative Control #3 siRNA (Ambion) composed of a 19 bp
scrambled sequence without significant homology to any known genes in rats. After the
seeded cells reached 60% confluency, the cells were transfected with non-sense siRNA or
siRNA against NNT using oligofectamine transfection reagent (Invitrogen) for 24 h
before differentiation.
3.3.3. Measurement of the redox status of the cell – (a) GSH and GSSG levels. GSH and
GSSG concentrations were analyzed using HPLC electrochemical detection (Yap et al.,
2010b). (b) Pyridine dinucleotides. NAD
+
, NADH, NADP
+
, and NADPH levels were
measured by HPLC (Klaidman et al., 1995). (c) Cellular redox status. The cellular redox
status was quantified by the Nernst equation (E
hc
= E
0
+ 30 log ([GSSG]/[GSH]
2
) where
60
[GSH] and [GSSG] are molar concentrations; E
0
was taken as −264 mV at pH 7.4, as
described previously (Yap et al., 2010b). The Nernst potential was calculated from the
molar concentration of GSH and GSSG in cell volume at the value of ∼10 μl/mg of
protein.
3.3.4. H
2
O
2
release and oxygen radical measurement – H
2
O
2
generation from PC12 cells
was determined by the Amplex Red Hydrogen Peroxide/Peroxidase Assay kit (Invitrogen)
following the manufacturer’s instructions. For oxygen radical staining, PC12 cells were
incubated with 10 μM of cell-permeant H
2
DCFDA (2',7'-dichlorodihydrofluorescein
diacetate) for 30 min and 2',7'-dichlorodihydrofluorescein (DCF) fluorescence was
monitored at 495 nm excitation and 520 nm emission.
3.3.5. Metabolic Flux Analysis: XF-Extraflux Analyzer – PC12 cells were cultured on
Seahorse XF-24 plates at a density of 5 x 10
4
cells/well. On the day of metabolic flux
analysis, cells were changed to unbuffered DMEM (DMEM base medium supplemented
with 25 mM glucose, 2 mM sodium pyruvate, 31 mM NaCl, 2 mM GlutaMax, pH 7.4)
and incubated at 37°C in a non-CO
2
incubator for 1 h. All medium and injection reagents
were adjusted to pH 7.4 on the day of assay. Baseline measurements of oxygen
consumption rate (OCR, measured by oxygen concentration change) and extracellular
acidification rate (ECAR, measured by pH change) were taken before sequential injection
of treatments / inhibitors: oligomycin (ATP synthase inhibitor, 4 μM), FCCP
61
(mitochondrial respiration uncoupler, 1 μM), and rotenone (Complex I inhibitor, 1 μM).
After the assays, plates were saved and protein readings were measured for each well to
confirm equal cell number/well.
3.3.6. Mitochondrial membrane potential – Mitochondrial membrane potential was
determined by measuring the ΔΨ
m
-dependent distribution of JC-1 using a fluorescence
spectrometer as described (Salvioli et al., 1997).
3.3.7. Lactic acid and ATP measurements – Differentiated PC 12 cells medium was
collected, acidified with an equal volume of perchloric acid (2 M), and centrifuged for 10
min at 12000 g. The supernatant was neutralized with KHCO
3
(3 M) and centrifuged at
12000 g. 100 μl of the extract was added to 500 μl of reaction buffer and the
concentration of lactic acid was measured in terms of NADH absorbance at 340 nm
(assay kit from Abnova). For ATP measurements, cells were lysed using perchloric acid
(2 mol/L); cell extracts ere neutralized using KHCO
3
as described above. ATP in cell
extracts was quantitatively measured by a bioluminescence assay that uses recombinant
firefly luciferase and D-luciferin (assay kit from Invitrogen).
3.3.8. Measurement of succinyl-CoA:3-oxoacid CoA transferase (SCOT) activity - SCOT
activity was measured as previously described (Williamson et al., 1971). Briefly, cell
lysates were disrupted by sonication and then centrifuged at 1000 g for 20 min. The
62
supernatants were incubated with 50 mM Tris-HCl, 5 mM MgCl
2
, 4 mM iodoacetamide,
0.2 mM acetoacetate, and 0.1 mM succinyl-CoA, pH 8.0. Catalytic activity was measured
spectrophotometrically at 313 nm.
3.3.9. Western blotting – Cell lysate was solubilized in SDS sample buffer, separated by
Laemmli SDS/PAGE, and transferred onto PVDF membranes. Using appropriate
antibodies, the immunoreactive bands were visualized with an enhanced
chemiluminescence reagent.
3.3.10. Apoptosis detection and cell viability assay – (a) Apoptosis detection. Cells were
collected and resuspended at a concentration of 10
6
cells/ml in the binding buffer (10 mM
HEPES/NaOH (pH 7.4), 140 mM NaCl, 2.5 mM CaCl
2
) and incubated with FITC
conjugated Annexin V and propidium iodide (PI) for 15 min at room temperature. Cell
apoptosis was analyzed by FACSDiva (BD Biosciences). (b) Cell viability. Cell viability
was assessed by incubating cells with MTT reagent (5 mg/ml) (Sigma) for 30 min at
37°C in a humidified 5% CO
2
incubator. The reduced intracellular formazan product was
dissolved by replacing 2 ml of differentiation medium with the same volume of DMSO.
The absorbance at 590 nm was measured with a microplate reader.
63
3.3.11. Statistical analysis - Data are reported as means ± SEM of at least three
independent experiments. Significant differences between mean values were determined
by student t-test. Means were considered to be statistically distinct if P < 0.05.
3.4. RESULTS
3.4.1. siRNA knockdown of NNT in PC12 cells
PC12 cells were transfected with siRNA to NNT; a scramble (nonsense) siRNA was
used as a negative control. Expression of NNT mRNA was mostly abolished (∼80%
decrease) by siRNA targeted against it, but was not affected in mock-transfected cells or
those transfected with nonsense siRNA (Fig. 3-1A). Likewise, NNT protein expression
was not affected after transfection of non-sense siRNA but was decreased by ∼60% after
transfection of siRNA against NNT (Fig. 3-1B). NNT mRNA and proteins were
normalized to mitochondrial cytochrome c oxidase subunit IV (COX IV) mRNA and
protein levels, respectively.
64
Figure 3-1. siRNA knockdown of the NNT gene in PC12 cells. PC 12 cells were
transfected with mock, scramble (non-sense) siRNA or NNT siRNA and differentiated for 5
days before RT-PCR and western blot analyses. A. Upper panel: RT-PCR bands of NNT and
COX IV (loading control) mRNA levels; lower panel: relative density of the NNT mRNA
level normalized to COX IV. B. Upper panel: Western Blots of NNT and COX IV (loading
control) protein levels; lower panel: relative density of NNT protein level normalized to COX
IV. *P < 0.05, n = 3.
65
3.4.2. NADPH production, H
2
O
2
release, and the redox status of the cell
The NADPH/NADP
+
ratio in differentiated PC12 cell lysate (collected 6 days after
transfection) significantly decreased in siNNT-transfected PC12 cells as compared to that
of cells transfected with nonsense siRNA (Fig. 3-2A), thus suggesting that NNT is an
important source of NADPH in mitochondria and its function can affect the total cellular
NADPH and NADP
+
pools.
The steady-state level of mitochondrial H
2
O
2
([H
2
O
2
]
ss
) (Boveris and Chance, 1973;
Boveris et al., 1972), is determined at the equilibrium by its sources (mainly the electron
leak in the mitochondrial respiratory chain) and its removal. The latter is driven by
mitochondrial thiols (glutathione, thioredoxin) that are supported by the ultimate electron
donor, NADPH. H
2
O
2
release from PC12 cells with different glucose loadings was
assessed by the peroxidase-based Amplex Red method (Fig. 3-2B): there was no
significant difference in H
2
O
2
release between NNT knockdown group and control group
in low glucose media; at high concentrations of glucose in the media (100 mM), H
2
O
2
released from NNT-suppressed cells was about 35% higher than that from control cells,
thus indicating a compromised H
2
O
2
removal system in NNT-suppressed cells upon
increased metabolic loading and H
2
O
2
generation. It may be surmised that decreased
availability of NADPH after NNT suppression affects the reserved antioxidants capacity
of the cell, i.e., H
2
O
2
removal. In the control group, high glucose (up to 100 mM) did not
induce an increased H
2
O
2
release, which suggests that increased H
2
O
2
release in NNT-
silenced cells is not merely a consequence of high metabolic loading. This is supported
66
Figure 3-2. Effect of NNT knockdown on NADPH production and H
2
O
2
release.
A. NADPH/NADP
+
ratio in differentiated PC12 cells transfected with non-sense or NNT
siRNA. The concentration of NADPH and NADP
+
in cell lysate was measured by HPLC and
the ratio was calculated. B. H
2
O
2
release from live PC12 cells with different glucose
loadings. H
2
O
2
release from PC12 cells incubated in different concentrations of glucose (0-
100 mM) was monitored for 30 min using Amplex Red fluorescence dye and the release rate
was calculated. C. Non-specific detection of cellular oxidants using H
2
DCFDA. PC12 cells
stained with H
2
DCFDA were incubated with different concentrations of glucose for 30 min
and the fluorescence density was measured. *P < 0.05, n = 4.
67
by the fact that increased DCF fluorescence was observed in NNT-suppressed cells even
at low glucose levels (Fig. 3-2C). H
2
DCFDA is a non-specific oxidant detector, but its
good cell penetrability enables the capture of various oxidants intracellularly; hence the
difference in cellular oxidant levels in control and NNT-silenced cells can be observed
regardless of metabolic loadings. The lack of specificity and accuracy of DCFH in
detecting H
2
O
2
and other oxidants (Murphy et al., 2011) is recognized here; in these
experimental conditions, DCF data confirmed the impairment of cellular antioxidant
capacity due to NNT dysfunction in addition to H
2
O
2
release data. Because the specific
detection of H
2
O
2
by Amplex-Red requires horseradish peroxidase (HRP), which is not
membrane-permeant, Amplex-Red/HRP can only measure extracellularly diffusing H
2
O
2
,
hence is less sensitive to the fluctuations of intracellular H
2
O
2
levels. This could explain
the unchanged H
2
O
2
release at low glucose concentrations. A high glucose concentration
was used to magnify the H
2
O
2
production to reveal the impairment of cellular antioxidant
defense in NNT-silenced cells, even though this concentration departs from the
physiological condition. Taken together, the sole utilization of DCFH or Amplex Red
method to detect intracellular oxidant levels has its limitations, either non-specific or
insensitive. A combination of both methods can better characterize the cellular level of
oxidants tentatively before a specific and in situ detection method is developed.
Increased levels of H
2
O
2
can shift the cell from a reduced- to an oxidized state. GSH
plays a central and important role in the removal of H
2
O
2
generated by the electron
transport chain (Chance et al., 1979; Hurd et al., 2005) and the GSH and GSSG
68
concentrations are determinant of the redox status of the cell. After NNT knockdown,
there is a slight decrease of cellular GSH levels and a significant increase of GSSG levels
(Table 3-1). Accordingly, GSH/GSSG ratio decreases from 136.50 ± 32.77 in control
cells to 68.78 ± 10.12 in siNNT-transfected cells, thus accounting for a less negative
redox potential value in siNNT-transfected cells (–235.40 ± 2.56 mV compared to –
245.62 ± 0.67 mV in control cells, Table 3-1). This indicates the importance of NNT in
maintaining the mitochondrial and cellular redox status by regulating NADPH-dependent
GSH regeneration.
TABLE 3-1. GSH AND GSSG LEVELS AND THE REDOX POTENTIAL OF THE CELL
___________________________________________________________________________________________
non-sense siNNT
_____________________________________________________
GSH (nmol/mg protein) 19.20 ± 0.50 16.72 ± 0.84*
GSSG (nmol/mg protein) 0.16 ± 0.03 0.26 ± 0.08*
GSH/GSSG 136.50 ± 32.77 68.78 ± 10.12*
Redox potential (mV) –245.62 ± 2.56 –235.40 ± 0.67*
_________________________________________________________________________________________________
PC12 cells were transfected with non-sense or NNT siRNA and differentiated for 5
days before collection. Cell lysates were subjected to HPLC analyses of GSH and
GSSG. Concentrations expressed in nmol/cell lysate protein. The redox potential of the
cell was estimated by the equation: E
hc
= E
0
+ 30 log ([GSSG]/[GSH]
2
. *p < 0.05; n = 4.
69
3.4.3. Energy metabolism in siNNT-transfected PC12 cells
Mitochondria provide most of the energy needed for cellular functions by the
conversion of energy in fuel molecules into ATP through oxidative phosphorylation.
Oxygen consumption rate (OCR) by mitochondria reflects the activity of mitochondrial
bioenergetics, and is thus an important parameter of mitochondrial function. At 6 days
after transfection, siNNT-transfected cells had a substantially lower basal OCR relative to
control cells (Fig. 3-3A); following the addition of oligomycin, OCR declined in both
control- (334 pmols/min decrease, ~65% of basal) and siNNT-transfected cells (84
pmols/min decrease, ~50% of basal) (Fig. 3-3A; 25-50 min), indicating that ATP
turnover was significantly lower in NNT knockdown cells than in control cells. Maximal
respiratory capacity –measured after the addition of the uncoupler FCCP– was
substantially lower in PC12 cells with NNT knocked down (Fig. 3-3A; 50-75 min). The
addition of the complex I inhibitor rotenone resulted in a further reduction in OCR (12%
of basal OCR remained for control and 16% of basal OCR remained for siNNT); the
residual O
2
consumption was accounted for by non-mitochondrial O
2
-consuming
pathways (Fig. 3-3A; 75-100 min).
Anaerobic glycolysis, the conversion of glucose to lactate, provides a modest amount
of ATP. The extracellular acidification rate (ECAR) was used to detect the glycolytic
activity by monitoring pH changes due to lactic acid generation. There were no
significant differences in ECAR between control and siNNT-transfected cells (Fig. 3-3B).
The increase in ECAR in response to oligomycin (in both siNNT- and non-sense
70
transfected cells) indicated a shift to ATP production by glycolysis via the Pasteur effect
(Guppy et al., 1995). Decreased ATP levels in NNT knockdown cells (Fig. 3-3C)
suggests that the overall energy metabolism is compromised with NNT suppression,
because mitochondrial oxidative phosphorylation contributes to the majority of ATP
production in the cell; the extracellular lactic acid levels (Fig. 3-3D) is consistent with the
ECAR data indicating an unchanged anaerobic glycolysis rate. Compared to the dramatic
decrease in OCR in siNNT-transfected cells, the unchanged ECAR values suggested that
a larger proportion of the total cellular ATP was generated through glycolysis. In these
experiments, OCR and ECAR values were normalized to total protein concentration in
each well. Considering the decrease of cell viability in NNT-suppressed cells (as shown
below in section 3.5), the dramatic decline in mitochondrial respiration should be
reflecting the combined effects of impaired mitochondrial function in individual living
cells and a decrease of overall cell density. The increased glycolysis level in individual
cells was seemingly offset by decreased cell number, which made the final output reading
unchanged. Hence, an OCR/ECAR ratio analysis was conducted that confirmed that ratio
of O
2
consumption- and extracellular acidification rate were significantly lower in
siNNT-transfected PC12 cells (Fig. 3-3E). Another factor that may be taken into
consideration regarding unchanged glycolysis is that some components of the glycolytic
machinery in the cytosol are sensitive to redox modification (such as GAPDH (Yap et al.,
2010a)) and may thereby be inhibited due to redox state change after NNT knockdown.
71
Figure 3-3. Cellular energy metabolism in siNNT-transfected PC12 cells. PC12 cells
were transfected with non-sense or NNT siRNA and differentiated for 5 days. OCR and
ECAR were determined using Seahorse XF-24 Metabolic Flux Analyzer. Vertical dashed
lines indicate time of addition of mitochondrial inhibitors: oligomycin (4 μM), FCCP (1 μM),
or rotenone (1 μM). A. OCRs in NNT-suppressed cells (open circles) have lower basal rates
and maximal rates (after the addition of FCCP) of mitochondrial respiration than those of
control cells (filled circles). B. No significant difference in ECAR was observed after NNT
knockdown (open circles), compared to control groups (filled circles). C. Cellular ATP levels
decreased after NNT knockdown. D. No significant change in extracellular lactate levels was
observed in NNT suppressed cells; E. The OCR/ECAR ratio of NNT-suppressed cells (open
circles) is lower than that of control group (filled circles) at basal conditions and after the
addition of FCCP. *P < 0.05, n = 5. OCR, ECAR, ATP and lactate readings were normalized
to total protein concentration in each well.
72
3.4.4. Components in mitochondrial bioenergetics
NNT generates NADPH from NADP
+
using NADH as reducing equivalents and
proton gradient across mitochondrial inner membrane as the driving force (Fig. 3-4A).
Both, NADH and H
+
gradients are highly involved in mitochondrial energy metabolism.
The NADH/NAD
+
values were substantially lower in differentiated siNNT-transfected
cells than in control cells (Fig. 3-4B); suppression of NNT activity resulted in a decline of
NADH/NAD
+
ratio (Fig. 3-4B); likewise, the decrease in ΔΨ
m
(Fig. 3-4C) could be a
consequence of limited NADH supply. ΔΨ
m
is established by H
+
pumping along with
electron flow through Complex I-IV of the respiratory chain, and NADH is the initial
electron donor.
73
Figure 3-4. Effect of NNT knockdown on mitochondrial bioenergetic machinery. A.
Schematic representation of the interaction between NNT function and mitochondrial
bioenergetic components. NADH generated in TCA cycle can either be used by respiratory
chain to build up membrane potential or be used by NNT to reduce NADP
+
to NADPH;
membrane potential is also the driving force for NNT to generate NADPH in physiological
conditions. B. NADH/NAD
+
ratio in differentiated PC12 cells decrease after transfection of
NNT siRNA. The concentration of NADH and NAD
+
in cell lysate was measured by HPLC
and the ratio was calculated. C. Mitochondrial membrane potential (ΔΨ
m
) is lower in NNT-
suppressed cells than control group. PC12 cells were differentiated and stained with JC-1
dye, and the ratio of red and green fluorescence intensity was calculated. *P < 0.05, n = 4.
74
These data suggest that the decreased OCR in siNNT-transfected cells could be
ascribed to decreased NADH availability accompanied by a decline in mitochondrial
membrane potential. It was expected that suppression of NNT activity would lead to an
accumulation of NADH (which is consumed by NNT to generate NADPH in
physiological conditions) rather than a decline of NADH. We therefore performed a time-
response study analyzing the effect of NNT knockdown on the energy and redox status of
the cell. As shown in Fig. 3-5, 1 day after siNNT transfection, the NADPH/NADP
+
ratio
has decreased by about 35% (Fig. 3-5C), while the NADH/NAD
+
ratio increased by
about 25% (Fig 3-5D). At the same time point, the OCR and ECAR is not changed
compared to control group (Fig. 3-5A and 3-5B). These data indicate that the alteration of
redox status of the cell precedes the impairment of mitochondrial energy metabolism.
The increase in NADH/NAD
+
ratio is in agreement with Eq. 1, i.e., NNT knockdown
initially leads to NADPH shortage and NADH accumulation. It may be suggested that
indirect effects following NNT suppression are involved in the control of NADH levels
in siNNT-transfected cells. In this regard, the pyruvate dehydrogenase (PDH) complex is
critical to the cellular energy metabolism inasmuch as it controls the entry of substrates
(acetyl-CoA) to the TCA cycle and the further generation of reducing equivalents
(NADH). Western blotting showed a substantial increase in the phosphorylated (inactive)
form of the E
1α
subunit of PDH along with a slight decrease in total PDH in siNNT-
transfected cells (Fig. 3-6A), which result in an increased inhibition of PDH (Fig. 3-6B).
75
The inhibition of PDH activity may account for the decreased NADH/NAD
+
values
observed in cells with a diminished NNT activity (Fig. 3-4B).
Succinyl-CoA-transferase (SCOT) alternatively provides acetyl-CoA to the TCA
cycle upon metabolism of ketone bodies (e.g. acetoacetate and β-hydroxybutyrate);
SCOT activity was also found decreased by ~34% in NNT-transfected cells (Fig. 3-6C),
thus limiting the alternative NADH generation.
Taken together, the decreased NADH/NAD
+
values observed in differentiated cells
upon suppression of NNT activity appear to be a consequence of a limited acetyl-CoA
supply (and further generation of NADH) to the TCA cycle imposed by an inhibited PDH
complex and decreased SCOT activity. This could also accounts for the substantially
diminished OCR and ATP levels in siNNT-transfected cells.
76
Figure 3-5. Alteration of redox status after NNT knockdown precedes the impairment
of energy metabolism. 1 day after siNNT transfection, OCR and ECAR were determined
using Seahorse XF-24 Metabolic Flux Analyzer. Vertical dashed lines indicate time of
addition of mitochondrial inhibitors: oligomycin (4 μM), FCCP (1 μM), or rotenone (1 μM).
A. OCRs in NNT-suppressed cells (open circles) remain unchanged in basal rates and
maximal rates (after the addition of FCCP) of mitochondrial respiration compared to control
cells (filled circles). B. No significant difference in ECAR was observed after NNT
knockdown (open circles), compared to control groups (filled circles). C. NADPH/NADP
+
ratio in differentiated PC12 cells 1 day after transfection with non-sense or NNT siRNA; D.
NADH/NAD
+
ratio in differentiated PC12 cells 1 day after transfection with non-sense or
NNT siRNA.
77
Figure 3-6. PDH and SCOT activities in siNNT-transfected PC12 cells. A. Western blots
of phospho-PDH-E
1α
and total PDH-E
1α
levels. COX IV was used as loading control. B.
PDH activity was inhibited after NNT knockdown as shown by significantly increased
relative phosphorylation level of PDH-E
1α
. C. SCOT activity was inhibited after NNT siRNA
transfection in differentiated PC12 cells. *P < 0.05, n = 4.
78
3.4.5. Activation of JNK and apoptotic pathways
Previous work demonstrated that JNK, a member of the MAPK family, translocates
to mitochondria upon activation and initiates a signaling cascade across both
mitochondrial membranes resulting in phosphorylation (and inhibition) of mitochondrial
matrix PDH (Zhou et al., 2008; Zhou et al., 2009). Besides, mitochondrion-generated
H
2
O
2
is involved in the modulation of redox-sensitive cell signaling such as JNK
pathway (Chen et al., 2001; Foley et al., 2004; Nemoto et al., 2000; Saitoh et al., 1998);
accordingly, the increased H
2
O
2
release from NNT-suppressed PC12 cells (Fig. 3-2B)
was associated with JNK1 activation (its phosphorylated form) (Fig 3-7A) as well as an
increase in p-JNK1/JNK1 values (Fig. 3-7B). This links the altered mitochondrial redox
status to impaired mitochondrial energy metabolism through H
2
O
2
levels, JNK activation,
and PDH inhibition. The interaction of JNK with mitochondria suggests a network
involving cytosolic and mitochondrial processes that control cellular energy levels and
the redox environment. Expectedly, these redox-energy changes are associated with a
marked decrease in cellular viability (Fig. 3-7C), partly supported by the effect of JNK on
mitochondrion-dependent apoptosis (Harris et al., 2002; Kharbanda et al., 2000; Putcha et
al., 1999; Schroeter et al., 2003). FACS analyses of NNT-suppressed PC12 cells (Fig. 3-
7D) showed a significant higher population of apoptotic cells (early and late stages of
apoptosis, Fig. 3-7E). These results suggest the important role of NNT in the regulation
of cellular function though mitochondria-generated H
2
O
2
(as second messenger) and
redox-sensitive signaling.
79
Figure 3-7. Activation of JNK and initiation of apoptosis in siNNT-transfected PC12
cells. A. Western blots of phospho-JNK1 and total JNK1 levels. β-actin was used as loading
control. B. JNK activity was enhanced after NNT knockdown as shown by its increased
relative phosphorylation level. C. Cell viability decreased after NNT siRNA transfection in
differentiated PC12 cells using MTT assay at 590 nm. D. Representative image of FACS
analyses of apoptosis in PC12 cells using FITC-conjugated AnnexinV and propidium iodide
(PI) double staining. E. NNT-suppressed group has more than 4-fold higher population of
apoptotic cells as compare with control group. *P < 0 .05, n = 4.
0
150
300
p-JNK1/JNK1
(Relative Density to non-sense %)
siNNT non-sense
β-actin
p-JNK1
JNK1
A B
0
0.5
1
590nm absorbance (AU)
non-sense siNNT
C
*
non-sense siNNT
0
20
40
Apoptotic Cells (%)
non-sense siNNT
*
– 46
– 46
– 42
*
E D
kD
80
3.5. DISCUSSION
The results reported here with NNT-suppressed PC12 cells showing that NNT plays a
critical role in regulating cellular redox status and energy metabolism as well as cytosolic
redox-sensitive signal pathways further confirm the significant functional role of NNT as
a mitochondrial NADPH source in other organisms or tissues (Arkblad et al., 2005;
Sheeran et al., 2010). siRNA silencing of NNT in PC12 cells results in decline of
NADPH production and an oxidized cellular redox status as inferred by increased GSSG
levels, decreased GSH/GSSG ratio, and a less negative redox potential. Furthermore, a
decrease in NADPH supply in NNT-knockdown cells results in an augmented H
2
O
2
release, consistent with previous reports on an insulin-secreting β-cell line (Freeman et al.,
2006), in which an increase in DCF fluorescence was found in NNT
-/-
β cells at glucose
loading of 20 mM.
The consequences of altered redox status on regulation of mitochondrial energy
metabolism may be effected through the redox modulation of cytosolic signaling
cascades and through redox-mediated post-translational modifications of mitochondrial
proteins (Yap et al., 2009). The first pathway is primarily through the
activation/inhibition of numerous redox-sensitive signaling pathways (such as JNK and
other MAPKs) by H
2
O
2
released from mitochondria. In the NNT-suppressed PC12 cell
model, increased H
2
O
2
production from mitochondria and a more oxidized redox
environment are associated with activation of cytosolic JNK, which further translocates
81
to mitochondria (as shown with primary cortical neurons (Zhou et al., 2008)) and initiates
a phosphorylation cascade that inhibits PDH activity upon phosphorylation of the E
1α
subunit, thereby increasing energy deficits. SCOT, which catalyzes the rate-limiting step
of generation of acetyl-CoA from ketone bodies, also exhibited a lower activity in NNT
knockdown cells; previous work in our laboratory showed that SCOT could be post-
translationally modified upon altered redox status, either by glutathionylation (Garcia et
al., 2010) or nitration (Lam et al., 2009), and both modifications decreased its activity.
Inhibition of PDH and SCOT is expected to lead to a decrease of substrate (i.e., acetyl-
CoA) entry into the TCA cycle, thereby limiting the NADH generation, as confirmed by
the reduced NADH/NAD
+
ratios. A decreased NADH supply to the mitochondrial
respiratory chain would also account for the reduced membrane potential and maximal
respiratory capacity (Fig. 3-4C and Fig. 3-3A). The compromised mitochondrial
respiration in NNT knockdown PC12 cells (Fig. 3-3A) is associated with unchanged
anaerobic glycolysis levels (Fig. 3-3B and 3-3D), indicating an increased contribution of
anaerobic glycolysis to cellular ATP production whereas the total ATP levels is declined
(Fig. 3-3C). A similar shift towards glycolysis was reported in JNK-treated primary
cortical neurons with an inhibited PDH activity (Zhou et al., 2008).
NNT activity provides a link between the mitochondrial metabolic function (energy-
transducing activity) and redox homeostasis by coupling NADPH generation to the TCA
cycle and active respiration. Hence, NNT plays a critical role in the maintenance of the
82
mitochondrial energy – redox axis. This supports the notion that decline in cellular
bioenergetics and changes in the redox status of the cell cannot be viewed as independent
events, but rather as an interdependent relationship centered on the mitochondrial energy
– redox axis (Yap et al., 2009). Disruption of electron flux from fuel substrates to redox
components (as a result of NNT dysfunction) induces not only altered redox status by
also impaired energy metabolism. It is also suggested that in addition to mitochondrial
energy-transducing capacity and redox homeostasis, impairment of NNT activity also
affects the cellular functions through interactive communication between mitochondrion-
generated second messengers and cytosolic redox-sensitive signaling. This study shows
that the modulation of NNT function could be important in the collective impairments of
the interdependent mitochondrial energy-redox axis and the regulation of cytosolic redox-
sensitive signaling inherent in several pathophysiological situations. Data obtained in this
study potentially explain the underlying mechanisms of the poor response of NNT
-/-
C57Bl/6J mice to glucose (Freeman et al., 2006), and they also explain the lethal effect of
combining a deficiency of both SOD2 and NNT (Huang et al., 2006). Moreover,
investigations of the physiological and pathological roles of NNT (Freeman et al., 2006)
will expand the understanding of the mechanisms that support energy- and/or redox
deficits in the early or late stages of some neurodegenerative diseases (Lin and Beal,
2006; Yao et al., 2009; Zhou et al., 2009), diabetes (Kaneto et al., 2005; Lowell and
Shulman, 2005), cardiovascular disease (Sheeran et al., 2010), and aging (Mattson and
Magnus, 2006; Navarro and Boveris, 2007b; Van Remmen and Jones, 2009).
83
3.6. CONCLUSION
As an important mitochondrial NADPH source, knockdown of NNT alters the
mitochondrial and cellular redox status (GSH depletion and H
2
O
2
production), along with
compromised mitochondrial bioenergetics by inhibition of mitochondrial metabolism
(including inhibition of PDH and SCOT activity). The limited NADH availability
imposed by inhibition of the latter enzymes could account for the decline of membrane
potential, ATP turnover, and maximal respiratory capacity. Activation of redox-sensitive
signaling (JNK) by H
2
O
2
induces mitochondrion-dependent intrinsic apoptosis and
results in decreased cell viability.
84
CHAPTER 4
LOSS OF NICOTINAMIDE NUCLEOTIDE TRANSHYDROGENASE
IN C57BL/6J MICE INDUCES DISTINCT RESPONSES
IN BRAIN AND IN LIVER
4.1. ABSTRACT
Regulation of mitochondrial H
2
O
2
homeostasis and its involvement in the regulation of
redox-sensitive signaling is the consequence of the concerted activities of the
mitochondrial energy- and redox systems. The energy-dependent nicotinamide nucleotide
transhydrogenase (NNT) produces the majority of mitochondrial NADPH for the redox
systems as reducing equivalents. Loss of NNT function accounts for the impaired glucose
tolerance in C57BL/6J mice. In this study, the role of NNT in linking the mitochondrial
energy status to the redox environment was investigated in brain and liver of these mice.
The results show that NNT regulates mitochondrial NADP(H) levels in an energy-
sensitive manner in both brain and liver. In liver, the NADP(H) pool is more affected by
loss of NNT activity and is associated with altered NAD(H) pool, which is not seen in
85
brain. The diminished impact of NNT dysfunction in brain could be due to an
upregulation of another NADPH-producing enzyme, IDH2. In primary hepatocytes, loss
of NNT led to impaired mitochondrial energy-transducing capacity, which could be
mediated by the modulation of multiple redox-sensitive cytosolic signaling pathways by
altered mitochondrial redox status in the absence of NNT. Taken together, loss of NNT in
C57BL/6J mice induces more significant consequences in liver than in brain, indicating a
more important role of this enzyme in liver.
4.2. INTRODUCTION
Mitochondrial NADPH is the ultimate reductant that controls the mitochondrial redox
homeostasis through glutathione- and thioredoxin-based systems. Sources of NADPH in
mitochondria are mainly isocitrate dehydrogenase-2 and nicotinamide nucleotide
dehydrogenase (NNT). The former, isocitrate dehydrogenase-2, catalyzes the reduction
from NADP
+
to NADPH by decarboxylating isocitrate to α-ketoglutarate. The latter,
NNT, catalyzes the hydride transfer from NADH to NADP
+
using proton gradient across
the inner mitochondrial membrane as driving force. Under physiological conditions, NNT
regulates NADPH levels by receiving reducing equivalents from NADH to produce high
86
concentrations of NADPH, and also buffers uncontrolled changes in metabolites
associated with NAD(H) (Hoek and Rydström, 1988 ).
NNT activity is regulated by multiple factors including its product concentration and
more importantly, the mitochondrial energy status. Firstly, NNT is largely thought to be
product-inhibited at high NADPH/NADP
+
values, and its maximal activity (~30
nmol/min/mg mitochondrial protein) is only reached upon a consumption of the product
NADPH at a rate exceeding the capacity of NNT (Rydstrom, 2006). Secondly, under
coupled conditions, the extent of the inhibition by NADPH is influenced by the
prevailing proton motive force (Δp), because the affinities of NNT for its substrates are
Δp-dependent (Rydstrom et al., 1971). Thirdly, NNT activity is highly dependent on
availability of energy substrates because of the connection between its activity and the
NAD pool. Its affinity for NADP
+
increases 5-fold and that for NAD
+
decreased 5-fold in
the presence of energy sources, which in turn strongly stimulate the forward reaction, i.e.,
the generation of NADPH. The inhibition of the forward reaction by NADPH or NAD
+
decreases several fold with an increase in Δp, whereas the inhibition of the reverse
reaction by NADH or NADP
+
shows a corresponding increase by energization (Hoek and
Rydström, 1988 ).
The crucial role of NNT in the maintenance of redox environment is revealed by the
fact that ablation of NNT renders C. elegans more susceptible to oxidative stress and
decreases the cellular GSH/GSSG ratio (Arkblad et al., 2005). Mice deficient in SOD2
die much earlier if they also lack functional NNT (Huang et al., 2006). Moreover, in β-
87
cells from mice carrying loss-of-function mutations in NNT, glucose causes a dramatic
increase in H
2
O
2
level and declined insulin secretion (Freeman et al., 2006). These
evidences indicate the importance of NNT in maintaining NADPH level and redox
homeostasis. NNT is also proposed to be involved in a substrate cycle with NAD- and
NADP-linked isocitrate dehydrogenase, which contributes to regulation of the TCA cycle
in mitochondria (Sazanov and Jackson, 1994), indicating potential interaction between
NNT and mitochondrial energy components. NNT is important for integrating
mitochondrial and cytosolic metabolism and for maintaining mitochondrial function
under conditions of anoxia or high energy demand (Hoek and Rydström, 1988 ).
C57BL/6J mice exhibit impaired glucose tolerance that is independent of obesity
(Kooptiwut et al., 2002). The intraperitoneal glucose tolerance test (IPGTT) shows the
blood glucose levels of non-obese C57BL/6J mice are higher and take longer to regain
the resting level than that of other mouse strains (Toye et al., 2005). High-fat diets induce
insulin resistance, hyperglycemia, and diabetes in these mice (Burcelin et al., 2002;
Rossmeisl et al., 2003; Surwit et al., 1991). Toye et al identified three main genetic loci
that influence the difference in glucose homeostasis under fasting conditions and in
response to glucose challenge in C57BL/6J mice by intercrossing non-obese C57BL/6J
and C3H/HeH mice and applying quantitative trait locus (QTL) mapping (Toye et al.,
2005). These loci were located on chromosomes 9, 11, and 13, and NNT gene is encoded
within the chromosome 13 locus. Two mutations were identified after sequencing NNT
coding region in C57BL/6J mice: a missense mutation (M35T) in the mitochondrial
88
localization sequence (MLS) of the NNT precursor protein, and an in-frame 5-exon
deletion that removes four predicted trans-membrane helices and their connecting linkers.
The 5-exon deletion deletes part of domain I of NNT and the first four trans-membrane
domains and leads to a marked downregulation of NNT mRNA in liver and islets (Toye
et al., 2005).
NNT is expressed at high levels in mouse heart, kidney, testes, adrenal, liver, pancreas,
bladder, lung, ovary, and brain and at low levels in skeletal muscle and spleen; NNT
expression in liver is ~2-2.5 fold-higher than in brain (Arkblad et al., 2002; Chico et al.,
1977). Currently, it is not clear about the exact role of NNT in regulating mitochondrial
NAPDH pool and redox status, especially how that is connected to mitochondrial NADH
pool and bioenergetic function. It is also unknown that how the different expression
levels of NNT in various tissues influence mitochondrial function and the redox status of
those tissues.
In this study, by comparing C57BL/6J mice containing mutant NNT (essentially a
NNT KO mice) with its wild type control C57BL/6NJ mice, we examined the
consequences of NNT dysfunction on mitochondrial pyridine dinucleotides pool as well
as its regulation by mitochondrial energy status. The different responses of brain and liver
mitochondria to the absence of NNT were also compared. Moreover, in primary
hepatocytes isolated form C57BL/6J and C57BL/6NJ mice, we validated the role of NNT
that are found in NNT-suppressed PC12 cells in terms of modulating mitochondrial
89
energy metabolism and redox homeostasis, and of regulating cytosolic signaling
pathway(s) that are modulated by metabolic or oxidative signals.
4.3. MATERIALS AND METHODS
4.3.1. Animals
The NNT-mutant C57BL/6J male mice and their wild type control C57BL/6NJ were
purchased from Jackson Lab. The animals are bred under stringent genetic and
environmental controls and each stock and strains are well characterized.
4.3.2. Mitochondria isolation
Mitochondria from mouse brain and liver were isolated by using discontinuous Percoll
gradient and differential centrifugation, respectively (Garcia et al., 2010).
Discontinuous Percoll gradient – Brain mitochondria were isolated based on
methodology previously described (Anderson and Sims, 2000). Brain tissue are excised,
chopped into fine pieces, washed with and homogenized in isolation buffer containing
210 mM mannitol, 70 mM sucrose and 2 mM HEPES, pH 7.4, plus 0.05% (w/v) BSA.
The homogenate was centrifuged at 800 g for 8 min, the pellet was removed, and the
centrifugation process was repeated. The supernatant was centrifuged at 8000 g for 10
min, the pellet was washed with the isolation buffer, and the centrifugation was repeated.
90
The pellet containing a mixture of organelles was further fractionated by centrifugation at
8500 g for 10 min in a Percoll gradient (consisting of three layers of 18, 30 and 60%
(w/v) Percoll in sucrose/Tris buffer (0.25 M sucrose, 1 mM EDTA and 50 mM Tris/HCl),
pH 7.4). Mitochondria were collected from the interface of 30% and 60% Percoll and
washed with the sucrose/Tris buffer. Mitochondrial protein concentration was determined
using protein assay reagent (Biorad).
Differential centrifugation – Livers from mice were excised, washed, and homogenized
in isolation buffer above using a loose Teflon pestle. The homogenate was centrifuged at
1000 g for 10 min at 4 °C, the pellet was removed, and the centrifugation process was
repeated. The resulting supernatant was centrifuged at 9,000 g for 15 min to generate the
mitochondria pellet, which was washed with isolation buffer, and the high speed
centrifugation was repeated. The mitochondria pellet was resuspended in isolation buffer
(without BSA) before HPLC analysis and other assays.
4.3.3. Pyridine dinucleotides levels
NAD
+
, NADH, NADP
+
, and NADPH levels were measured by HPLC (Klaidman et al.,
1995). Briefly, isolated mitochondria or cell pellet were homogenized in buffer (0.06 M
KOH, 0.2 M KCN, and 1 mM bathophenanthroline disulfonic acid) followed by
chloroform extraction. Chloroform extraction was carried out by centrifugation at 14,000
rpm in a microcentrifuge at 4 °C; the resulting aqueous supernatant with soluble pyridine
nucleotides was collected and extracted thrice to remove lipids and proteins. Finally, it
91
was filtered with a 0.45-μm positively charged filter (Pall Life Sciences) to remove RNA
and DNA in microcentrifuge at 4 °C. The mobile phase consisted of 0.2 M ammonium
acetate (buffer A) at pH 5.5 and HPLC grade methanol (buffer B). A gradient program
with initial conditions as 100% buffer A and 0% buffer B was set. From 0 to 4 min, we
used 0 to 3% B, and from 4 to 23 min, we used 3 to 6.8% B, followed by washing the
column with 50% A and 50% B, and re-equilibrated to initial conditions for next run.
Quantitation of pyridine nucleotides was performed by integrating the peaks and adding
the cyanide adducts as detected by the fluorescence spectrophotometer (λ
exc
= 330nm; λ
em
= 460 nm).
4.3.4. H
2
O
2
release
H
2
O
2
generation from freshly isolated mitochondria was determined by the Amplex Red
Hydrogen Peroxide/Peroxidase Assay kit (Invitrogen) following the manufacturer’s
instructions, with the presence of 5mM of glutamate, 5mM of malate and no ADP.
4.3.5. Western blotting
Cell or mitochondria lysate was solubilized in SDS sample buffer, separated by
SDS/PAGE, and transferred onto PVDF membranes. Using appropriate antibodies, the
immunoreactive bands were visualized with an enhanced chemiluminescence reagent.
92
4.3.6. Metabolic Flux Analysis: XF-Extraflux Analyzer
Primary hepatocytes were cultured on Seahorse XF-24 plates at a density of 5 x 10
4
cells/well. On the day of metabolic flux analysis, cells were changed to unbuffered
DMEM (DMEM base medium supplemented with 25 mM glucose, 2 mM sodium
pyruvate, 31 mM NaCl, 2 mM GlutaMax, pH 7.4) and incubated at 37°C in a non-CO
2
incubator for 1 h. All medium and injection reagents were adjusted to pH 7.4 on the day
of assay. Baseline measurements of oxygen consumption rate (OCR, measured by
oxygen concentration change) and extracellular acidification rate (ECAR, measured by
pH change) were taken before sequential injection of treatments / inhibitors: oligomycin
(ATP synthase inhibitor, 4 μM), FCCP (mitochondrial respiration uncoupler, 1 μM), and
rotenone+antimycin A (Complex I and III inhibitor, 1 μM each). OCR and ECAR were
automatically calculated and recorded by the Seahorse XF-24 software. After the assays,
protein level was determined for each well to confirm equal cell density per well. The
percentage of change relative to the basal rate was calculated as the value of change
divided by the average value of baseline.
4.3.7. Statistical analyses of experimental data
The general analytic approach utilized a comparison of means across experimental
conditions using analysis of variance (ANOVA) procedures (which reduce to a t-test in
the case of 2 experimental conditions).
93
4.4. RESULTS
4.4.1. Pyridine dinucleotide pool in brain mitochondria
NNT levels were examined in brain mitochondria from C57BL/6NJ (WT) and
C57BL/6J mice (NNT mutant). NNT expression was absent in brain mitochondria from
C57BL/6J mice (Fig. 4-1A). In the absence of substrates, NADPH and NADP
+
levels in
the brain mitochondria of NNT mutant mice were lower than those in brain mitochondria
from WT mice (Fig. 4-1B and 4-1C) but no differences were observed in
NADPH/NADP
+
ratios between these two lines (Fig. 4-1D), indicating that NAPDH
production was not affected by NNT dysfunction in brain mitochondria if no energy
substrate was added. In energized mitochondria (supplemented with complex I substrates,
i.e., malate/glutamate), NADPH levels decreased while NADP
+
levels increased in both
mouse lines compared (as compared to a non-energized condition). Lower NADPH levels
in NNT mutant mice resulted in a lower NADPH/NADP
+
ratios compared with WT
group (2.61 versus 1.84). This suggests that energized mitochondria, NADPH production
is moderately impacted by NNT dysfunction. The addition of CCCP, which collapses
mitochondrial proton gradient, led to decreased NADPH levels, increased NADP
+
levels,
as well as declined NADPH/NADP
+
ratios in both lines, but no difference were observed
between WT and NNT mutant mice. This suggests that the differences in NADPH
production in these two lines due to disparity of NNT activity were eliminated upon loss
of proton gradient, which is the driven force of NNT-catalyzed NADPH production.
94
Figure 4-1. Loss of NNT function moderately affects mitochondrial NADP(H) pool in
brain. Brain mitochondria from WT and NNT mutant mice were isolated, and the NADPH
and NADP
+
levels were determined under different conditions: (1) without any substrates, (2)
with the addition of 2.4 mM glutamate and 2.4 mM malate, and (3) with 4 μM of CCCP in
addition to glutamate/malate. A. Expression of NNT in brain of WT and NNT mutant mice;
B. NADPH concentrations in brain mitochondria of WT and NNT mutant mice under
different conditions; C. NADP
+
concentrations in brain mitochondria of WT and NNT
mutant mice under different conditions; D. NADPH/NADP
+
ratios in brain mitochondria of
WT and NNT mutant mice under different conditions.
0.0
0.5
1.0
NADP
+
(nmol/mg)
0.0
1.5
3.0
NADPH/NADP
+
0.0
0.5
1.0
NADPH (nmol/mg)
NNT
COX IV
~110 kD
~19 kD
WT NNT-KO
G/M
CCCP
−
+
+ +
− −
G/M
CCCP
−
+
+ +
− −
G/M
CCCP
−
+
+ +
− −
WT
NNT KO
A
C D
B
WT
NNT KO
WT
NNT KO
95
NNT links the mitochondrial NADP pool with the NAD pool. The mitochondrial
levels of NADH and NAD
+
and their relative ratios were determined in brain
mitochondria from WT and NNT mutant mice. In both lines, the addition of
glutamate/malate strongly stimulated the NADH production (Fig. 4-2A) and NAD
+
consumption (Fig. 4-2B), while CCCP accelerated NADH consumption by the electron
transport chain and led to an accumulation of NAD
+
. Of note, no significant differences
were found in NADH, NAD
+
or their ratio (Fig. 4-2C) where comparing WT and NNT
mutant mice, indicating that in brain, NNT activity does not elicit a significant effect on
the mitochondrial NAD pool.
96
Figure 4-2. Loss of NNT function does not affect mitochondrial NAD(H) pool in brain.
Brain mitochondria from WT and NNT mutant mice were isolated, and the NADH and
NAD
+
levels were determined under different conditions: (1) without any substrates, (2) with
the addition of 2.4 mM glutamate and 2.4 mM malate, and (3) with 4 μM of CCCP in
addition to glutamate/malate. A. NADH concentrations in brain mitochondria of WT and
NNT mutant mice under different conditions; B. NAD
+
concentrations in brain mitochondria
of WT and NNT mutant mice under different conditions; C. NADH/NAD
+
ratios in brain
mitochondria of WT and NNT mutant mice under different conditions.
0.0
2.0
4.0
6.0
NAD (nmol/mg)
0.0
0.5
1.0
NADH/NAD
0.0
1.5
3.0
NADH (nmol/mg)
G/M
CCCP
−
+
+ +
− −
G/M
CCCP
−
+
+ +
− −
G/M
CCCP
−
+
+ +
− −
A
C
B
WT
NNT KO
WT
NNT KO
WT
NNT KO
97
4.4.2. Pyridine dinucleotide pool in liver mitochondria
With the same conditions applied in brain mitochondria, pyridine dinucleotide pools
in liver mitochondria were also examined. As in brain, functional NNT is also completely
abolished in liver mitochondria isolated from NNT mutant mice (Fig. 4-3A). The loss of
NNT function elicited distinct effect in liver mitochondria compared to that in brain.
Even without the addition of energy substrates, there is a difference in NADP
+
levels in
these two lines: higher NADP
+
(Fig. 4-3C) resulted in decreased NADPH/NADP
+
ratio
(Fig. 4-3D) in the NNT mutant mice. This suggests that the NADP pool in liver
mitochondria is more sensitive to the loss of NNT activity than that in brain. With the
addition of glutamate/malate, WT mitochondria responded with an increased NADPH
levels, decreased NADP
+
levels and increased NADPH/NADP
+
ratio (4.12±1.07 to
7.10±1.05); neither NADPH nor NADP
+
levels were responsive to glutamate/malate in
NNT mutant mitochondria (NADPH/NADP
+
ratio: 2.23±0.90 to 2.19±0.68). The
comparison between these two lines with substrate present showed a dramatic decrease in
NADPH/NADP
+
ratio in NNT mutant mice (7.10±1.05 versus 2.19±0.68). This suggests
the critical role of NNT in producing NADPH by utilizing reducing equivalents generated
from energy substrates in liver mitochondria. Upon uncoupling by CCCP, mitochondria
from WT mice exhibited a significant decline in NADPH levels and increase in NADP
+
levels, but these changes were less significant in NNT mutant mice: the NADPH/NADP
+
ratio decreased from 7.10±1.05 to 0.50±0.42 in WT mice and from 2.19±0.68 to
0.53±0.44 in NNT mutant mice. The addition of CCCP eliminated the functional
98
differences of WT and NNT mutant mitochondria in NADPH production, because of the
dependence of NNT function on the proton driven force across the membrane.
Figure 4-3. Loss of NNT function elicits substantially impacts on mitochondrial
NADP(H) pool in liver. Liver mitochondria from WT and NNT mutant mice were isolated,
and the NADPH and NADP
+
levels were determined under different conditions: (1) without
any substrates, (2) with the addition of 2.4 mM glutamate and 2.4 mM malate, and (3) with 4
μM of CCCP in addition to glutamate/malate. A. Expression of NNT in liver mitochondria of
WT and NNT mutant mice; B. NADPH concentrations in liver mitochondria of WT and
NNT mutant mice under different conditions; C. NADP
+
concentrations in liver mitochondria
of WT and NNT mutant mice under different conditions; D. NADPH/NADP
+
ratios in liver
mitochondria of WT and NNT mutant mice under different conditions. Bars represent mean
values ± SEM of 4 animals per group (* p < 0.05).
NNT
COX IV
~110 kD
~19 kD
WT NNT-KO
A
C D
B
WT
NNT KO
WT
NNT KO
WT
NNT KO
G/M
CCCP
−
+
+ +
− −
0.0
2.0
4.0
NADPH (nmol/mg)
G/M
CCCP
−
+
+ +
− −
0.0
1.5
3.0
NADP (nmol/mg)
G/M
CCCP
−
+
+ +
− −
0.0
5.0
10.0
NADPH/NADP
99
Mitochondrial levels of NADH and NAD
+
and their relative ratios were
simultaneously determined in liver mitochondria from WT and NNT mutant mice. As
shown in Fig. 4-4, the addition of glutamate/malate stimulated the NADH production
(Fig. 4-4A) and NAD
+
consumption (Fig. 4-4B), while CCCP led to a decreased NADH
concentration and an increased NAD
+
concentration. The change of NADH levels in
NNT mutant mice in response to substrate was less pronounced than that in WT mice;
this may suggest a compromised response of mitochondrial bioenergetic machinery to
substrate when NNT is absent. Fig. 4-4C shows that NNT mutant mice had a higher
NADH/NAD
+
ratio in liver mitochondria than that in WT mitochondria, under both non-
energized and energized conditions. These differences in NAD pools between these two
lines were abolished after the addition of CCCP, which suggests that these distinctions
are elicited by disparity in the proton-gradient-sensitive NNT activity.
100
Figure 4-4 Mitochondrial NAD(H) pool in liver is perturbed by loss of NNT function.
Liver mitochondria from WT and NNT mutant mice were isolated, and the NADH and
NAD
+
levels were determined under different conditions: (1) without any substrates, (2) with
the addition of 2.4 mM glutamate and 2.4 mM malate, and (3) with 4 μM of CCCP in
addition to glutamate/malate. A. NADH concentrations in liver mitochondria of WT and
NNT mutant mice under different conditions; B. NAD
+
concentrations in liver mitochondria
of WT and NNT mutant mice under different conditions; C. NADH/NAD
+
ratios in liver
mitochondria of WT and NNT mutant mice under different conditions. Bars represent mean
values ± SEM of 4 animals per group (* p < 0.05).
G/M
CCCP
−
+
+ +
− −
G/M
CCCP
−
+
+ +
− −
A
C
B
WT
NNT KO
WT
NNT KO
WT
NNT KO
G/M
CCCP
−
+
+ +
− −
0.0
1.5
3.0
NAD (nmol/mg)
0.0
1.0
2.0
NADH(nmol/mg)
0.0
1.5
3.0
NADH/NAD
101
4.4.3. NADPH production is more affected in liver than in brain
To further compare the difference of the role of NNT in brain and liver, pyridine
dinucleotide levels were measured in brain homogenate of WT and NNT mutant mice.
The NADPH/NADP
+
ratio was 1.35±0.11 in NNT mutant mice and 1.66±0.06 in WT
mice, while the NADH/NAD
+
ratio remained the same in these two lines; hence, in brain,
the NADPH generation was affected by NNT dysfunction but the energy-linked NADH
pool was not affected (Fig. 4-5A). Corresponding to the decline of NADPH/NADP
+
ratio,
the H
2
O
2
generation rate was ~12% higher in the NNT mutant mice (Fig. 4-5B). The
effect of NNT dysfunction on brain mitochondria function was also associated with the
upregulation of the expression of IDH2 in brain, which is another important source of
NADPH (Fig. 4-5C).
102
Figure 4-5. Less effect of NNT dysfunction on mitochondrial redox homeostasis is
associated with upregulation of IDH2 in brain. A. Isolated brain mitochondria from WT
and NNT mutant mice were measured for pyridine dinucleotide levels by HPLC method and
the ratios of NADPH/NADP
+
and NADH/NAD
+
were calculated; B. H
2
O
2
release from
freshly isolated brain mitochondria from WT and NNT mutant mice was monitored for 30
min using Amplex Red fluorescence dye in the presence of 5 mM L-malate and 5 mM L-
glutamate, and the release rate was calculated.; C. IDH2 expression in brain mitochondria of
WT and NNT mutant mice was examined by Western blot (lower panel), and the density of
bands were quantified and normalized to 6-month AL group. Bars represent mean values ±
SEM of 4-5 animals per group (* p < 0.05).
0.0
1.0
2.0
Pyridine Nucleotide Ratio
NADPH/NADP
NADH/NAD
IDH2
NNT KO WT
0
30
60
WT NNT-KO
H
2
O
2
generation rate
(pmol min-1 mg-1)
NNT KO WT
A
C
B
VDAC
0
50
100
150
200
250
Relative levles to WT (%)
103
In liver, the lack of expression of NNT led to significant effects on mitochondrial
function. Similar to the results in liver mitochondria, the NADPH/NADP
+
ratio in
isolated primary hepatocytes was 1.76 in WT mice and 1.24 in NNT mutant mice, while
the NADH/NAD
+
ratio increased from 0.699 in WT mice to 1.231 in NNT mutant mice
(Fig. 4-6A). Meanwhile, the expression levels of IDH2 and glucose-6-phosphate
dehydrogenase (G6PD, primary NADPH producer in cytosol) were not upregulated in
NNT mutant mice (Fig. 4-6B), which possibly explains the more significant effect in
liver induced by NNT dysfunction. Moreover, the expression of major components of the
mitochondrial redox system that require NADPH as electron donor, such as Prx3, Trx2,
and Grx2 were not affected in NNT mutant mice (Fig. 4-6B); this suggests that NADPH
production by NNT is the rate-limiting step of the redox system in liver mitochondria.
104
Figure 4-6. Loss of NNT function in hepatocytes affects mitochondrial NADP(H) and
NAD(H) pools but not on expression of other redox enzymes. A. Primary hepatocytes
were isolated from WT and NNT mutant mice, and the concentrations of pyridine
dinucleotide were measured and the ratios of NADPH/NADP
+
and NADH/NAD
+
were
calculated. Bars represent mean values ± SEM of 4-5 animals per diet per group (* p < 0.05);
B. Lysate of primary hepatocytes isolated from WT and NNT mutant mice IDH2 expression
in brain mitochondria of WT and NNT mutant mice was immunoblotted by antibodies
against enzymes involved in cellular NADPH production (IDH2 and G6PD) and redox status
regulation (Prx3, Trx2 and Grx2), and β-actin was used as loading control.
Prx-3
Trx-2
Grx-2
NNT
NNT KO WT
IDH2
G6PD
β-actin
0.0
1.0
2.0
Pyridine Nucleotide Ratio
NADPH/NADP
NADH/NAD
NNT KO WT
A B
105
4.4.4. Mitochondrial energy capacity is compromised in hepatocytes from NNT mutant
mice
Mitochondria provide most of the energy needed for cellular functions by the
conversion of energy in fuel molecules into ATP through oxidative phosphorylation.
Oxygen consumption rate (OCR) by mitochondria reflects the activity of mitochondrial
bioenergetics, and was found decreased after NNT silencing in PC12 cells (see Chapter
3). Similarly, primary hepatocytes isolated from NNT mutant mice had a substantially
lower basal OCR relative to control cells (Fig. 4-7A); following the addition of
oligomycin, OCR declined in both WT- (from 168 to 119 pmols/min) and NNT-mutant
cells (from 124 to 87 pmols/min) (Fig. 4-7A; 25-50 min), indicating that ATP turnover
was significantly lower in NNT mutant hepatocytes than that in control groups. Maximal
respiratory capacity –measured after the addition of the uncoupler FCCP– was also
substantially lower in cells without NNT (Fig. 4-7A; 50-75 min). The addition of the
complex I inhibitor rotenone in combination with complex III inhibitor antimycin A,
resulted in a further reduction in OCR; the residual O
2
consumption was accounted for by
non-mitochondrial O
2
-consuming pathways (Fig. 4-7A; 75-100 min). To evaluate the
mitochondrial bioenergetic capacity affected by the absent of NNT, OCR data at all of
the time points in both lines were base-lined to their respective basal respiration. As
shown in Fig. 4-7B, the NNT mutant hepatocytes had a much lower increase in maximal
respiration after adding FCCP compared with WT cells (41% versus 7%), indicating a
106
substantial decrease of mitochondrial reserved energy-transducing capacity when NNT is
not present.
Anaerobic glycolysis, the conversion of glucose to lactate, provides a modest amount
of ATP. The extracellular acidification rate (ECAR) was used to detect the glycolytic
activity by monitoring pH changes due to lactic acid generation. There were no
significant differences in ECAR between WT and NNT mutant hepatocytes (Fig. 4-7C).
Compared to the substantial decrease in OCR in NNT mutant hepatocytes, the unchanged
ECAR values suggested that a larger proportion of the total cellular ATP was generated
through glycolysis. An OCR/ECAR ratio analysis was also conducted that confirmed that
ratio of O
2
consumption- and extracellular acidification rate were significantly lower in
NNT mutant cells (Fig. 4-7D).
107
Figure 4-7. Cellular energy metabolism in primary hepatocytes of WT and NNT mutant
mice. Primary hepatocytes were isolated from WT and NNT mutant mice. OCR and ECAR
were determined using Seahorse XF-24 Metabolic Flux Analyzer. Vertical dashed lines
indicate time of addition of mitochondrial inhibitors: oligomycin (4 μM), FCCP (1 μM), or
rotenone+antimycin A (1 μM + 1 μM). A. OCRs in NNT mutant hepatocytes (open circles)
have lower basal rates and maximal rates (after the addition of FCCP) of mitochondrial
respiration than those of WT cells (filled circles). B. The percentage of OCR change relative
to the basal rate of WT and NNT mutant groups was calculated as the value of change
divided by the average value of their respective baseline; C. No significant difference in
ECAR was observed in NNT mutant hepatocytes (open circles), compared to WT groups
(filled circles); D. The OCR/ECAR ratio of in NNT mutant hepatocytes (open circles) is
lower than that of control group (filled circles) at basal conditions and after the addition of
FCCP. *p < 0.05, n = 10. OCR and ECAR readings were normalized to total protein
concentration in each well.
A
C D
B
108
4.4.5. Cytosolic redox signaling induced in NNT mutant hepatocytes
The PI3K/Akt signaling pathway is the classic signaling pathway mediating insulin-
stimulated glucose uptake and metabolism. H
2
O
2
is an important modulator of the
PI3K/Akt signaling, and previous studies suggested that decreased mitochondrial H
2
O
2
production activates this signaling pathway (Chen et al., 2008). To address whether
oxidized redox status induced by NNT dysfunction plays a role in modulating the
PI3K/Akt pathway in liver, we measured phosphorylated (active form) and total levels of
Akt. In NNT mutant hepatocytes, Akt is significantly inactivated compared to WT cells
(Fig. 4-8). GSK3β is a downstream target of Akt by its phosphorylation at Ser
9
, and plays
important roles in regulating glucose homeostasis in liver, and activated GSK3β is also a
pro-apoptotic factor because its activation stimulates the phosphorylation of the anti-
apoptotic members of the Bcl-2 family, Mcl-1 (Maurer et al., 2006; Pap and Cooper,
1998). Consistent with a decreased Akt phosphorylation, phosphorylated GSK3β
(inactive form) levels were significant lower in NNT mutant cells (Fig. 4-8), indicating
an enhanced glycogen synthesis and apoptotic assembly in NNT mutant hepatocytes.
Furthermore, over-activation of GSK3β may lead to energy hypometabolism through the
phosphorylation and inhibition of mitochondrial pyruvate dehydrogenase (PDH)
(Horbinski and Chu, 2005; Hoshi et al., 1996); accordingly, the levels of the
phosphorylated PDH-E1α subunit are higher in NNT mutant mice (Fig. 4-8). The Akt-
GSK3β-PDH pathway could partially contribute to the decline of mitochondrial
respiration shown in Fig. 4-7. Another signaling pathway that could be regulated by
109
mitochondrion-generated H
2
O
2
is JNK pathway, which was found to be activated upon
NNT silencing in PC12 cells (see Chapter 3). Previous work demonstrated that JNK
translocates to the mitochondrion upon activation (bisphosphorylation) and initiates a
signaling cascade that results in phosphorylation and inhibition of PDH (Zhou et al.,
2008; Zhou et al., 2009). In NNT mutant hepatocytes, active JNK1 (the phosphorylated
form) was substantially upregulated compared to that in WT cells (Fig. 4-8), suggesting
the contribution of activation of JNK on PDH phosphorylation and inhibition on
mitochondrial energy-transducing activity. JNK, together with the Akt-GSK3β pathway,
could link the oxidized mitochondrial redox status to impaired mitochondrial energy
metabolism when NNT is absent. These results suggest the important role of NNT in the
regulation of cellular function though mitochondria-generated H
2
O
2
(as second
messenger) and redox-sensitive signaling, and again confirmed the importance of NNT in
maintaining cellular energy levels and redox homeostasis in liver.
110
Figure 4-8. Loss of NNT function induces the modulation of cytosolic redox-sensitive
signaling. A. Primary hepatocytes were isolated from WT and NNT mutant mice and the cell
lysate was immunoblotted with antibodies against phosphorylated form and total levels of
Akt, GSK3β, JNK and PDH; B. Quantification of the levels of p-Akt, p-GSK3β and p-JNK
in NNT mutant hepatocytes normalized to their respective expression in WT cells, and the
relative value of p-PDH/PDH in NNT mutant cell normalized to the raio in WT cells. *p <
0.05, n = 4.
p-Akt
NNT KO WT
p-GSK3β
p-JNK
JNK
β-actin
PDHE1α
p-PDHE1α
Akt
GSK3β
A B
0
50
100
150
200
250
p-Akt p-GSK3β p-JNK p-PDH/PDH
Relative levels to WT (%)
WT
111
4.5. DISCUSSION
The study described in Chapter 3 revealed that knockdown of NNT in PC12 cells alters
the mitochondrial and cellular redox status (GSH depletion and H
2
O
2
production) along
with a compromised mitochondrial bioenergetics by inhibition of mitochondrial
metabolism (including inhibition of PDH and SCOT activities). Activation of redox-
sensitive signaling (JNK) by H
2
O
2
activates the mitochondrion-driven apoptotic pathway
(intrinsic apoptosis) and results in decreased cell viability (Yin et al., 2012b). In this
Chapter, we sought to validate those findings in animal models of NNT dysfunction. By
comparing the regulation of mitochondrial NAD(P) pools by NNT in WT and the NNT
mutant mice, we obtained distinct conclusions regarding brain and liver. In brain, the
NAPDH/NADP
+
ratio was slightly lower in mitochondria from NNT mutant mice when
the substrate was present, but the differences vanished when the proton motive force was
collapsed by CCCP (Fig. 4-1); NADH availability was not affected in NNT mutant mice
(Fig. 4-2). This suggested that in brain of NNT mutant mice, NAPDH production was
moderately affected, but not as severe as observed in NNT-silenced PC12 cells, in which
NNT suppression affected not only NADP but also the energy-linked NAD pool. The
effect of NNT dysfunction in brain was associated with a compensatory upregulation of
IDH2 expression observed in NNT mutant mouse brain (Fig. 4-5). However, in liver, the
impact of NNT on the mitochondrial pyridine dinucleotide pools appeared to be more
substantial than that in brain: first, NAPDH/NADP
+
ratio was lower in NNT mutant mice
112
under both non-energized and energized conditions (Fig. 4-3); second, in energized
mitochondria, the extent of decreased NAPDH/NADP
+
ratio was higher in liver (3.24
fold decrease, NNT mutant versus WT, Fig. 4-3D) than that in brain (1.42 folds decrease,
NNT mutant versus WT, Fig. 4-1D); third, NAPDH and NADP
+
levels and their ratios
were very sensitive to substrate energization in brain of NNT mutant mice (Fig. 4-1), but
not responsive to substrate in liver of the same mice (Fig. 4-3), which suggested that
there are considerable alternative sources of NADPH (such as IDH2) in the brain
mitochondria of NNT mutant mice but these sources are not induced in the liver of these
mice (Fig. 4-6); and fourth, in liver, NNT dysfunction also led to changes in the NAD
pool (Fig. 4-4) that was not affected in brain (Fig. 4-2). Notably, in both brain and liver,
when CCCP was applied, differences between WT and NNT mutant mice in all
parameters mentioned above were abolished; this suggested the importance of the
maintenance of mitochondrial H
+
gradient in not only the energy- transducing capacity
but also the redox homeostasis of the organelles.
Our results are consistent with earlier findings in liver mitochondria that NADPH
generation in these mitochondria after the addition of a NADH-linked substrate (2-
hydroxybutyrate) is less sensitive to oxidation inducer (tert-butylhydroperoxide) than
mitochondria provided with a NADPH-linked substrate (isocitrate, substrate for IDH2)
(Bellomo et al., 1984). Moreover, it is known that NNT expression in liver is about 2
folds of that in brain (Chico et al., 1977), which is associated with higher
NADPH/NADP
+
ratio in liver as we found (Fig. 4-1 and 4-3). Our findings in liver also
113
conformed the notion that NNT tends to generate a high NADPH/NADP
+
ratio at a
relatively low rate, in contrast to reaction catalyzed by IDH2 which generates a lower
NADPH/NADP
+
ratio but at a higher maximal rate (Rydstrom, 2006). Taken these
together, it can be speculated that in liver, the high NADPH/NADP
+
ratio is primarily
contributed by NNT, and the loss of NNT function, either due to NNT mutation (in
C57BL/6J mice) or the collapse of proton motive force (by CCCP), seriously impairs the
ability of mitochondria to build and maintain this high ratio of NADPH/NADP
+
. In the
brain, on the other hand, the less significant effects of NNT dysfunction on NADP pools
can be explained in two folds: (a) a relative low physiological ratio of NADPH/NADP
+
accompanied by lower physiological expression of NNT in brain makes NNT less
important in generating NADPH, and (b) increased expression of IDH2 in the brain of
NNT mutant mice compensates the effects of NNT dysfunction on NADPH production
and redox regulation.
The prominent role of NNT in liver mitochondria led to the further investigations of
NNT function in this tissue at a cellular level in isolated hepatocytes from the WT and
NNT mutant mice. In these hepatocytes, alternative cellular NADPH generators, such as
IDH2 in the mitochondria and G6PD in the cytosol, remained the same levels as in the
WT hepatocytes (Fig. 4-6), making the absent of NNT rather harmful for these cells, as
indicated by the decrease of mitochondrial energy-transducing capacity (Fig. 4-7). This
impairment due to NNT dysfunction is associated with the modulation of several redox-
sensitive signaling pathways including the PI3K/Akt and the JNK pathways. We have
114
demonstrated that in NNT-suppress PC12 cells, activation of JNK is associated with
phosphorylation and inhibition of PDH and commitment impairment of mitochondrial
respiration. Here in primary hepatocytes, we confirmed the activation of JNK in NNT
mutant mice, and we also observed the inhibition of the Akt and the subsequent
activation of its downstream target, GSK3β (Fig. 4-8). The cascades of JNK and Akt-
GSK3β signaling point to a same target in mitochondria, the PDH, which was found
substantially inhibited in NNT mutant cells, in which JNK and GSK3β were activated.
JNK is redox sensitive, and oxidative stress conditions, entailing also enhanced
generation of mitochondrial H
2
O
2
, result in its activation (Nemoto et al., 2000; Zhou et
al., 2008). H
2
O
2
may act at multiple levels to activate JNK: dissociation of thioredoxin
from the ASK-1 complex (Saitoh et al., 1998), disruption of the glutathione transferase
(GT)-JNK complex (Adler et al., 1999), or inhibition of MAPK phosphatase activity
(Foley et al., 2004). Mitochondrial H
2
O
2
is also involved in the regulation of the
PI3K/Akt route of insulin/IGF-1 signaling. Akt inhibits GSK3β upon phosphorylation at
Ser
9
and thereby protects cell against apoptosis (Pap and Cooper, 1998). It is still not
clear that how H
2
O
2
levels regulate Akt activity. Interestingly, previous studies show that
exogenous H
2
O
2
activates the PI3K/Akt signaling pathway through oxidation of specific
cysteine residues that promote tyrosine autophosphorylation of the insulin receptor
(Storozhevykh et al., 2007) and through the inhibition of phosphatases (PTEN and
PTP1B) involved in the insulin receptor substrate (IRS) node of insulin signaling upon
oxidation of critical cysteines to disulfides (Mahadev et al., 2001; Salmeen et al., 2003).
115
In this study, increased H
2
O
2
release from mitochondria due to NNT dysfunction actually
led to inactivation of Akt, which is against the above-mentioned mechanism. However, in
the NNT mutant mice, H
2
O
2
that signals to Akt is endogenous and its concentration is
much lower than concentration of exogenous H
2
O
2
used in other studies. And our results
were consistent with another study, in which decreased mitochondrial H
2
O
2
generation by
Prx3 overexpression activates Akt signaling (Chen et al., 2008). Hence, regulation of
PI3K/Akt pathway by redox modulation appears to be more comprehensive than simply
“activation” or “inhibition”. It is likely dependent on the local concentration of H
2
O
2
. In
addition, JNK signaling inhibits PI3K/Akt signaling (Karpac and Jasper, 2009); a likely
mechanism entails the phosphorylation of the insulin receptor substrate-1 (IRS-1) at
Ser
307
, leading to the inhibition of the insulin-promoted tyrosine phosphorylation of IRS
(Aguirre et al., 2000). It is possible that the activation of JNK inhibits Akt signaling and
this inhibition may overcome the direct activation by H
2
O
2
, and results in a decreased
activity of Akt. Actually, JNK activity was found elevated in peripheral tissues including
liver in multiple models of obesity and insulin resistance (Hirosumi et al., 2002), and the
inhibition of the insulin pathway by JNK is a major factor in the etiology of type II
diabetes (Karpac and Jasper, 2009). Our results could explain the diabetic phenotype
found in the NNT-absent C57BL/6J mice, and the lack of phenotype in brain in the same
line. Nevertheless, this study strengthens the notion that disruption of electron flux from
fuel substrates to redox components imposed by NNT dysfunction not only alters
mitochondrial redox status but also impair bioenergetic capacity and other cellular
116
processes through redox-sensitive signaling. The communication between mitochondria
and other components of the cell therefore establishes a regulatory devise that controls
cellular energy levels and the redox environment.
117
CHAPTER 5
CONCLUSIONS
Mitochondrial dysfunction is an inherent feature of brain aging and is involved in the
development of neurodegenerative diseases. As the powerhouses of the cell and effective
sources of cellular H
2
O
2
, mitochondria play a central role in the biology of aging due to
(a) the decline of basal metabolic rate and (b) the impairment of redox signaling and
transcription that occurs in aging. As such, mitochondria have been often regarded as the
pacemaker of tissue aging. Age-related changes in energy production and redox status
cannot be viewed as independent variables, but rather as an interdependent relationship
reflected in the mitochondrial energy-redox axis. The communications between
mitochondria and the rest of the cell by redox-sensitive signaling establish a master
regulatory devise of the energy levels and the redox environment of the cell.
Mitochondrial NNT provides ultimate reducing equivalents to the redox systems using
the proton gradient as the driving force and NADH as the electron donor. We thus
hypothesized that NNT provide a critical link between the mitochondrial metabolic
function and redox homeostasis, and the modulation of NNT is critical for mitochondrial
and cellular function that is compromised with aging. This hypothesis encompasses a
118
disruption of electron flow between the energy and redox component of the mitochondria
(i.e., disruption of the energy-redox axis), perturbations of mitochondrial redox
homeostasis, a collapse of mitochondrial bioenergetics, and alterations of cell function
through redox-sensitive signaling pathways.
In specific aim 1, functional changes of the mitochondrial energy-redox axis in
brain, entailing components involved in mitochondrial bioenergetics and redox status
maintenance, in the aging process and their modulation by short-term CR were
determined. It was demonstrated that NNT function and NADPH availability decreased
with age in brain, accompanied by compromised mitochondrial bioenergetic capacity and
altered redox status. 2-month of CR significantly reversed the age-dependent impairment
of the axis by (a) upregulating ketone body metabolism, (b) increasing OXPHOS
capacity, (c) maintaining redox equilibrium, and (d) enhancing NNT function and
NADPH supply. Notably, the effects of short-term CR on the modulation of
mitochondrial function were only observed in old rats, but not in young animals.
In specific aim 2, the role of NNT in regulating mitochondrial redox status and energy
metabolism, as well as its further impact on cellular function was investigated in a NNT-
suppressed neuronal cell model. Silencing of NNT in PC12 cells resulted in decreased
cellular NADPH levels, altered redox status of the cell in terms of decreased GSH/GSSG
ratios and increased H
2
O
2
levels, thus leading to an increased redox potential. Increased
H
2
O
2
levels activates JNK, followed by a decrease of mitochondrial oxygen consumption
in association with (a) inhibition of mitochondrial PDH and SCOT activity, (b) reduction
119
of NADH/NAD
+
ratio, (c) decline of mitochondrial membrane potential, and (d) decrease
of ATP levels. Active JNK also induced intrinsic apoptosis, and resulted in decreased cell
viability.
In specific aim 3, the function of NNT in linking the mitochondrial energy status to the
redox environment was evaluated in brain and liver of a NNT mutant mouse model. It
was demonstrated that NNT regulates mitochondrial NADP(H) levels in an energy-
sensitive fashion in brain and liver. In liver, the NADP(H) pool is more affected by loss
of NNT activity, and is associated with an altered NAD(H) pool and impaired energy-
transducing capacity. It was confirmed that redox-sensitive signaling pathways including
JNK and Akt are involved in mediating the signal initiated from altered mitochondrial
redox status to metabolic function. And the diminished impact of NNT dysfunction in
brain could be due to an upregulation of IDH2.
Taken together, these studies strengthens the notion that oxidized cellular redox state
and decline in cellular bioenergetics in aging and age-related diseases cannot be viewed
as independent events, but rather as an interdependent relationship coordinated by the
mitochondrial energy-redox axis, a dual pronged approach to assess the changes in
mitochondrial function. The transmission of electrons from fuel molecules to redox
components by NNT is crucial not only for the integrity of the axis itself but also for
cellular functions with the involvement of mitochondrial second messengers and redox-
sensitive signaling.
120
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Abstract (if available)
Abstract
The mitochondrial energy-transducing capacity is essential for the maintenance of neuronal function: impairment of energy metabolism and redox homeostasis –integrated in the mitochondrial energy-redox axis– is a hallmark of brain aging and is accentuated in the early stages of neurodegenerative diseases. The energy component of the axis entails the formation of reducing equivalents (NADH) and their flow through the respiratory chain with consequent electron leak to generate O₂.⁻ and H₂O₂. The redox component of the axis entails the removal of H₂O₂ by NADPH-dependent, thiol-based systems. Mitochondrial NADPH generation is largely dependent on nicotinamide nucleotide transhydrogenase (NNT) that catalyzes the reduction of NADP⁺ to NADPH utilizing the proton gradient as the driving force and NADH as the electron donor, thereby linking the energy- and redox components of the axis. ❧ The hypothesis to be tested is that NNT activity is critical for the maintenance of the mitochondrial energy-redox axis and is compromised during aging leading to mitochondrial dysfunction and loss of cellular redox homeostasis. This hypothesis was tested by three specific aims that encompassed three different experimental models. ❧ I have shown that NNT dysfunction impairs cellular redox homeostasis and energy metabolism in PC12 cells. Knockdown of NNT results in decreased cellular NADPH supply, increased H₂O₂ levels, and increased redox potential. Altered redox status further leads to the impairment of mitochondrial bioenergetic function through the activation of redox-sensitive c-Jun N-terminal kinase (JNK) signaling and concomitant inhibition of pyruvate dehydrogenase. Active JNK also initiates mitochondrion-dependent intrinsic apoptosis after NNT suppression. ❧ The role of NNT in linking the mitochondrial energy status to the redox environment was further investigated in brain and liver of C57BL/6J mice –essentially NNT knockout mice– that show impaired glucose tolerance independent of obesity. The results showed that NNT regulates mitochondrial NADPH levels in an energy-sensitive manner in both tissues but with the NADP pool in the liver more affected. In primary hepatocytes of C57BL/6J mice, loss of NNT also leads to impaired mitochondrial energy-transducing capacity, a process related by the co-regulation of redox-sensitive signaling pathways. The diminished impact of NNT in brain was ascribed to the upregulation of another mitochondrial NADPH-generating enzyme, isocitrate dehydrogenase-2. ❧ The functional changes of the mitochondrial energy-redox axis in brain were further characterized in an aging model and the effects of short-term caloric restriction (CR) were assessed. It was shown that the substrate supply (pyruvate and ketone bodies) and mitochondrial energy-transducing capacity are enhanced in 26 month-old rats following short-term CR
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Yin, Fei
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The mitochondrial energy – redox axis in aging and caloric restriction: role of nicotinamide nucleotide transhydrogenase
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Doctor of Philosophy
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Pharmaceutical Sciences
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energy metabolism
hydrogen peroxide
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nicotinamide nucleotide transhydrogenase
redox signaling