University of Southern California Dissertations and Theses

Protein post-translational modifications in the cell's redox and energy states

(USC Thesis Other)

Protein post-translational modifications in the cell's redox and energy states

PDF

Download

Open document

Flip pages

Download a page range

Download transcript

Copy asset link

Request this asset

Transcript (if available)

Content

PROTEIN POST-TRANSLATIONAL MODIFICATIONS IN THE CELL’S REDOX
AND ENERGY STATES

by

Jerome Vincent Garcia

_____________________________________________________________________

A Dissertation Presented to the
FACULTY OF THE GRADUATE SCHOOL
UNIVERSITY OF SOUTHERN CALIFORNIA
In Partial Fulfillment of the
Requirements for the Degree
DOCTOR OF PHILOSOPHY
(MOLECULAR PHARMACOLOGY AND TOXICOLOGY)

May 2007

Copyright 2007 Jerome Vincent Garcia
ii

Dedication

To my wife Sheila and my parents, Bernie and Agnes Garcia, for their unconditional
love and support.
iii
Acknowledgements

I would like to thank my mentor Dr. Enrique Cadenas for his guidance and support
throughout the years. Thank you Dr. Derick Han and Li-Peng Yap, for your help and
encouragement. My education would not have been possible without the comfortable
working environment created by former and current laboratory colleagues, Dr.
Siranoush Sharzhrad, Dr. Havan Nguyen, Dr. Dominique Nguyen-Wascher, Dr. Allen
Chang, Dr. Qiong Zhang, Yue Tang, Philip Lam, and Ryan Hamilton.

In addition, I would like to thank all my committee members: Dr. Alex Sevanian, Dr.
Dr. Ronald Alkana, and Dr. Tzung Hsiai for their insightful suggestions.
iv
Table of Contents

Dedication ii
Acknowledgements iii
List of Tables v
List of Figures vi
Abstract ix
Chapter 1: Introduction 1
Hypothesis 3
Specific Aims 3
Background 4
Significance 13
Synopsis 15
Chapter 2: Regulation of redox status in brain mitochondria 18
Chapter 3: Synergistic action of dopamine and nitric oxide in neuronal injury 54
Chapter 4: Critical post-translational mitochondrial protein modifications 80
Chapter 5: Conclusion 113
Chapter 6: Future Perspectives 117
References 119
v
List of Tables

Table I: GSH concentrations subsequent to different mitochondrial
isolation methods 27

Table II: Potentiation of nitric oxide-mediated damage by dopamine in
PC12 cells 73

Table III: ONOO- induced brain mitochondrial dysfunction by nitric oxide
and dopamine autoxidation 73

Table IV: LC/MS/MS analysis of aconitase amino acid modifications 98

Table V: ATPase activity: NADH consumption 107

vi
List of Figures

Figure 1: Cellular redox status and function 17
Figure 2: NAD(P)H formation upon complex I substrate (glutamate/malate
-G/M) supplementation. 30

Figure 3: GSH and GSSG concentration of (non)energized liver mitochondria 31

Figure 4: GSH and GSSG concentration of (non)energized liver mitochondria 33

Figure 5: Substrate dependent GSH formation in brain mitochondria 35

Figure 6: Similar GSH and GSSG concentrations in energized liver
mitochondria (DC) versus non-energized mitochondria exposed to a
reducing agent 37

Figure 7: Similar GSH and GSSG concentrations are achieved once the liver
mitochondria (DPG) are energized versus non-energized mitochondria
exposed to a reducing agent 38

Figure 8: Similar GSH and GSSG concentrations are achieved once the brain
mitochondria (DPG) are energized versus non-energized mitochondria
exposed to a reducing agent 40

Figure 9: Substrate-induced protein sulfhydryl recovery (deglutathionylation) of
glutathione-protein adducts and identification of immunoprecipitated
proteins by LC/MS/MS 42

Figure 10: H
2
O
2
induced GSH depletion and GSSG formation in rat brain
mitochondria 44

Figure 11: H
2
O
2
-induced GSH depletion and GSSG formation in rat brain
mitochondria supplemented with complex I substrates and ADP 45

Figure 12: GSH formation depends on substrate availability not electron flow
through the ETC 50

vii
Figure 13: Determination of O
2
.-
, from dopamine oxidation, using electron
paramagnetic resonance 59

Figure 14: ONOO
-
formation by dopamine in the presence of
.
NO 60

Figure 15: Simplified scheme of the dopamine oxidation process and its
byproducts 62

Figure 16: Dopamine-o-semiquinones decay during dopamine autoxidation
in RCR 63

Figure 17: Dopaminochrome formation and oxygen consumption during
dopamine autoxidation 66

Figure 18: Effects of GSH on oxygen consumption, and formation of
dopamine-o-semiquinone, and neuromelanin 68

Figure 19: Proposed mechanism of dopamine recycling/reduction of
dopamine semiquinone by GSH 69

Figure 20: Detection of the glutathionyl radical using electron paramagnetic
resonance 71

Figure 21: GSH depletion associated with dopamine oxidation 79

Figure 22: ONOO
–
-mediated inactivation of aconitase: Effect of citrate 91

Figure 23: Effect ONOO
–
on the Fe-S cluster in the active site of aconitase:
EPR analysis 92

Figure 24: Detection of an immobilized DMPO/cysteinyl radical-protein
radical adduct after ONOO
–
treatment 94

Figure 25: Loss of aconitase thiols and nitrotyrosine formation following
ONOO
–
treatment 97

viii
Figure 26: MS/MS spectra for aconitase tryptic peptides carrying an
oxidative/nitrative modification 100

Figure 27: Structure of aconitase and amino acid modifications elicited by
ONOO
–
102

Figure 28: Effect of GSSG on aconitase activity and aconitase protein thiols.
Glutathionylation of aconitase 105

ix
Abstract

The increased generation of reactive oxygen and nitrogen species and
glutathione depletion creates profound changes in the redox status of the cell and has
been implicated in the progression of neurodegenerative diseases such as Parkinson’s
disease. Redox regulation has become exceedingly important in understanding the
cellular adaptation to oxidative stress. However, it remains unclear how changes in
the cellular and mitochondrial redox environment affect redox sensitive protein
function during exposure to
.
NO and dopamine and the consequences of such changes
with respect to mitochondrial function and redox signaling (achieved through post-
translational modifications).
The work in this dissertation demonstrates that glutathionylation of
mitochondrial proteins occurs during oxidative stress. This modification functions as a
store of GSH and is released in response to changes in mitochondrial redox and energy
status. Protein sulfhydryl recovery (de-glutathionylation) of mitochondrial proteins in
energized mitochondria increased the GSH buffering capacity of mitochondria against
physiological and non-physiological concentrations of hydrogen peroxide (H
2
O
2
), thus
preventing drastic changes in mitochondrial redox status. This is exemplified using a
Parkinson’s disease model with NO and dopamine as modifiers of the redox
environment. The reaction between NO and dopamine oxidation produces ONOO-, a
powerful oxidant that leads to a 72% inhibition of mitochondrial respiration in isolated
mitochondria and 71% inhibition in PC12 cells.
x
Additionally, we identified aconitase and ATP synthase as targets of protein
post-translational modifications. Nitration of aconitase by ONOO
-
resulted in a dose-
dependent decrease in aconitase function. In the presence of citrate, a substrate for
aconitase, a 66-fold higher concentration of ONOO- was required for half-maximal
inhibition. Glutathionylation of aconitase and ATP synthase also resulted in an
inhibition of enzyme function. De-glutathionylation of ATP synthase resulted in a
221% change in activity. It is well established that damage to proteins under
conditions of oxidative and nitrosative stress is highly specific, and plays an important
role in many diseases. Characterization of post-translational modifications of
mitochondrial proteins provides a mechanism for
.
NO and dopamine elicited damage
to mitochondrial function leading to apoptosis of dopaminergic neurons in Parkinson’s
disease.
1
Chapter 1: Introduction

The increased generation of reactive oxygen and nitrogen species and
glutathione depletion has been implicated in the progression of neurodegenerative
diseases such as Parkinson’s disease (PD). From a neuropathological view, PD is
defined as the selective degeneration of pigmented dopamine synthesizing/storing
(dopaminergic) neurons found in the substantia nigra pars compacta (Forno 1996).
Although the etiology of PD remains a longstanding debate, it has been well
established that significant depletion of total glutathione levels is observed in the
substantia nigra of early idiopathic PD patients (Chinta and Andersen 2006).
Glutathione (GSH) is a major antioxidant and redox regulator in cells and
functions namely as a buffer against reactive oxygen and nitrogen species. Increased
generation of reactive oxygen and nitrogen species from different sources or
mechanisms, i.e. inflammation, dopamine autoxidation, and mitochondrial oxidative
stress, can regulate the redox status of dopaminergic neurons through the modulation
of GSH and GSSG pools. This in turn affects neuronal function through (i) altering
the redox status of the cell through diminishing the buffering capacity of the
GSH/GSSG couple, (ii) redox modulation of protein sulfhydryl groups reflected by
thiol/disulfide exchange and (iii) mitochondrial function. Redox regulation has
become exceedingly important in understanding cellular adaptation to oxidative stress.
Individual signaling and control events occur through discrete redox circuits or
pathways rather than through global mechanisms where there is a general disruption of
thiol/disulfide balance. Thus it was proposed recently by Dean Jones, that oxidative
2
stress should be viewed as a disruption of redox signaling (Jones 2006). Alterations
in normal redox balance and energy status can alter signaling pathways and protein
function through protein post-translational modifications (e.g. nitration and
glutathionylation) and should be considered as early events in neurodegenerative
disorders and also in PD. This notion is tenable for:
1) Nitric oxide (
.
NO) has been reported to be increased in PD (Iravani, Kashefi et al.
2002; Zhang, Dawson et al. 2006).
.
NO, generated by the inducible form of nitric
oxide synthase (iNOS) in glial cells (Liberatore, Jackson-Lewis et al. 1999) or the
neuronal form (nNOS), participates in the cascade of events leading to the
degeneration of dopamine-containing neurons.
2) The selective loss of dopaminergic neurons has focused much attention on the
dopamine molecule itself as a potential mediating factor in PD. Dopamine,
because of its catechol structure can easily oxidize to form superoxide anion (O
2
.-
),
it’s dismutation product hydrogen peroxide (H
2
O
2
), and neuromelanin, a dark
pigment found in dopaminergic neurons (Herlinger, Jameson et al. 1995).
3) There is evidence that GSH levels, a component of the cell’s antioxidant defense,
are decreased in the substantia nigra in Parkinson’s disease (Perry, Godin et al.
1982; Sian, Dexter et al. 1994; Bharath, Hsu et al. 2002). Glutathione depletion is
the earliest reported change in the substantia nigra and the magnitude of
glutathione depletion appears to parallel the severity of the disease (Chinta and
Andersen 2006).
4) PD cybrids (engineered cell lines containing PD mitochondria) have implicated
mitochondria as a major component in the selective death of dopaminergic
3
neurons, as these cells are more susceptible to methyl phenylpyridinium (MPP
+
),
a toxin known to cause PD development (Hodaie, Neimat et al. 2007). The role of
mitochondria is further illustrated by experiments using rotenone, a mitochondrial
complex I inhibitor. Inhibition of complex I by rotenone resulted in the loss of
dopaminergic neurons in the substantia nigra (Greenamyre, Sherer et al. 2001;
Greenamyre, Betarbet et al. 2003). Administration of animals treated with MPP
+

with glutathione ethyl ester protected against oxidative stress or chronic
mitochondrial impairment (Zeevalk, Manzino et al. 2007).

HYPOTHESIS - The hypothesis to be tested is that changes in cell redox status –
elicited by
.
NO overproduction and dopamine autoxidation – affect mitochondrial
and cell function through specific and critical protein modifications leading to
apoptotic cell death.

SPECIFIC AIMS
a) Regulation of redox status in brain mitochondria: Determine the role of
glutathionylation, a new protein post-translational modification, in the
mitochondria by (i) assessing changes in GSH and GSSG concentrations in the
mitochondria (ii) glutathionylation patterns and (iii) the protective effects of
mitochondrial GSH against oxidative stress.
b) Synergistic action of dopamine and nitric oxide in neuronal injury: Dopamine
and nitric oxide represent an abundant source of oxidants that can deplete GSH.
Their potential synergistic effects on mitochondrial and cellular function were
4
investigated by (i) first identifying the reaction products of
.
NO and dopamine,
(ii) the effects of dopamine,
.
NO and its reaction products on mitochondrial and
cellular function and (iii) the potential protective effects of GSH against
dopamine,
.
NO and it’s reaction products.
c) Critical post-translational mitochondrial protein modifications: Sites,
mechanisms, and consequences of nitration, cysteine oxidation and
glutathionylation of mitochondrial proteins exposed to distinct redox
environments were assessed using proteomics approach utilizing LC/MS/MS
approach. Specific experiments were employed to determine changes in
individual protein function after post-translational modifications.

BACKGROUND

The cellular redox environment: An indicator of cellular health
a) Definitions and circuits
The redox status of the cell is associated with the cell’s progression through its
life cycle: as the redox environment becomes increasingly oxidative, the cell
progresses from proliferation to differentiation to apoptosis and necrosis (Schafer and
Buettner 2001). Work carried out in our laboratory by Antunes et. al. (Antunes and
Cadenas 2001) provided a quantitative analysis of the cellular steady-state levels of
H
2
O
2
and thresholds linked to proliferation, apoptosis, and necrosis.
The redox status of the cell is best defined by the GSH/GSSG redox couple
(Han, Hanawa et al. 2006). Glutathione (GSH) is most abundant non-protein thiol (~1-
5
11 mM) (Han, Canali et al. 2003). The protective effects of GSH can be attributed to
the thiol (SH) containing side chain of cysteine which allows formation of protein
disulfide bonds and the removal of electrophiles and oxidants (DeLeve and Kaplowitz
1991; Hammond, Lee et al. 2001), resulting in a depletion of GSH and the production
of GSH disulfide (GSSG), the oxidized from of GSH. Its concentration and
antioxidant properties are the reasons that this compound best defines the redox status.
GSH is synthesized in the cytosol from glycine, glutamate, and cysteine in a two-step
process catalyzed by γ-glutamylcysteine synthetase and GSH synthase. Although its
synthesis is primarily within the cytosol, distinct pools of GSH are created through the
sub cellular distribution within organelles, such as mitochondria. The oxidation
product of GSH, glutathione disulfide (GSSG), is reduced back to GSH by glutathione
reductase, a NADPH- dependent enzyme ubiquitously distributed throughout tissues.
Under non-oxidative or nitrosative stress conditions, the concentration of GSSG is
negligible at 1/100
th
of the total GSH pool (Schafer and Buettner 2001). As the
concentration of GSH far exceeds any other redox active couple, measurement of GSH
and or GSSG levels have been used to define the redox environment of the cell,
calculated by the Nernst equation, E
hc
= E
o
– RT/nF ln Q (where R is the gas constant,
T is the temperature (Kelvins) and F is the faraday constant, n is the number of
electrons exchanged and Q is the mass action exchanged). Accordingly, the redox
potential of the GHS/GSSG redox couple at 25
o
C, ph7.0 is defined as E= -240 –
(59.1/2) log([GSSG]/[GSH]
2
) (Schafer and Buettner 2001). This allows comparison
of the reducing force available from the latter redox couple with respect to other redox
couples. The redox potential is not only dependent upon the ratio of GSH to GSSG but
6
also the absolute concentration of GSH, which represents the reducing capacity of the
cell. Generation of ROS and RNS can lead to the conversion of GSH to GSSG or
GSNO [GSH +
.
NO → GSNO]. Thus, changes of ROS and RNS production can lead
to significant changes of the redox environment and affect the reducing capacity of the
cell. Consideration of the Nernst potential provides direct quantifiable information on
the pro-oxidant/antioxidant balance. The Nernst potential provides an index of the
reducing force and thus, calculating the Nernst potential provides a means to evaluate
oxidative stress, the direction of electron flow and redox signaling in different
compartments (Jones 2006).
b) Compartmentalization of mitochondrial GSH pools: Unique redox control
The sub cellular compartments differ in their redox states where the most
reducing to most oxidizing are mitochondria > nuclei > cytoplasm > endoplasmic
reticulum > extracellular space. Due to the relatively alkaline pH of the mitochondria,
mitochondrial protein thiols (-SH) are exceptionally sensitive and vulnerable to
inhibition of activity via oxidation, therefore GSH concentrations must be high in this
organelle to protect sensitive protein thiols. Unfortunately, mitochondria lack the
machinery to perform de novo synthesis of GSH and depend on the cytosol pool as its
source of GSH. Transport of GSH into mitochondria can be stimulated by energizing
mitochondria through specific transporters (Hansen, Go et al. 2006). The lack of
machinery to perform de novo synthesis of GSH and the inability to export its
oxidized form, GSSG by the mitochondria, gives significance to the glutathione
reductase enzyme, an NADPH dependent enzyme that reduces GSSG back to GSH.
This enzyme is important in maintaining mitochondrial GSH content. Mitochondrial
7
NADPH:NADP
+
ratio remains high due to NADP-dependent isocitrate
dehydrogenase (Jo, Son et al. 2001) and by a transhydrogenase that drives electrons
from NADH to NADP
+
(Bizouarn, Fjellstrom et al. 2000) to ensure efficient reduction
of GSSG to GSH.
Evidence amassed from numerous studies in various cell types exposed to
cytotoxic insults have shown that mitochondrial GSH levels, as opposed to cytosolic
GSH, play a far more significant role in determining cellular function and viability
(Fernandez-Checa, Garcia-Ruiz et al. 1991; Dhanbhoora and Babson 1992; Uhlig and
Wendel 1992; Shan, Jones et al. 1993; Colell, Garcia-Ruiz et al. 1998; Wullner,
Seyfried et al. 1999; Colell, Coll et al. 2001). It is also well established that although
mitochondrial and cytosolic pools of GSH are similar in concentration, they are
independent of each other. This allows GSH redox independence of each of the
respective compartments. Mitochondrial thiols are important targets of oxidant
induced apoptosis and necrosis (Hansen, Go et al. 2006). Taken into consideration
that mitochondria themselves are the most important source of cellular superoxide and
hydrogen peroxide, contain highly oxidizable thiols, and is central in mitochondrion
driven apoptosis (Cadenas 2004), understanding the modulation of mitochondrial GSH
pools and mitochondrial redox status is of paramount importance with respect to
cellular health and apoptosis.
A plethora of mitochondrial proteins have been demonstrated to be post-
translationally modified during oxidative stress such as the nitration of pyruvate
dehydrogenase during ischemic reperfusion (Richards, Rosenthal et al. 2006).
Additionally, complex I of the mitochondrial respiratory chain is persistently
8
glutathionylated (formation of mixed disulfides) under conditions of oxidative stress
(Beer, Taylor et al. 2004). The GSH/GSSG redox couple dynamically regulates
protein function through the reversible formation of mixed disulfides between protein
cysteine sulfhydryls and GSH in a process termed glutathionylation. Mixed disulfide
formation can occur mainly through three mechanisms. (i) thiol disulfide exchange
between protein sulfhydryl groups and GSSG (ii) GSH reduction of protein sulfenic
acids, and (iii) the nucleophilic attack of thiolate on the S-NO bond of GSNO (Klatt
and Lamas 2000; Schafer and Buettner 2001). Cysteinyl groups can be oxidized to
form sulfineic acid derivatives, which can lead to irreversible loss of protein function.
Cysteinyl groups can also undergo covalent modification by
.
NO through
transnitrosylation reactions with GSNO or nitrosylation (N
2
O
3
). In many instances
where S-nitrosylation has been described, the protein cysteines are also oxidized
resulting in the subsequent formation of disulfide bonds (S-thiolation) (Martinez-Ruiz
and Lamas 2004). In vivo, the most abundant protein thiol is glutathione, which exist
in milimolar concentration in the cytoplasm. Hence, S-nitrosylation is likely to
promote S-glutathionylation. The formation of mixed disulfides is expected in part to
reflect the redox status of the cell or organelle (Schafer and Buettner 2001). The end
result of these redox-induced modifications are typically an inhibition of protein
activity that may lead to cell death, therefore it is crucial to characterize different
modifiers of the redox environment.

9

Modifiers of the redox environment in neurodegenerative disorders
a) Nitric Oxide overproduction and dopamine autoxidation
Nitric oxide (
.
NO) has been reported to be increased in PD (Iravani, Kashefi et
al. 2002; Zhang, Dawson et al. 2006). Neuroinflammation in both animal and humans
with PD is a ubiquitous finding (Whitton 2007). During neuroinflammation, activated
microglia, which are the resident immune cells of the brain, generate a whole slew of
neurotoxic factors that include cytokines and reactive oxygen and nitrogen species. Of
particular interest is the increased micromolar concentrations of
.
NO and the long-
lasting effects produced by the inducible form of nitric oxide synthase (iNOS) in glial
cells (Liberatore, Jackson-Lewis et al. 1999) or the neuronal form (nNOS). Although
a necessary component of the body’s defense system, neuroinflammation can have
detrimental effects when over stimulated. Additionally,
.
NO has been shown to play a
role in the selective degeneration of dopaminergic neurons associated with
Parkinson’s disease. Although the role of neuroinflammation (
.
NO) and dopamine
individually has been extensively studied in the development of PD, little has been
done to study their interactions. Reiterating the facts stated above, dopamine is
released in the presence of
.
NO during PD and their interactions need to be elucidated
and determined if they are an important component of this disease.
Dopamine is an archetypical neurotransmitter. Synthesis of dopamine from the
amino acid tyrosine is a two-step process: the first is catalyzed by tyrosine
hydroxylase, resulting in the formation of L-DOPA and the second is the deamination
of L-DOPA, catalyzed by DOPA decarboxylase, to form dopamine. Dopamine is than
10
actively pumped (ATP dependent) into storage vesicles, where it stays until released
into the synaptic space. These vesicles not only provide a physical barrier to protect
dopamine from enzyme degradation they also favor the protonated over the de-
protonated form of dopamine. The de-protonated dopamine is more prone to
(auto)oxidation, therefore to displace the equilibrium (pKa =8.9) towards the
protonated species, the storage vesicles maintain a pH of 5.5, where the ratio of
protonated:de-protonated is 3000:1. Once dopamine is released to the synaptic cleft or
cytosol the pH shifts to 7.4, and the ratio changes to 30:1. With respect to its
molecular structure, dopamine is an unstable and oxidizable molecule due to its
catechol moiety. The process of auto-oxidation entails two one-electron transfer steps:
the first and rate-limiting step involves the conversion of dopamine to its semiquinone.
This process can be accelerated by transition metal ions, certain enzymes (xanthine
oxidase, prostaglandin H synthase, and tyrosinase (Stokes, Hastings et al. 1999) but
more importantly by
.
NO (Rettori, Tang et al. 2002). Previous work done in our
laboratory demonstrated that
.
NO released by a steady NO donor, DETA-NO, into an
aerobic solution of dopamine resulted in a 20-fold increase in a EPR dopamine
semiquinone signal intensity (Rettori, Tang et al. 2002), suggesting that
.
NO
accelerates dopamine oxidation and potentially produces O
2
.-
at a faster rate
(Herlinger, Jameson et al. 1995).
Additionally the presence of
.
NO during dopamine autoxidation may lead to
peroxynitrite (ONOO
-
) formation. ONOO
-
is formed by the rapid reaction between
.
NO and O
2
.-
(reaction k
2
= 1.9 x 10
10
M
-1
s
-1
). This reaction takes precedent over the
disproportionation of O
2
.-
to hydrogen peroxide by superoxide dismutase, as the
11
reaction occurs at ~10-fold slower rate (2.3 x 10
9
M
-1
s
-1
). ONOO
-
is an oxidant and
can partake in many chemical reactions within the cell such as modulating the redox
status of the cell, oxidize lipids, and modulate protein function through chemical post-
translational modifications of amino acids such as nitration of tyrosine to form
nitrotyrosine. It has been demonstrated that nitrotyrosine accumulates in the substantia
nigra of patients with Parkinson’s disease (Good, Hsu et al. 1998), which has led to
the hypothesis that ONOO
-
might be a key oxidant in the causation of cellular
oxidative damage, which leads to neurodegeneration (Torreilles, Salman-Tabcheh et
al. 1999). Increased
.
NO generation through MPTP activation of iNOS in
dopaminergic neurons resulted in cellular damage (Liberatore, Jackson-Lewis et al.
1999). These findings are in agreement with the hypothesis that
.
NO released by glial
cells or within the dopaminergic neurons themselves, may participate in the cascade of
events inherent in PD either directly or through its reaction product ONOO
-
.
b) Glutathione in neurodegeneration with emphasis of Parkinson’s disease
Patients diagnosed early with PD exhibited a significant decrease of total GSH
(GSH + GSSG) in the substantia nigra (Perry and Yong 1986). Although GSH is only
one part of the antioxidant defense, its depletion tends to coincide with the severity of
the disease as well as precedes both substantia nigral degeneration and the loss of
mitochondrial complex I activity through post translational modification (Jenner 1998;
Chinta and Andersen 2006). Work published by Chinta et.al demonstrated clearly that
chronic glutathione depletion leads to inhibition of mitochondrial complex I activity
through a ONOO
-
mediated event, which is reversible by a thiol reducing agent.
Additionally, chronic GSH depletion lead to an increase in intracellular NO levels and
12
S-nitrosation of mitochondrial proteins. Complex I inhibition due to chronic GSH
depletion was reversible upon replenishment of GSH levels to normal (Chinta and
Andersen 2006). As mentioned before, treatment of MPP
+
exposed rats with
glutathione ethyl ester could protect against oxidative stress or chronic mitochondrial
impairment (Zeevalk, Manzino et al. 2007). Thus, therapeutics towards maintenance
of GSH levels within the brain would be beneficial in terms of PD.
c) Mitochondrial dysfunction and oxidative stress’ involvement in Parkinson’s disease
There are several lines of evidence implicating oxidative/nitrosative stress and
mitochondrial dysfunction in a variety of cellular pathologies, including
neurodegeneration. The role of mitochondria in cellular pathology emanates from the
unique features of these organelles, such as oxidative phosphorylation, and generation
of reactive oxygen species generalized below:
1) Progressively increasing concentrations of
.
NO are required to observe the
following changes in mitochondria: reversible binding of cytochrome oxidase
(complex IV), inhibition of the bc
1
segment (complex III) of the respiratory
chain, and oxidation of the mobile carrier ubiquinol. The second effect is
similar to that elicited by antimycin A and leads to H
2
O
2
formation by
mitochondria; likewise ubiquinol oxidation by
.
NO is an effective source of
O
2
.-
, which, depending on steady-state levels of reactants, may dismutate to
H
2
O
2
(via Mn-superoxide dismutase-catalyzed reaction) or ONOO
-
(upon the
reaction with
.
NO). Approximately 59% of
.
NO in mitochondria is consumed
in the formation of ONOO-, whereas 85% of O
2
.-
dismutates into H
2
O
2
and
15% to ONOO
-
(Poderoso, Lisdero et al. 1999).
13
2) Maintenance of an electrochemical gradient, essential for neurotransmission,
requires the constant activity of the 3Na
+
/2K
+
active transporter, therefore it
can reasonably assumed that a loss in mitochondrial ability to maintain ATP
above a critical threshold level would deleteriously affect the overall function
(neurotransmission)/health of the dopaminergic neurons. Mitochondria, being
the site of oxidative phosphorylation, play a central role in determining the
energy supply inherent in necrotic, apoptotic, and other forms of cell death
(Brown 2000).
3) Two main processes contribute to the steady state level of H
2
O
2
in the
mitochondrial matrix: at the inner membrane (Han, Williams et al. 2001), and
the mitochondrial respiratory chain (Boveris and Cadenas 1975; Boveris,
Cadenas et al. 1976; Cadenas, Boveris et al. 1977; Turrens and Boveris 1980;
Kwong and Sohal 1998) where H
2
O
2
is formed from O
2
.-
disproportionation
(O
2
Æ O
2
.-
Æ H
2
O
2
) and released into either the matrix or intermembrane space.
The above mitochondrial features gain further significance when considering that
complex I impairment, energy deprivation, overproduction of free radicals, and
mitochondrial signaling of apoptotic cascades are all hallmarks of dopaminergic cell
death (Schapira 1998; Olanow and Tatton 1999; Tatton and Olanow 1999).

SIGNIFICANCE
PD is a chronic, late onset disease that leads to devastation of the substantia
nigra and an irreversible loss of quality of life for patients. As the disease etiology
remains complex, the therapeutic options available at present are effective at
14
controlling the symptoms but neither treat the disease or delay the progression of
this disease. The work presented in this dissertation has several novel features that
contribute to a better understanding of the role of the redox status, energy status, and
protein post-translational modification of proteins in the pathogenesis of PD.
a) Importance of the current experimental model
In the brain, the production of NO and dopamine does not occur exclusively in
contained areas, but rather dopamine and
.
NO exist and are released into the same
microenvironments. Therefore, it is of paramount importance to understand the
relationship between NO and dopamine in terms of the potential reactions that can
occur between the two, the products of these reactions and the consequences of these
products to cellular as well as mitochondrial function. The work presented in this
dissertation demonstrates for the first time, the reaction between NO and dopamine
produces ONOO
-
, a powerful oxidant, and the synergistic effects of NO and dopamine
on mitochondrial respiration. This is of significance not only for cellular oxidative/
nitrosative stress but also changes the extracellular redox environment.
b) Protein Modifications: Linking the redox and energy signaling
Damage to proteins under conditions of oxidative and nitrosative stress is highly
specific, and plays an important role in many diseases. Characterization of post-
translational modifications of mitochondrial proteins that occur in PD may provide a
mechanism for NO and dopamine elicited damage to mitochondrial function and
apoptosis in PD. Cell and mitochondrial dysfunction are accomplished through
specific protein modifications in response to the cell’s redox status. Furthermore,
identification of these chemically modified proteins can potentially provide targets for
15
therapeutic intervention therapies. The work presented in this dissertation
demonstrates for the first time that glutathionylation of mitochondrial proteins not
only alter protein activity but also acts as stores of GSH and can be released in
response to changes in mitochondrial redox status as well as protect mitochondria
from oxidants. Additionally, we identified aconitase and ATP synthase as a target of
protein post-translational modification. We demonstrated that aconitase could be
nitrated as well as glutathionylated. ATP synthase, was also identified to be
glutathionylated. These proteins’ post-translational modifications in turn modulated
protein function.

SYNOPSIS
This dissertation is presented in three sections as summarized below:
a) Redox and energy status of brain mitochondria – regulation by glutathionylation:
GSH plays a significant role in the detoxification of ONOO- in the cytosolic and
mitochondrial portion of the cell. GSH depletion is a well-defined characteristic in PD,
but the actual ramifications on mitochondrial welfare are poorly understood, in part,
due to the lack of understanding mitochondrial GSH regulation. This section explores
the pathways involved in mitochondrial GSH regulation and the consequences related
to it.
b) Synergistic mitochondrial damage by dopamine and nitric oxide is mediated by
ONOO
-
production.
.
NO and dopamine and their individual causative roles in
dopaminergic cell death have been studied extensively. This section takes into account
both factors, a more physiologically relevant model, and shows for the first time a
16
synergistic effect on mitochondrial damage and the potential protective effects of
GSH.
c) Critical post-translational protein modifications – altered signaling. ONOO- has
been shown to inactivate the aconitase protein (Castro, Rodriguez et al. 1994), but the
actual nature of ONOO- mediated modification is unknown. This section delves into
the direct and indirect mechanism of ONOO- toxicity. On the one hand ONOO-
induced aconitase inactivation and the other GSH depletion induced glutathionylation
of ATP synthase. This gains further significance, as these proteins are essential in
maintaining the proper energy status of the neuron (Fig 1).
17

Fig. 1. Cellular redox status and function
Inflammation: NADPH oxidase
Mitochondria: UQ
.–
autoxidation
Dopamine autoxidation
O
2
O
2
.–
H
2
O
2
RNH
2
.
NO
Inflammation: iNOS
superoxide
anion
hydrogen
peroxide
molecular
oxygen
guanidino
of L-arginine
nitric
oxide
ONOO
–
peroxynitrite
ROS / RNS
Cell Redox Status
(RSH / RSSR)
Critical Protein Modifications
(Altered Signaling)
Mitochondrial- and Cell
Dysfunction
Apoptotic Cell Death
(Neurodegeneration)
18
Chapter 2: Regulation of redox status in brain mitochondria

INTRODUCTION
Traditionally defined as the cell’s powerhouse, mitochondria are subcellular
organelles whose function was presumed to only synthesize ATP. Since then,
mitochondria have now been also recognized as important sources of free radicals, as
a target itself for free radical regulation, and as a source of signaling molecules that
can modulate redox sensitive cellular processes. The primary free radical generated by
mitochondria is the superoxide anion (O
2
.-
) which is the stoichiometric precursor of
mitochondrial hydrogen peroxide (H
2
O
2
) (Boveris and Cadenas 1975; Dionisi,
Galeotti et al. 1975; Cadenas, Boveris et al. 1977). H
2
O
2
is an effective signaling
molecule, as it is uncharged and highly diffusible. The release of mitochondrial H
2
O
2
(Boveris, Oshino et al. 1972; Boveris and Chance 1973) and its precursor O
2
.-
(Han,
Antunes et al. 2003), into the cytosol, must be carefully regulated as slight alterations
in steady state concentrations of H
2
O
2
have the capacity to regulate the cell’s fate of
proliferation, apoptosis, or necrosis through the redox potential (Antunes and Cadenas
2001).
Detoxification of H
2
O
2
and O
2
.-
is afforded by the mitochondria’s free thiol
pool, glutathione (GSH). In hepatoctyes mitochondrial GSH plays a far more
significant role than cytosolic GSH. Treatment of hepatocytes with 2µM of usnic acid,
a liver toxicant, resulted in necrosis in 10% of the cells. Depletion of cytosolic GSH
by 0.25 mM diethylmaleate and treatment with usnic acid had little to no effect.
Interestingly, a decrease of mitochondrial GSH by 0.50 mM diethylmaleate produced
19
a synergistic effect, as 79% of hepatocytes were necrotic (Han, Matsumaru et al.
2004). Evidence amassed from numerous studies in various cell types exposed to
cytotoxic insults corroborated this finding and further established the importance of
mitochondrial GSH in determining cellular function and viability (Fernandez-Checa,
Garcia-Ruiz et al. 1991; Dhanbhoora and Babson 1992; Uhlig and Wendel 1992;
Shan, Jones et al. 1993; Colell, Garcia-Ruiz et al. 1998; Wullner, Seyfried et al. 1999;
Colell, Coll et al. 2001). Although it is well established that depletion of mitochondrial
GSH pool sensitizes cells to various toxicant induced cell death, the effects of GSH
depletion on mitochondrial function is still ill defined.
Glutathionylation, or protein s-thiolation, of redox sensitive proteins has
emerged as a potential mechanism that sensitizes cells after mitochondrial GSH
depletion. Glutathionylation is a reversible protein post-translational modification,
where redox sensitive cysteinyl thiols are covalently modified by GSH, forming a
protein mixed disulfide (Gilbert 1984). Mitochondrial protein thiols can be
characterized as (i) essential thiols that reside in the active sites of enzymes, (ii)
exposed thiols, and (iii) unexposed thiols. Essential and exposed thiols are of primary
interest as they can interact with the GSH pool and possibly regulate protein
function/activity and influence the mitochondrial redox environment. Intact rat liver
mitochondria contain ~65-70 nmol of thiol/mg of total protein and ~20-25 nmol of
thiols are essential or exposed (Hurd, Costa et al. 2005). Because the concentration of
these two sub-groups is high, it may have a significant impact on mitochondrial
environment and/or function.
20
Protein glutathionylation can occur through several mechanisms; (i) nitric
oxide induced (ii) thiol oxidation and (iii) thiol-disulfide exchange. Nitric oxide
induced glutathionylation is initiated by the s-nitrosation of protein thiols (PrSNO)
and results in glutathionylation through the displacement of the nitroxyl anion by the
glutathionylate anion (GS
-
) (Reaction 1) (Foster and Stamler 2004).
PrSNO + GS
-
Æ PrS-SG + NO
-
Reaction 1
Oxidation of protein thiols can occur through either the two or one electron oxidation
of the thiol to form sufenic acid (RSOH) or thiyl radical (PrS
*
) respectively. Sulfenic
acid will react with the glutathionylate anion and displace the hydroxyl group to
generate the glutathionylated protein (Schafer and Buettner 2001).
PrSOH + GS
-
Æ PrS-SG + OH
-
Reaction 2
Thiyl radical and GS
-
must first form a radical mixed disulfide that is then oxidized by
oxygen to form O
2
.-
and the glutathionylated protein (Thomas, Poland et al. 1995).
PrS* + GS
-
Æ PrS
*-

-
SG + H
+
Reaction 3
PrS
*-

-
SG + O
2
Æ PrS-SG + O
2
*-
Reaction 4
The bulk of protein glutathionylation typically occurs through thiol-disulfide
exchange. Nucleophilic attack by GSSG on the protein’s thiolate anion leads to the
glutathionylation of the protein and the release of GSH.
PrS
-
+ GSSG Æ PrS-SG + GS
-
Reaction 5
The high mitochondrial GSH:GSSG ratio drastically reduces the occurrence of
reaction 5, however an oxidized environment can favor and potentially maintain the
glutathionylated protein indefinitely (Gilbert 1984; Beer, Taylor et al. 2004).
21
Regulation of mitochondrial protein function/activity by glutathionylation is
attractive as this modification is reversible, specific, and sensitive to the redox status
of the mitochondria. Although glutathionylation of a number of mitochondrial proteins
such as complex I (Taylor, Hurrell et al. 2003; Beer, Taylor et al. 2004) and aconitase
(Han, Canali et al. 2005) have been observed, the physiological significance of this
protein post-translational modification remains unclear. Potential roles for this protein
post-translational modification, under physiological conditions, are the regulation and
protection of mitochondria. Regulation – (a) Glutathionylation of key cysteines may
regulate protein activity. Protection – (a) The modification can safeguard essential
thiols by preventing further oxidation to sulfinic and sulfonic acid, which are more
stable-irreversible type modifications (Hurd, Costa et al. 2005). (b) Because the
concentration of essential and exposed thiols is high (Hurd, Costa et al. 2005), direct
reactions with ROS may be an important antioxidant defense (Thomas, Poland et al.
1995). (c) Glutathionylation may also be a mechanism whereby low GSH/GSSG is
sensed and a compensatory mechanism is activated. Glutathionylation of proteins
(Reaction 5) can decrease mitochondrial GSSG and increase GSH, therefore shifting
the GSH:GSSG ratio from low to high. In essence glutathionylation may play a key
protective role in maintaining a reduced mitochondrial environment, regardless of
transport from the cytoplasmic GSH pools. Therefore mitochondria can independently
adapt to increases in oxidative stress.
As mitochondria cannot synthesize GSH nor export GSSG, recycling
mechanisms (GR) as well as GSH/GSSG equilibrium with redox sensitive protein
cysteinyl thiols might represent very important mechanisms that preserve
22
mitochondrial GSH pools, GSH/GSSG and mitochondrial function. Therefore, the
aim of this study was to: (a) to assess mitochondrial redox status in terms of
thiol/disulfide exchange, (b) to establish the significance of glutathionylation of
mitochondrial proteins, and (c) the significance of glutathionylation with respect to
mitochondrial adaptation towards oxidative stress (H
2
O
2
).

Materials and Methods
Chemicals – DTT, percoll, metaphosphoric acid, CHAPS, rotenone, potassium
cyanide (KCN), antimycin were from Sigma Chemical Co. (St. Louis, MO, USA).
Mitochondrial Isolation – Mitochondria from rat brain were isolated by
previously described procedures for differential centrifugation (Sciamanna and Lee
1993) and discontinuous percoll gradient (Anderson and Sims 2000; Schroeter, Boyd
et al. 2003). Sub-Mitochondrial Particles (SMPs) were prepared from freshly isolated
mitochondria after proper treatments and incubations. Mitochondria were sonicated,
by Branson Sonifier 150 Internal Sonicator, 6 times with 10-20% power output each
time for 10 seconds with a 1-minute interval. Intact mitochondria were spun down at
8250 g for 10 minutes. Supernatant containing the SMPs were transferred and diluted
with the mitochondrial isolation buffer and spun down at 80,000 g for 40 minutes
using a Type 60 Ti rotor. Mitochondria were kept on ice throughout the procedure.
Broken mitochondria were prepared by 3 series of freezing and thawing in a 2%
CHAPS solution. Intact mitochondria were spun down at 8250 g for 10 minutes.
Measurement of NAD(P)H Oxidation/Reduction State – Reduced NAD(P)H
was measured flourimetrically with a PerkinElmer LS 55 luminescence spectrometer
23
using an excitation wavelength of 346 nm and an emission wavelength of 460 nm.
NAD(P)H standard curves were done prior to each experiment and used to determine
the concentration of NAD(P)H reduction in the mitochondria.
Western blotting and 2D gel electrophoresis – Mitochondrial samples were
solubilized in a (non)reducing SDS sample buffer, separated by Laemmli SDS/PAGE,
and transferred onto PVDF membranes. Using appropriate antibodies, the
immunoreactive bands will be visualized with an enhanced chemiluminescence
reagent. 2D gel electrophoresis will be performed by using 70% pH 5-7/30% pH 3.5-
10 ampholines in the isoelectric focusing and 10.5% acrylamide gel in the second
dimension. Gels will be stained with Sypro Ruby protein gel or blotted onto PVDF
membrane and probed against appropriate antibodies. Gel images will be acquired
using VersaDoc1000 imaging system (Bio-Rad).
LC/MS/MS – (a) In-gel tryptic digest – Protein bands from SDS-PAGE were
excised from the gels using biopsy punches (Acuderm, Lauderdale, FL, USA). In-gel
tryptic digest was carried out using trypsin that was reductively methylated to reduce
autolysis (Promega, Madison, WI, USA). The digestion reaction was carried out
overnight at 37°C. Digestion products were extracted from the gel with a 5% formic
acid/50% acetonitrile solution (2X) and one acetonitrile extraction followed by
evaporation using an APD SpeedVac (ThermoSavant, Milford, MA, USA). (b)
Analysis of tryptic peptide sequence tags by tandem mass spectrometry – The dried
tryptic digest samples were cleaned with ZipTip and resuspended in 10 µL 60%
formic acid. Chromatographic separation of the tryptic peptides was achieved using a
ThermoFinnigan Surveyor MS-Pump in conjunction with a BioBasic-18 100 mm X
24
0.18 mm reverse phase capillary column (ThermoFinnigan, San Jose, CA). Mass
analysis was done using a ThermoFinnigan LCQ Deca XP Plus ion trap mass
spectrometer equipped with a nanospray ion source (ThermoFinnigan) employing a
4.5 cm long metal needle (Hamilton, 950-00954) using data-dependent acquisition
mode. Electrical contact and voltage application to the probe tip took place via the
nanoprobe assembly. Spray voltage of the mass spectrometer was set to 2.9 kV and
heated capillary temperature at 190°C. The column was equilibrated for 5 min at 1.5
µL/min with 95% solution A, 5% solution B (A, 0.1% formic acid in water; B,
0.1%formic acid in acetonitrile) prior to sample injection. A linear gradient was
initiated 5 min after sample injection ramping to 35% A, 65% B after 50 min and 20%
A, 80% B after 60 min. Mass spectra were acquired in the m/z 400-1800 range. (c)
Protein Identification – Protein identification was carried out with the MS/MS search
software Mascot 1.9 (Matrix Science) with confirmatory or complementary analyses
with TurboSequest as implemented in the Bioworks Browser 3.2, build 41
(ThermoFinnigan). NCBI Sus scrofa protein sequences were used as the primary
search database and searches were complemented with the NCBI non-redundant
protein database.
Measurement of GSH/GSSG. GSH and GSSG will be detected using HPLC
with electrochemical detection as described previously (Harvey, Ilson et al. 1989).

RESULTS
DC and DPG are currently the most commonly used experimental methods to
isolate organelles, specifically mitochondria. Both methods are density-based
25
separation procedures relying on centrifugal force for the purification of
mitochondria. The main difference between DC and DPG is the use of a percoll
gradient. Though percoll increases sample purity, prolonged exposure can compromise
mitochondrial integrity and therefore must be removed by a series of buffer washes.
Elimination of percoll requires added centrifugations and therefore tends to be a more
rigorous method that may compromise mitochondrial integrity and mitochondrial GSH
content. Further understanding of how these two isolation methods affect
mitochondrial GSH content will allow careful interpretation of experimental data and
might provide an additional method that can be utilized for the study of diseases that
exhibit decreases in mitochondrial GSH content such as alcohol induced liver toxicity.
All experiments were carried out in coupled mitochondria unless stated otherwise.
Influence of Isolation Methods on Mitochondrial GSH and GSSG.
Brain and liver mitochondria were isolated using the two different isolation
methods, DC and DPG, to assess the affects of each method of mitochondrial GSH
concentrations. GSH and GSSG concentrations were analyzed using HPLC
electrochemical detection. After isolation, mitochondrial GSH and GSSG
concentrations were stabilized by the addition 5% metaphosphoric acid and also to
ensure protein precipitation. Liver mitochondria isolated via DC and DPG had
different GSH concentrations. DC-isolated liver mitochondrial GSH concentration was
13.91 + 0.64 nmol/mg, and was significantly higher than DPG-isolated liver
mitochondrial GSH concentration that was 5.23 + 0.95 nmol/mg. GSSG
concentrations were the opposite. DC-isolated liver mitochondrial GSSG
concentrations were 0.37 + 0.09 nmol/mg, significantly lower than DPG-isolated liver
26
mitochondrial GSSG that was 0.91 + 0.21 nmol/mg. Thus, liver mitochondrial GSH
concentrations decreased by 62% when using DPG as compared to DC, while GSSG
concentrations increased by 146%. As stated previously, DPG, a harsher method that
increases mitochondria purity, may compromise organelle integrity and as the data
suggest, decrease the initial mitochondrial GSH and increase GSSG concentrations.
GSH concentrations tend to differ between tissues. Therefore, in order to confirm
these findings and to exclude the possibility of tissue specificity, brain mitochondria
were also isolated and the GSH and GSSG concentrations were determined. The high
lipid content and the smaller quantity of mitochondria that can be obtained from the
brain makes isolation by DC a difficult method for obtaining pure mitochondria.
Therefore, DPG is the commonly used isolation method for brain mitochondria as it
removes lipids more efficiently. DPG isolation had a significant effect on brain
mitochondrial GSH concentrations. DPG-isolated brain mitochondrial GSH
concentrations was a mere 0.93 + 0.08 nmol/mg falling beneath its GSSG content of
1.06 + 0.03 nmol/mg. Compared to DC-liver mitochondria and DPG-liver
mitochondria there was 93% and 82% decrease of GSH and a 286% and 116%
increase in GSSG respectively (Table I). Overall DPG, the more laborious isolation
method, resulted in a loss of mitochondrial GSH and an increase in GSSG in both liver
and brain. However, between the two tissues the brain displayed a higher sensitivity to
the isolation methods with respect to GSH depletion.

27

Table I
GSH Concentrations Subsequent to Different Mitochondrial
Isolation Methods
(nmols/mg)

Method GSH GSSG
Differential Centrifugation
Liver 13.91 + 0.64 0.37 + 0.09

Discontinuous Percoll Gradient
Liver 5.23 + 0.95 0.91 + 0.21
Brain 0.93 + 0.08 1.06 + 0.03

Discontinuous Percoll Gradient + DTT
Liver 8.41 + 0.21 0.02 + 0.07
Brain 5.54 + 0.17 0.89 + 0.08
Assay Conditions: Mitochondria were isolated at 4
o
C using the different
methods. The presence of DTT in the buffers is indicated above.
Immediately after isolation mitochondrial protein concentration was
determined, using the Bradford method, 150µg was pelleted, and
resuspended in 5% metaphosphoric acid to precipitate proteins.
Supernatant was then analyzed for its GSH/GSSG content via HPLC.
28
Influence of Buffer Composition on Mitochondrial GSH and GSSG.
Dithiothreitol (DTT) is an excellent reducing agent that is commonly found in
mitochondrial isolation buffers. To determine if the presence of an exogenous
reducing agent can help retain mitochondrial GSH, isolation of mitochondria was
carried out in the presence or absence of DTT. The cellular redox potential has been
demonstrated to change from a more reduced to oxidized state as a cell moves through
different cell cycles. As the cellular redox potential becomes increasingly oxidized
cells progress from apoptosis to necrosis. Thus, this method may be used to represent
reduced (+DTT) or oxidized (-DTT) extra-mitochondrial environment. As
demonstrated previously, DPG isolation of mitochondrial resulted in more pronounced
changes in mitochondrial GSH, and therefore used for the comparison of isolation
buffers with and without DTT. DTT was absent in the isolation buffers used for DC
and DPG in the experiments described above. DPG-isolated liver mitochondrial GSH
and GSSG concentrations were 8.41 + 0.21 and 0.02 + 0.07 respectively, while DPG-
isolated brain mitochondrial GSH and GSSG concentrations were 5.54 + 0.17 and
0.89 + 0.08 respectively. In comparison to the previous data obtained, the buffer that
contained DTT increased GSH concentrations in liver and brain mitochondria by
161% and 597% respectively (Table I). By providing a reduced (+DTT) extra-
mitochondrial environment during isolation, mitochondria tend to retain GSH more
readily as opposed to an oxidized (-DTT) extra-mitochondrial environment. Once
again between the two tissues the brain displayed a hypersensitivity to the different
buffers, which may be comparable to a reduced/oxidized extra-mitochondrial
environment. It should be noted that the mitochondrial membrane integrity was not
29
compromised as the mitochondria isolated using either method were coupled. After
each isolation, measuring oxygen uptake and determining the respiratory control ratio
assessed structural and bioenergetic integrity of mitochondria.
Substrate-Dependent Modulation of Mitochondrial GSH Already published data has
suggested that mitochondrial substrates can prevent peroxynitrite (OONO-)-induced
permeability transition (Scarlett, Packer et al. 1996) and in the formation of free thiols
(Sabadie-Pialoux and Gautheron 1971). Substrate supplementation induces many
changes within the mitochondria such as the generation of NAD(P)H, mitochondrial
respiration, and ATP production. Before investigating GSH and GSSG concentration
changes upon substrate supplementation, NAD(P)H formation was measured
flourimetrically at absorbance 340nm to confirm that substrate supplementation lead
to the generation of reducing equivalents.
Complex I substrates glutamate/malate, or state 4 respiration, generated 6.43
nmol of NAD(P)H/mg of protein, suggesting a reduced environment upon substrate
supplementation (Fig 2). The addition of ADP, state 3 respiration, decreased the
NAD(P)H levels drastically, but a basal amount of ~ 1-2 nmol of NAD(P)H/mg was
left (data not shown).

30

02 46 8
0
5
10
15
20
B
A
6.43 nmol / mg
G/M Mitochondria
8.01 nmol / mg
Flourescence (a.u.)
Time (min)
Fig. 2. NAD(P)H formation upon complex I substrate (glutamate/malate-G/M)
supplementation.
Reduced NAD(P)H was measured flourimetrically using an excitation wavelength
of 346 nm and an emission wavelength of 460 nm. Intact mitochondria were incubated at
room temperature in a respiration buffer for a total of 8 minutes and given glutamate/malate
at the 4 min mark. NAD(P)H standard curves were generated before every experiment to
determine concentration. Complex I inhibitor, rotenone (1µM) was used to determine
maximal NAD(P)H production.
31

0
5
10
15
20
25

GSH (nmol/mg)
0 5 10 15 20 25 30
0.0
0.5
1.0
1.5
GSSG (nmol/mg)
Time (min)
A
B
Fig. 3. GSH and GSSG concentration of (non)energized liver mitochondria
Liver mitochondria were isolated via differential centrifugation (DC), incubated for the
appropriate time at 37
o
C, pelleted, and resuspended in 5% metaphosphoric acid to precipitate the
proteins. Supernatant was analyzed for GSH and GSSG by HPLC. ( ) Control, and ( S )
Glutamate/malate and ADP. All values are nmol/mg + standard error mean.
32
After confirming that substrate supplementation resulted in the generation of
reducing equivalents, liver and brain mitochondrial GSH and GSSG concentrations in
the presence of complex I substrates were monitored over time. Mitochondria were
incubated at 37
o
C in the presence and absence of glutamate/malate and ADP. At the
appropriate time point, mitochondria were removed, pelleted, resuspended in 5%
metaphosphoric acid, and analyzed. DC-isolated liver mitochondrial GSH
concentration, when supplemented was maximized at ~ 14-15 nmol/mg, while in
controls, the GSH concentration was 13-14 nmol/mg. GSSG concentrations decreased
upon supplementation from ~0.4 nmol/mg to ~0, while the GSSG concentrations in
control mitochondria remained unchanged (Fig 3). According to figure 2, NAD(P)H,
the electron donor for glutathione reductase, increased upon substrate
supplementation, therefore the slight increase of GSH upon substrate supplementation
might be attributed to a simple reduction of GSSG to GSH by glutathione reductase.
DPG-isolated liver mitochondria exhibited similar GSH and GSSG changes as
substrate supplementation resulted in slightly higher GSH and lower GSSG
concentrations than control mitochondria. One key difference between the GSH
concentrations between DC- and DPG-isolated liver mitochondria was the initial GSH
concentrations. GSH concentrations were much lower, which was expected as
mitochondria isolated with DPG previously showed a larger decrease in GSH
concentrations as compared to mitochondria isolated with DC. DPG-isolated liver
mitochondria had initial GSH concentrations of 5 nmol/mg which increased to ~13
nmol/mg after substrate supplementation and ~10 nmol/mg without substrate

33

0
5
10
15
20

GSH (nmol/mg)
0 5 10 15 20 25 30
0.0
0.5
1.0
1.5
GSSG (nmol/mg)
Time (min)
A
B
Fig. 4. GSH and GSSG concentration of (non)energized liver mitochondria
Liver mitochondria were isolated via discontinuous percoll gradient (DPG).
Preparation and analysis were carried out as described in figure 1. ( ) Control, and ( S )
Glutamate/malate and ADP.
34
supplementation. Additionally, initial GSSG concentrations decreased from ~0.9 to
0 with substrate supplementation and ~0.5 nmol/mg without substrate supplementation
(Fig 4). Regardless of being isolated by DC or DPG GSH concentrations increased
upon supplementation, but DPG-isolated liver mitochondrial GSH concentration
increase cannot be due to the simple reduction of GSSG back to GSH by glutathione
reductase. Supplementation increased the GSH concentration from ~5 to 13 nmol/mg
of protein, a 2.6-fold increase cannot be due to reduction of GSSG as there was only 1
nmol/mg. This is further demonstrated more effectively in DPG-isolated brain
mitochondria.
DPG-isolated brain mitochondria displayed a robust increase in mitochondrial
GSH upon substrate supplementation. After 5 minutes in state 3 respiration GSH
concentrations increased from ~1 nmol/mg to ~9 nmol/mg, while GSSG concentration
decreased from ~1 nmol/mg to 0. Control mitochondrial GSH concentration slightly
increased from ~1 nmol/mg to ~2 nmol/mg, while GSSG remained relatively the same
(~1 nmol/mg to ~ 0.9 nmol/mg). GSH concentrations plateaued after 5 minutes; a 427
fold increase that was maintained for at least 30 minutes. GSH concentrations in
supplemented brain mitochondria increased from ~ 1 nmol/mg to ~ 9 nmol/mg while
GSSG concentrations decreased from ~ 1 nmol/mg to 0 nmol/mg, once again
suggesting the increase cannot be completely accounted to reduction of GSSG to GSH
by glutathione reductase (Fig 5).
Cytosolic (extra-mitochondrial) environment can influence the redox status of
mitochondria. Isolation methods and the presence/absence of DTT strongly influenced
the GSH levels within the organelle. Under stressful conditions (DPG isolation, a
35

0 5 10 15 20 25 30
0.0
0.5
1.0
1.5

GSSG (nmol/mg)

A
0
2
4
6
8
10

GSH (nmol/mg)

B
Fig. 5. Substrate dependent GSH formation in brain mitochondria
Brain mitochondria were isolated via discontinuous percoll gradient (DPG).
Preparation and analysis were carried out as described in figure 1. ( ) Control, and ( S )
Glutamate/malate and ADP.
36
harsher method, and an oxidized extra-mitochondrial environment, -DTT)
mitochondria had decreased GSH levels. Since this antioxidant is key in detoxification
of ROS and RNS, mitochondria with low GSH concentrations are susceptible to
oxidative and nitrosative stress. This concept can easily be extrapolated to an in vivo
type setting in which an oxidized cytosol will deplete the mitochondrial GSH pool
ultimately compromising ATP production. When depleted of GSH substrate
supplementation of mitochondria was able to restore the GSH levels. This is an
interesting phenomenon as it suggests mitochondria have the ability to sequester GSH
and release it when needed, therefore equipping the organelle with an emergency
storage of GSH. The question then arises how and where is this storage?
Glutathionylation of mitochondrial proteins as potential mechanisms for conservation
of GSH
As shown previously, the decrease in GSSG concentrations cannot completely
explain the increase in the concentrations of GSH, which suggests that substrate
supplementation maybe releasing GSH by de-glutathionylating proteins. To
investigate the possibility that glutathionylation of mitochondrial proteins act as a
store for mitochondrial GSH, mitochondria were lysed by a series of freezing and
thawing in the presence of DTT in order to maximize disulfide bond reduction and the
release of any GSH conjugated to proteins. Mitochondria prepared in this manner were
compared to substrate supplemented mitochondria lysed without DTT. Both DC-
isolated mitochondria exposed to DTT and supplemented DC-isolated liver
mitochondrial GSH concentrations were relatively equal over time (~ 16 nmol/mg).
Slight deviations could once again be attributed to GSSG reduction (Fig 6). DPG-
37

0 5 10 15 20 25 30
0.0
0.5
1.0
1.5

Time (min)
GSSG (nmol/mg)

0
5
10
15
20
25

GSH (nmol/mg)
A
B
Fig. 6. Similar GSH and GSSG concentrations in energized liver mitochondria (DC) versus
non-energized mitochondria exposed to a reducing agent
Comparison of liver mitochondrial GSH and GSSG concentrations between
mitochondria supplemented with glutamate/malate and ADP or a reducing agent. ( S )
Glutamate/malate and ADP, and ( ) DTT. Upon completion of the allotted incubation time at
37
o
C, mitochondria were lysed in the presence/absence of DTT, and diluted with metaphosphoric
acid for a final concentration of 5%. Proteins were pelleted via centrifugation and supernatants
38

0
5
10
15
20

GSH (nmol/mg)
0 5 10 15 20 25 30
0.0
0.5
1.0
1.5
GSSG (nmol/mg)
Time (min)
A
B
Fig. 7. Similar GSH and GSSG concentrations are achieved once the liver mitochondria
(DPG) are energized versus non-energized mitochondria exposed to a reducing agent
Comparison of liver (DPG) mitochondrial GSH and GSSG concentrations between
mitochondria supplemented with glutamate/malate and ADP or exposed to a reducing agent.
Preparation and analysis were carried out as described in Fig 4. ( S ) Glutamate/malate and
ADP, and ( ) DTT. All values are nmol/mg + standard deviation.
39
isolated liver mitochondria exposed to DTT and supplemented DPG-isolated liver
mitochondria had initial GSH concentrations of ~12 nmol/mg and ~5 nmol/mg
respectively. DPG-isolated liver mitochondria exposed to DTT had a consistent GSH
concentration over time of ~12 nmol/mg, while supplementation was needed to equal
this concentration in DPG-isolated liver mitochondria (initial GSH ~5 nmol/mg Æ
supplementation Æ ~12 nmol/mg) (Fig 7). This is an agreement with data shown
previously. DPG-isolated brain mitochondria exposed to DTT and supplemented
DPG-isolated brain mitochondrial GSH began at ~8 nmol/mg and ~1 nmol/mg and
increased to ~9 nmol/mg and ~8 nmol/mg (Fig 8). The increase due to substrate
supplementation is an agreement with DC- and DPG-isolated liver mitochondria,
except that supplementation did not reach the maximum GSH concentration that DTT
exposed mitochondria attained. This could suggest some type of specificity, in the
sense that supplementation maybe activating enzymes that de-glutathionylate specific
mitochondrial proteins as opposed to reducing non-discriminately like DTT. Taken
together, substrate supplementation may increase mitochondrial GSH similarly to
DTT treatment through the release of GSH from glutathionylated proteins.
To confirm the possibility of a SH moiety conservation by proteins via
glutathionylation and that substrate supplementation de-glutathionylates protein to
release GSH, western analysis of isolated mitochondria was carried out to identify the
presence of GSH-protein adducts. These experiments were carried out in brain
mitochondria isolated by DPG in the absence of DTT as previous data has already
demonstrated that the observed changes in GSH concentrations were more dramatic.
To compare glutathionylated and de-glutathionylated mitochondrial proteins,
40

A
B
0
2
4
6
8
10
12

GSH (nmol/mg)

0 5 10 15 20 25 30
0.0
0.5
1.0
1.5

Tim e (m in)
GSSG (nmol/mg)

Fig. 8. Similar GSH and GSSG concentrations are achieved once the brain mitochondria
(DPG) are energized versus non-energized mitochondria exposed to a reducing agent
Comparison of brain (DPG) mitochondrial GSH and GSSG concentrations between
mitochondria supplemented with glutamate/malate and ADP or exposed to a reducing agent.
Preparation and analysis were carried out as described in figure 4. ( S ) Glutamate/malate and
ADP, and ( ) DTT.
41
incubations with and without substrates occurred for 10 minutes at 37
o
C and
eventually lysed with a 2% CHAPS non-reducing sample buffer. Mitochondria,
without substrate supplementation, were lysed with a reducing sample buffer (+DTT)
to act as a negative control. Previous data (Fig. 4) indicates that 10 minutes incubation
is sufficient for most of the potentially glutathionylated proteins to undergo de-
glutathionylation. Western blotting with anti-glutathionylation antibody revealed 3
bands at ~ 40kd, ~60kd, and 70kd in control mitochondria that were DPG isolated
mitochondria without DTT (oxidized extra-mitochondrial environment).
Consistent with the GSH concentration increase, the addition of substrates
decreased the 40 kd band intensity, while the 60 kd and 70 kd bands disappeared,
implying that supplementation promotes the process of de-glutathionylation (Fig 9A).
When comparing the GSH profile in Fig 8 and the western blot, supplementation does
not completely de-glutathionylate all proteins and the amount of free GSH released is
slightly less than that of DTT. On the other hand, DTT non-specifically de-
glutathionylates removing all GSH protein adducts and maximizes GSH release. This
also suggests specificity in the glutathionylation and de-glutathionylation process that
maybe regulated by mitochondrial substrates directly or indirectly.
Immunoprecipitation followed by LC/MS/MS was used to identify the
mitochondrial proteins that were glutathionylated. The 70 kd band was identified as
succinyl CoA: 3-oxoacid CoA-transferase (SCOT) (gi 34854196, 68.260kd), which
plays a role in ketone body metabolism. The 60 kd band was identified as ATP
synthase, H+ transporting, mitochondrial F1 complex, α subunit, isoform 1
42

Control
+ G/M
& ADP
+DTT
192.7
126.4
81.1
40.3
IP
59.8 kDa:
ATP Synthase F
1
α subunit isoform
1 (GI:40538724)
68.3 kDa:
Succinyl CoA:
3-oxoacid CoA-
transferase
(SCOT)
(GI:34854196)
A B
Fig. 9. Substrate-induced protein sulfhydryl recovery (deglutathionylation) of glutathione-
protein adducts and identification of immunoprecipitated proteins by LC/MS/MS
A. DPG isolated brain mitochondria were incubated at 37
o
C for 10 min. with the
appropriate substrates. Mitochondria were lysed in a 2% CHAPS/non-reducing and
reducing buffer, mitochondria underwent a series (3X) of freezing and thawing to
ensure maximal protein release, separated via SDS-PAGE, transferred to a
nitrocellulose membrane, and probed using a Virogen GSH antibody (1:500).
B. Immunopercipitation (IP) of mitochondrial proteins occurred in a 2% CHAPS/non-
reducing buffer containing the GSH antibody (1:200). IP proteins were then separated
via SDS-PAGE. Protein bands stained by sypro ruby were excised and identified by
LC/MS/MS.
43
(gi 40538742, 59.831 kd) (Fig 9B). The purified F1 sector is made up of five
different subunits with a stoichiometry of α
3
β
3
γδε (Amzel, McKinney et al. 1982;
Bianchet, Ysern et al. 1991). One of the cysteine residues in the α subunit is close
both spatially and in sequence to the glycine-rich loop, which is thought to be involved
in binding one or more of the ATP phosphates (Bianchet, Ysern et al. 1991).
Identification and abundance of these proteins within the mitochondria not
only represent a potential storage of GSH, it may also play a role in mitochondrial
respiration regulation, which will be examined in section 3. De-glutathionylation
represents a mechanism in which mitochondria will be able to independently adapt to
an oxidative or nitrosative stress regardless of GSH transport, or lack there of, from
the cytosol. This in turn will allow the organelle to continue to synthesize ATP and the
cell to respond properly to the particular stress.
Increased GSH content in brain mitochondria via de-glutathionylation can protect
mitochondria from pharmacological doses of H
2
O
2
.
Buffers with DTT and supplementation of DPG-isolated brain mitochondria
increased GSH content. Whether this increase has any significance in terms of
mitochondrial function is still a question. Therefore, to examine the potential
protective effects of GSH brain mitochondria were isolated by DPG using a buffer that
contained DTT and one that did not. The mitochondria isolated in the absence of DTT
were incubated with glutamate/malate and ADP prior to treatment. DTT was shown to
preserve mitochondrial GSH while supplementation increased GSH concentration
through the de-glutathionylation of mitochondrial proteins. Mitochondria were then

44

0
1
2
3
4
5
6
H
2
O
2
(mM)
1.0 0.5 0.1 0.02 0
GSH (nmol/mg)
0.0
0.5
1.0
1.5
2.0
1.0 0.5 0.1 0.02 0
GSSG (nmol/mg)
H
2
O
2
(mM)
Fig. 10. H
2
O
2
induced GSH depletion and GSSG formation in rat brain mitochondria
Brain (DPG) mitochondrial isolated using a buffer with DTT, to conserve
mitochondrial GSH. Once isolated mitochondrial were washed and resuspended in a
respiration buffer and incubated with H
2
O
2
for 10 min at 37
o
C. Mitochondria were washed
of any excess H
2
O
2
and measured for GSH and GSSG concentrations as described in Fig.
1.
45

0
2
4
6
8
10
12
5.0 1.0 0.5 0.1 0.02 0
GSH (nmol/mg)
H
2
O
2
(mM)
Fig. 11. H
2
O
2
-induced GSH depletion and GSSG formation in rat brain mitochondria
supplemented with complex I substrates and ADP
Brain (PG) mitochondrial isolated using a buffer without DTT. Once isolated
mitochondrial were resuspended in a respiration buffer and incubated with glutamate malate,
ADP, and with the appropriate concentration of H
2
O
2
for 10 minutes at 37
o
C. Mitochondria
were washed of any excess H
2
O
2
and measured for GSH and GSSG concentrations as
described in Fig. 1. GSSG was measured but present only in the mitochondrial sample that was
treated with 5.0 mM H
2
O
2
(0.36 nmol/mg + 0.03 GSSG).
46
exposed to both physiological and non-physiological concentrations of H
2
O
2
. H
2
O
2

dose-dependently decreased GSH with a concomitant increase of GSSG, when
mitochondrial GSH was conserved by DTT (Fig 10). However, when supplemented
with mitochondrial substrates, physiological concentrations of H
2
O
2
had little to no
effect on GSH content or GSSG formation. Only high or pharmacological H
2
O
2

concentrations were able to decrease GSH, and any detectable GSSG occurred only at
5 mM H
2
O
2
exposure (0.36 nmol/mg + 0.03) (Fig 11). Supplemented mitochondria
were isolated in the absence of DTT therefore the ability to buffer the H
2
O
2
was
strictly due to glutamate/malate and ADP supplementation induced de-
glutathionylation.

Discussion
Mitochondria lack the machinery to synthesize GSH (Griffith and Meister
1985) and thus are dependent on cytosolic GSH influx via a mitochondrial inner
membrane carrier-mediated transport protein (Martensson, Lai et al. 1990; Garcia-
Ruiz, Morales et al. 1995; Cummings, Angeles et al. 2000). Isolation methods (DC or
DPG) and buffer constituents (+/- DTT) play a key role in determining the initial
redox status of isolated mitochondria and can have significant effects on experimental
models/treatments and interpretation. This problem is not trivial, for less rigorous
isolation methods (DC) and buffers with reducing agent (+DTT) tend to conserve
mitochondrial GSH. Conservation maybe based on the activity of the GSH transport
proteins. 1-oxoglutarate, one of many carrier-mediated transport proteins, displayed
sensitivity to phenylsuccinate, membrane fluidity, ATP, and GSH (Martensson, Lai et
47
al. 1990; Coll, Colell et al. 2003; Lluis, Colell et al. 2003). The presence of GSH
provides a substrate to the carrier, but may also offer a reduced environment for
maximal protein activity. DTT in the isolation buffer, a reduced extra-mitochondrial
environment, may influence the activity of mitochondrial inner membrane carrier-
mediated GSH transport proteins to conserve the mitochondrial GSH. The influence of
DTT’s presence or absence in the isolation buffer to mitochondrial GSH maybe
compared to the cytosol’s redox status. When the cytosolic environment is oxidized
mitochondria release free GSH while a reduced environment conserves it. DPG-
isolated brain mitochondria displayed a higher sensitivity to the absence DTT in the
isolation buffer, with respect to GSH depletion, as it was very dramatic, compared to
liver. This sensitivity correlates to work done with respect to glutathione depletion and
neurotoxicity. When GSH is exhausted in neurons it causes: (1) increased formation of
reactive oxygen species and lipid peroxidation (Garcia-Ruiz, Colell et al. 1995), (2)
loss of mitochondrial membrane potential (Wullner, Seyfried et al. 1999), (3)
degeneration of brain mitochondria in the newborn rat (Jain, Martensson et al. 1991),
and (4) decreased activity of mitochondrial enzymatic complexes (Bolanos, Heales et
al. 1996) to name a few.
BSO, a glutamate-cysteine ligase inhibitor, has helped establish the importance
of glutathione in both the cytosol and mitochondria. Experiments with BSO
administered as a single dose, typically exhibit a biphasic response to cytosolic GSH
depletion, while mitochondrial GSH decrease is slow and gradual (Romero and Sies
1984; Griffith and Meister 1985). After hypoxia-ischemia, mitochondrial GSH
transiently decreased, recovered, and then decreased after 14 h (Wallin, Puka-Sundvall
48
et al. 2000). This phenomenon of rapid cytosolic GSH and slow mitochondrial GSH
depletion, suggested a separate pool of GSH within the mitochondria. Explanations
such as synthesis of GSH within mitochondria and faster GSH incorporation into the
mitochondria were proposed but found to be inconsequential as no glutamylcysteine
synthetase activity was found and incorporation of GSH into the cytosol and
mitochondria were equal (Griffith and Meister 1985). Glutathionylation of
mitochondrial proteins provides another alternative as it may function as both a post-
translational modification and storage of GSH within the mitochondria. As cytosolic
GSH decreases, mitochondria sequester the GSH moiety within glutathionylated
proteins and can release GSH when needed. The idea of storage and release is further
supported by the rapid increase of mitochondrial GSH when supplemented with
substrates and the concurrent de-glutathionylation of mitochondrial proteins.
SCOT and ATP synthase, H+ transporting, mitochondrial F1 complex, α
subunit, isoform 1 protein were identified as being endogenously glutathionylated.
SCOT and ATP synthase α subunit has 12 and 2 potentially modifiable cysteine
residues. The ATP synthase is made up of 3 α subunits, therefore one ATP synthase
protein has a total of 6 potentially modifiable cysteine residues. Reactive protein
thiols, compared to the thiol content in total GSH, is much greater and the interactions
between these two thiol pools plays a key role in the antioxidant defense (Thomas,
Poland et al. 1995; Jung and Thomas 1996; Schafer and Buettner 2001). Though there
may only be 18 potential sites for glutathionylation per the two proteins, when taking
into account protein amount within the mitochondria, a potentially large storage of
GSH is possible (Hurd, Costa et al. 2005). This concept was confirmed by measuring
49
protein thiol content using DTNB (Ellman 1959). Mitochondrial proteins were
precipitated and incubated with or without DTT, to de-glutathionylate all proteins, for
15 min and washed thoroughly. Protein thiol content without DTT was 18.8 nmol/mg
and increased to 28.3 nmol/mg in the presence of DTT, supporting its storage capacity
for GSH.
To determine potential mechanisms of de-glutathionylation, GSH was
monitored in mitochondria with different complex inhibitors. By using different
complex inhibitors it would be possible to determine if electron flow through the
electron transport chain is crucial as well as identify the specific complex that maybe
controlling GSH release. Rotenone (complex I inhibitor), antimycin (complex III
inhibitor), and KCN (complex IV inhibitor) had no effect on GSH content within
mitochondria, thus suggesting that GSH release was independent of the electron
transport chain (Fig 12). However, NAD(P)H formation coincided with the increase in
GSH formation (Fig 2). As discussed previously a reduced environment can activate
certain proteins, an increase in NAD(P)H manipulates the redox status (reduced)
within the mitochondria and/or provides reducing equivalents for proteins that may de-
glutathionylate proteins, one such enzyme that has been proposed to de-glutathionylate
proteins within the mitochondria is glutaredoxin 2 (Grx2). Grx2 was identified and
characterized as a mitochondrial thiol/disulfide oxidoreductase (Gladyshev, Liu et al.
2001; Lundberg, Johansson et al. 2001; Ehrhart, Gluck et al. 2002). The physiological
role of Grx2 is under investigation, but recent work has shown that when exposed to
H
2
O
2
Grx2 was able to protect cells against H
2
O
2
-mediated disruption of
mitochondrial transmembrane potential (Fernando, Lechner et al. 2006), reduce
50

0
2
4
6
8
10
12
nmol/mg
0
2
4
6
8
10
nmol/mg
GSH
GSSG
Succinate - + - + + + + +
ADP - + + - - - - -
Antimycin A - - - - + - + -
Rotenone - - - - - + + -
KCN ------- +
Glutamate/Malate - + - + + + + +
ADP - + + - - - - -
Antimycin A - - - - + - + -
Rotenone - - - - - + + -
KCN - ------ +
B
A
Fig. 12. GSH formation depends on substrate availability not electron flow through the ETC
DPG-isolated brain mitochondria were incubated for 10 min. at 37
o
C with the
appropriate factors. All inhibitors were added prior to supplementation with
glutamate/malate, succinate, or ADP. Upon completion of 10 min. mitochondria were
pelleted, resuspended in 5% metaphosphoric acid and analyzed for GSH and GSSG.
51
GSSG to GSH (Johansson, Lillig et al. 2004), and catalyze the glutathionylation/de-
glutathionylation reaction (Beer, Taylor et al. 2004; Johansson, Lillig et al. 2004). We
show that protein similar to 3-oxoacid CoA transferase and ATP synthase, H+
transporting, mitochondrial F1 complex, α subunit, isoform 1 as potential targets for
Grx2. Ultimately Grx2’s physiological role would be balancing the buffering of
cytosolic GSH depletion with conservation of the GSH moiety within mitochondria
and regulation of ATP production via glutathionylation/de-glutathionylation.
Glutathionylation and de-glutathionylation of mitochondrial proteins provide
potential explanations and mechanisms to: (i) the protective effects of mitochondrial
substrate supplementation, (ii) the biphasic response of BSO to cytosolic GSH as
opposed to the slow and gradual depletion of mitochondrial GSH, and (iii) a
physiological role for Grx2. It maybe surmised that substrate supplementation
protection and the initial depletion of cytosolic GSH by BSO and hypoxia-ischemia
are buffered by the mitochondria via de-glutathionylation of proteins, resulting in the
transient increase of GSH. Though this process may have a fundamental protective or
regulatory role, mitochondrial dysfunction maybe a consequence of persistent
glutathionylation or de-glutathionylation of mitochondrial proteins.
The reduction capacity, measured by the Nernst equation using the GSH/GSSG
redox couple, in mitochondria as compared to the cytoplasm and endoplasmic
reticulum is –270 mV, – 220 mV, and –150 respectively (Hansen, Go et al. 2006). The
mitochondrial compartment is the most reduced, but also the most susceptible to
oxidative stress. This susceptibility and the significance of this organelle justifies it
reduction capacity, which is strictly based on the concentration of free thiols. Taking
52
into account the ability to store GSH via glutathionylation of proteins only further
increases this reductive capacity as GSH can be released when needed.
These studies support the following notions: (a) extra-mitochondrial
environment can influence the mitochondrial redox status, (b) substrate
supplementation increases GSH levels that is mediated through de-glutathionylation,
and (c) that this increase in GSH, via de-glutathionylation is significant as it was able
to buffer pharmacological doses of H
2
O
2
. Glutathionylation represents a post-
translational modification that can not only regulate proteins, but also act as a support
for GSH when it becomes depleted. This becomes exceedingly significant under
oxidative/nitrosative stress conditions. As cytoplasmic GSH decreases to nullify the
increase in ROS and/or RNS the mitochondria become vulnerable as GSH transport
into the organelle ceases. Supplementation of GSH via de-glutathionylation allows the
continued protection /function of mitochondria. Though other portions of the cell
maybe compromised energy production is not, therefore allowing the possibility of the
cell to respond accordingly to the particular stress. Under acute exposures of ROS and
RNS, these active and critical cysteines within the mitochondrial proteins would only
be exposed for a short period as the cell would quickly re-establish the status quo and
replenish the mitochondrial GSH. Chronic exposures present a major obstacle as this
environment would never allow mitochondrial GSH replenishment, therefore these
active and critical cysteines would be susceptible to oxidation and more stable post-
translational modifications. PD, a neurodegenerative disease, is an appropriate
example of the latter circumstance as the neurons within the substantia nigra pars
compacta are constantly exposed to oxidants generated from both dopamine and
53
inflammation. The next section explores the consequences of GSH depletion, with
respect to mitochondrial respiration, in the presence of dopamine and
.
NO, a product
of inflammation.

54
Chapter 3: Synergistic action of dopamine and nitric oxide in neuronal injury

Introduction
Oxidative stress is classically defined as an imbalance between oxidants and
antioxidants. Overwhelming evidences suggest an integral role of oxidative stress in
numerous disease pathologies such as aging (Finkel and Holbrook 2000) and
neurodegenerative diseases (Bains and Shaw 1997) such as Parkinson’s disease (PD).
PD is defined as the selective death of dopaminergic neurons (synthesizes and uses
dopamine as a neurotransmitter) within the substantia nigra pars compacta. PD is
characterized by the progressive and selective loss of dopaminergic neurons in the
substantia nigra pars compacta. In cases of PD, various indices of oxidative stress
such as protein nitration in Lewy bodies (Good, Hsu et al. 1998), 40-50% decrease in
glutathione (GSH) levels (Sian, Dexter et al. 1994), and upregulation of manganese
superoxide dismutase (MnSOD) (Saggu, Cooksey et al. 1989) have been observed in
the specific regions of the brain, consistently linking oxidative stress to dopaminergic
neuronal loss.
Although the mechanisms that lead to selective loss of dopaminergic neurons
in PD remains unclear, previous work carried out in our laboratory have demonstrated
that mitochondrial damage by nitric oxide occurs in PC12 cells. Dopamine, on one
hand is utilized as a neurotransmitter by dopaminergic neurons but on the other hand
sensitizes dopaminergic neurons to additional damage. This is supported in part by
the generation of superoxide (O
2
.-
), electrophilic quinones/semiquinones that can cause
cellular damage and neuromelanin through dopamine autoxidation.
55
Additionally, it has also been demonstrated that neuroinflammation plays a
role in the progression of Parkinson’s disease. Nitric Oxide (
.
NO) is produced by the
resident immune cells in the brain, microglia, as well as glial cells such as astrocytes.
Examination of post partum brain sections from individuals afflicted with PD
demonstrates a co-localization of activated microglia with neuronal death in the
substantia nigra (Mcgeer, Itagaki et al. 1988). In vitro studies have further
demonstrated that
.
NO released from astrocytes compromise mitochondrial oxidative
phosphorylation in the surrounding neurons (Barker, Bolanos et al. 1996). Mutant
mice lacking inducible nitric oxide synthase were more resistant to MPTP-induced
dopaminergic neurodegeneration (Liberatore, Jackson-Lewis et al. 1999; Dehmer,
Lindenau et al. 2000). Taken together, the aforementioned data suggests that the role
of
.
NO in PD is also extremely important in disease progression.
Therefore, it may be surmised based on the aforementioned evidences that
dopamine and
.
NO can have a synergistic effect on mitochondrial function and the
toxic effects of dopamine and
.
NO can be mediated in part through the formation of
peroxynitrite (ONOO
-
), a potent oxidant formed through the reaction of O
2
.-
and
.
NO
at diffusion limited rates. This hypothesis is supported by the observation that
glutathione depletion occurs prior to mitochondrial complex I inhibition through a
peroxynitrite mediated inhibition in dopaminergic cells (Chinta and Andersen 2006).
Although the role of NO on mitochondrial respiration in dopaminergic cells has been
investigated, the possible synergistic inhibition of dopamine and
.
NO on PC12 and
brain mitochondria remains unclear. Additionally, the potential mechanisms that lead
to mitochondria impairment and cellular function remain ill defined. In this study, we
56
demonstrate: (i) ONOO
-
production by dopamine in the presence of
.
NO, (ii) the
importance of GSH in maintaining dopamine levels by preventing its oxidation, and
(iii) that dopamine and
.
NO can synergistically inhibit of mitochondrial respiration
through formation of ONOO
-
.

Materials and methods
Chemicals – Dopamine and digitonin were from Fluka (Buchs, Switzerland).
Diethylamine/
.
NO (DEA-NO) complex, Diethylenetriamine/
.
NO (DETA-NO)
adduct, Cu,Zn-superoxide dismutase (from bovine red blood cells), malate, glutamate,
ADP, were from Sigma Chemical Co. (St. Louis, MO, USA).
Oxygen consumption – Oxygen consumption was measured amperometrically
with a Clark-type electrode (Hansatech, UK) assembled to a thermostatic water jacket.
Solutions in the electrode chamber were maintained under continuous stirring with a
magnetic agitator. For calibration, the saturating O
2
concentration in air equilibrated
distilled water at 37
o
C was taken as 253 µM (Wilhelm et al. 1977) and the zero was
established in the presence of sodium dithionite.
Electron paramagnetic resonance (EPR) measurements – EPR spectra were
obtained with a Bruker ECS 106 spectrometer (operating at X-band) equipped with a
cylindrical room temperature cavity operating in TM
110
mode. Aliquots (150 µl) of the
reaction mixtures were transferred to bottom-sealed Pasteur pipettes and measured at
room temperature under the instrument settings described in the figure legends.
57
Spectrophotometric measurements – All the spectrophotometric
measurements were done with an Agilent 8453 (diode-array) UV-visible
spectrophotometer equipped with a Peltier temperature control accessory. The cuvette
holder was thermostatically maintained at 37
o
C, and a magnetic stirrer was used for a
continuous mixing of the sample. RCR, without any addition, was used as the
reference measurement (blank).
Mitochondrial Isolation – Mitochondria from rat brain were isolated by
discontinuous percoll gradient as previously described (Anderson and Sims 2000;
Schroeter, Boyd et al. 2003) .
Cell culture – PC12 cells from ATCC were cultured in complete medium
(RPMI-1640 medium supplemented with 10% horse serum, 5% fetal calf serum, L-
glutamine, and antibiotics). Cells were incubated at 37˚C in humidified air with 5%
CO
2
and kept in logarithmic phase by routine passage.
Incubation conditions - PC12 cells (1.25 mg of protein) were incubated with
the
.
NO donor (di-ethylamine/nitric oxide complex) for 30 minutes at 37ºC (
.
NO
release rate at t
0
was 3 µM × s
–1
). The half-life of the donor was ~2.1 min; hence,
every 2.1 min,
.
NO release was decreased by 50%. Incubations were carried out in the
presence or absence of exogenous dopamine (1 mM). Cells were spun down,
collected, and used for mitochondrial respiration. Isolated brain mitochondria were
incubated for 3 minutes at 37ºC with the
.
NO donor (Diethylenetriamine/nitric oxide
(DETA-NO):
.
NO release rate at t
0
was 3.8 nM × s
–1
) and/or dopamine (1 mM) within
the chamber of the oxygen electrode to allow immediate determination of respiration.
The half life of this donor is ~ 1200 min.
58
Mitochondrial damage – Complex I-driven respiration was measured in
respiration buffer in the presence of malate/glutamate (state 4) and malate/glutamate
plus ADP (state 3) at room temperature in digitonin-permeabilized cells (1.25 mg of
protein). Mitochondrial damage was expressed as inhibition of the respiratory control
(RC) calculated as (RC
control
– RC
sample
) / RC
control
– 1).

Results
ONOO
-
formation via dopamine and
.
NO
The production of O
2
.-
as a product of dopamine oxidation has already been
demonstrated (Herlinger, Jameson et al. 1995) and was corroborated by trapping the
radical with DMPO and detected using electron paramagnetic resonance. DMSO was
added to differentiate between the formation of hydrogen peroxide and O
2
.-
. The
spectrum revealed the formation of methyl radical (methyl group cleaved by hydroxyl
radical from hydrogen peroxide), hydroxyl radical, and superoxide (Fig 13A). Though
many species were detected, the addition of SOD significantly decreased the spectrum,
suggesting that all the species detected originated from O
2
.-
(Fig 13B).

59

Fig. 13. Determination of O
2
.-
, from dopamine oxidation, using electron paramagnetic
resonance
Spin trapping of O
2
.-
was performed using DMPO (experimental conditions – A:
RCR solution contained 10 mM dopamine, 1% DMSO, and 500 mM DMPO and B: A +
SOD). EPR spectra were recorded with a Bruker ECS 106 spectrometer (operating at X-
band) equipped with a TM
110
(Bruker’s TM 8810) room temperature cavity. Aliquots of
the reaction mixtures were promptly transferred to bottom-sealed Pasteur pipettes and
measured at room temperature. Instrumental conditions: microwave frequency 9.77 GHz;
microwave power 20 mW; modulation frequency 100 kHz; modulation amplitude 1.0 G;
receiver gain 8x10
5
; scan rate 1.0 G s
-1
; time constant 655 ms; number of scans
accumulated = 5.
3440 3460 3480 3500 3520
B)
A)
magnetic field (Oe)
60

0 5 10 15 20 25 30
0
200
400
600
800
1000
1200
1400
1600
1800
2000
Dopamine
C
B
A
.
NO Electrode Current (pA)
Time (min)
Fig. 14. ONOO
-
formation by dopamine in the presence of
.
NO
ONOO
-
is generated by the reaction of
.
NO with O
2
.-
. The presence of these two
species and the formation of ONOO
-
was measured either directly or indirectly using a
Clark-type
.
NO electrode in the presence and absence of the protein SOD. At the one-
minute mark DETA-NO was added and allowed to accumulate for 15 minutes to reach its
plateau. The different conditions are as follows: A – nitric oxide 3.85 nM/s, B – A + 1
mM Dopamine, and C – A + 1 mM Dopamine + SOD.
61
To confirm the formation of ONOO
-
by dopamine and
.
NO,
.
NO
concentrations in the presence or absence of dopamine were monitored using a Clark-
type
.
NO electrode. DETA-NO, a slow releasing
.
NO donor, reached a 1 µM steady
state concentration in 15 minutes, and was maintained for up to 3 hours (Fig. 14A).
The addition of dopamine immediately decreased the
.
NO current/signal as well as
prohibited any further release by the donor (Fig 14B) suggesting a consumption of
.
NO. The addition of SOD prevented the consumption of
.
NO in the presence of
dopamine (Fig 14C), further confirming the formation of ONOO- by O
2
.-
, generated
from dopamine oxidation, and its reaction with
.
NO. Though ONOO- formation has
been proposed as a key player in PD development and the presence of nitrotyrosine (a
finger print of ONOO
-
induced nitration) was detected in the substantia nigra of PD
patients (Good, Hsu et al. 1998), the actual source of it remains unclear. Dopamine
oxidation and the subsequent O
2
.-
generation, during inflammation, represent a novel
source of intracellular and extracellular ONOO- for dopaminergic neurons (Fig 15).
To date there are no known
.
NO or ONOO
-
scavengers, therefore in order to cope with
ONOO
-
toxicity, the cell must prevent the diffusion rate limited reaction of O
2
.-
and
.
NO. The only portion of this reaction that can be manipulated, from a PD
perspective, is O
2
.-
generated by dopamine oxidation. Comprehension of the dopamine
oxidation process and the effects of antioxidants, such as GSH and SOD, are crucial in
the prevention of O
2
.-
production and consequently the formation of ONOO
-
.
EPR spin stabilization of dopamine-o-semiquinones.
Characterization of dopamine oxidation was performed by monitoring the
formation of dopamine-o-semiquinone, the first product generated in the oxidation
62

Inflammation
H
2
O
2
.
NO
ONOO-
Dopamine
Semiquinone
.
O
-
O
NH
2

Dopamine
NH
2
HO
HO
Dopaminochrome
precursor to neuromelanin
H
2
O
2
O
2
.-
O
2
.-
O
2
O
2
O
2
.-
O
2
Transition metals, Enzymes,
.
NO, O
2
O
O
NH
2
Dopamine
Quinone
N
O
HO
Fig. 15. Simplified scheme of the dopamine oxidation process and its byproducts
63

0 5 10 15 20 25
0
2
4
6
8
10
12
14
28 min
16 min
11 min
2 min
Magnetic Field (G)
EPR Signal Intensity (a. u.)
Time (min)
3465 3470 3475 3480 3485 3490 3495

Fig. 16. Dopamine-o-semiquinones decay during dopamine autoxidation in RCR
At time point zero, dopamine (final concentration of 10 mM) was added to RCR
containing MgCl
2
(0.5 M). EPR spectra were recorded at time intervals of approximately 3
min. The EPR signal intensity was obtained by double integration of the time point spectra
(inset) and plotted against time. Instrument settings were: microwave frequency, 9.77 GHz;
microwave power, 20 mW; field modulation frequency, 100 kHz; field modulation amplitude,
1 G; receiver gain, 4 x 10
5
; time constant, 21 ms; scan rate, 1.4 G s
-1
; number of scans
accumulated, 3.
64

process, dopaminochrome, a downstream product, as well as oxygen consumption in
the presence and absence of GSH. Electron paramagnetic resonance spectroscopy was
used to verify the formation of the dopamine-o-semiquinones intermediates. Recent
work from our group (Rettori, Tang et al. 2002) demonstrated that the one electron
oxidation of dopamine generates two isomeric o-semiquinones, which present a
characteristic composite EPR spectrum. By applying the spin-stabilization technique
with Mg
2+
, the characteristic EPR spectrum (Fig. 16-inset) was obtained during the
autoxidation of dopamine, confirming the formation of the semiquinones. Figure 16
shows the decay profile of the EPR signal intensity, obtained by double integration of
the time point spectra, as a function of time. After ~ 25 min, the signal disappeared as
oxygen within the buffer was consumed, re-oxygenation (data not shown) was able to
recover the EPR signal−dopamine was present in large excess. This suggests that
semiquinone formation during dopamine oxidation is dependent upon oxygen
concentrations.
Formation of dopaminochrome coincides with oxygen consumption
It is well established that further oxidation of dopamine semiquinone leads to
the formation of dopaminochrome, which is rapidly converted to neuromelanin a
neurotoxic compound. To determine if dopaminochrome formation is also oxygen
dependent, we measured simultaneously, the consumption of oxygen using a Clark
type oxygen electrode and monitored dopaminochrome formation by monitoring
absorbance changes. During the reaction, the formation of colloidal melanines
contributes to a general increase of absorbance due to light scattering (Herlinger,
65
Jameson et al. 1995), therefore, in order to follow properly the levels of
dopaminochrome formation, absorbance values at 480 nm (at different time points)
were subtracted from the time-correspondent absorbance values at 700 nm. These
subtractions were possible because a UV-visible spectrophotometer with diode-array
technology, which records a full spectrum (190-1100 nm) at specified time intervals
(min. 0.5 s), was employed for the experiments. The addition of dopamine
immediately caused a consumption of oxygen that coincided with the increase in
dopaminochrome formation, further supporting the oxidation route of dopamine into
dopaminochrome with the consumption of oxygen (Fig 17). As dopamine was present
in great excess, after anaerobiosis has taken place, re-oxygenation re-established
dopamine oxidation with dopaminochrome formation as indicated by further increase
of absorbance at 480 nm (data not shown). Thus, the further conversion of dopamine
semiquinone to dopaminochrome is oxygen dependent.
Dopamine autoxidation results in a number of key events, such as: (i) oxygen
consumption with subsequent O
2
.-
generation, (ii) dopamine semiquinone formation, a
early product of the oxidation process, and (iii) dopaminochrome generation, a late
product of the oxidation process and is the precursor of neuromelanin. Typically the
latter products of dopamine oxidation are more toxic than O
2
.-
, but in the presence of
.
NO, ONOO
-
can be formed. Though, experimentally, SOD was able to prevent
ONOO- production, physiologically the reaction of the two radicals would out-
compete SOD, leaving GSH as the only potential antioxidant defense against ONOO
-
.
66

0 5 10 15 20 25
0
50
100
150
200
250

Time (min)
[O
2
] (µM)
Oxygen
0.00
0.02
0.04
0.06
0.08
0.10
∆A
480-700
Dopaminochrome
Fig. 17. Dopaminochrome formation and oxygen consumption during dopamine autoxidation
At time point 1 min (indicated by arrows), dopamine was added to RCR buffer at
37
o
C−final concentration of dopamine was 10 mM. Dopaminochrome formation was followed
spectrophotometrically at 480 nm−measurements were corrected by subtracting the time
correspondent absorbance values at 700 nm. Oxygen consumption was monitored
amperometrically with a Clark type oxygen electrode.
67
Inhibiting the formation of dopamine semiquinone and dopaminochrome by GSH.
Glutathione concentrations in the brain have been estimated to lie in the low
micromolar range extracellularly, and in the milimolar range within the cell. Aside
from its ability to react with reactive oxygen species, GSH can also react with
electrophiles, thus enabling the extrusion of the GSH conjugates out of cells through
specific transporter systems. Due to its pivotal role in antioxidant defense and cellular
detoxification, we investigated the protective effects of GSH against dopamine
autoxidation products. The addition of GSH delayed oxygen consumption and the
formation of dopamine-o-semiquinone and dopaminochrome.
Using a Clark-type electrode, the consumption of oxygen was monitored
during dopamine oxidation in the presence of GSH. As demonstrated previously, the
autoxidation of dopamine lead to the consumption of oxygen (Fig 17). However,
when GSH was included in the reaction, there was a lag phase in dopamine oxidation
driven oxygen consumption. Dopamine driven oxygen consumption occurred in two
phases in the presence of GSH. Initial consumption of oxygen was slower than that
observed in the absence of GSH. At approximately 18 minuets, the rate of oxygen
depletion increased to a rate similar to that observed in the absence of GSH (Fig 18A).
Additionally, this biphasic response, in the presence of GSH, was also observed in the
formation of dopamine-o-semiquinone and dopaminochrome.
Autoxidation of dopamine in the absence of GSH resulted in almost
instantaneous detection for both the dopamine-o-semiquinone and dopaminochrome.
However, in the presence of GSH, no detectable signal for both dopamine-o-
semiquinone and dopaminochrome was observed until after the 18 minute mark (Fig
68

0
50
100
150
200
A
[O
2
] (µM)
0
1
2
3
4
5
6
50 min
35 min
22 min
14 min
3 min
B
Magnetic Field (G)
EPR Signal Intensity (a. u.)
3470 3480 3490

0 102030 40 50
0.00
0.02
0.04
0.06
0.08
C
∆A
480-700
Time (min)
Fig. 18. Effects of GSH on oxygen consumption, and formation of dopamine-o-
semiquinone, and neuromelanin
Oxygen, dopamine semiquinone, and dopaminochrome were measured as described
in fig. 3 and 4, but in the presence of 0.1 mM of GSH.
69

O
2
O
2
.-
HO
HO
NH
2
Dopamine
NH
2
.
O
-
O
Dopamine
Semiquinone
GSH
GS
.
Dopaminochrome
precursor to neuromelanin
N
O
HO
Fig. 19. Proposed mechanism of dopamine recycling/reduction of dopamine semiquinone by GSH
70
18B & 18C). A slightly slower rate of oxygen consumption and the inhibition of
dopamine-o-semiquinone and dopaminochrome formation within this 18-minute time
frame suggest that GSH may be reducing the semiquinone back to form dopamine.
Once GSH is depleted, it can no longer prevent the consumption of oxygen and the
subsequent formation of dopamine oxidation products (Fig 19).
To confirm if GSH is indeed reducing the dopamine-o-semiquinone back to
dopamine by donating one electron, EPR analysis in conjunction with the spin trap
DMPO was used to detect the formation of glutathionyl radical, the one electron
oxidation product of GSH. Indeed, in the presence of dopamine, GSH underwent a
one electron oxidation to form the glutathionyl radical. (Fig.20)
In PD GSH levels are significantly low, suggesting that in the presence of
.
NO,
ONOO
-
will be formed. To test the consequences of this oxidants formation, an
intracellular and extracellular model was developed using PC12 cells, a dopaminergic
cell line, and isolated brain mitochondria.
Synergistic effects of dopamine and nitric oxide on mitochondrial respiration.
The effects of
.
NO on mitochondrial respiration of PC12 cells displayed a
sigmoid pattern of inhibition rather than a linear one (data not shown). This indicates a
threshold level of
.
NO, below which little damage is exerted. However, once this
threshold is reached inhibition of respiration is almost complete. At low
.
NO
concentrations (2.5 µM x s
-1
), 15% inhibition of mitochondrial respiration occurred
while at higher
.
NO concentrations (4.0 µM x s
-1
), the minimum needed to reach the
threshold, mitochondria respiration was inhibited by 90%. The effects of
.
NO at low
concentrations on PC12 cell mitochondrial respiration were dramatically augmented in
71

3440 3460 3480 3500 3520

Magnetic Field (G)
Fig. 20. Detection of the glutathionyl radical using electron paramagnetic resonance
Spin trapping of the electron donor radical, GS
.,
was performed using DMPO
(experimental conditions: RCR solution contained 10 mM dopamine; 0.1 mM GSH; 500
mM DMPO). Instrument settings: A) modulation amplitude = 1G, time constant = 41 ms,
scan rate = 2 G/s, number of scans accumulated = 14. B) modulation amplitude = 2G, time
constant = 82 ms, scan rate = 0.2 G/s, number of scan accumulated = 15.
72
the presence of dopamine. While dopamine itself had little effect on mitochondrial
respiration, dopamine and
.
NO lead to a 70% inhibition of mitochondrial respiration.
It has been hypothesized that dopamine, though necessary for
neurotransmission, can sensitize mitochondria of dopaminergic neurons to further
damage. In the previous experiment exogenous or extracellular dopamine was added
to the PC12 cells, as it has been proposed that dopamine can diffuse away from the
synapse into the extracellular area (Vizi 2000). In order to test the significance of the
endogenous or intracellular dopamine, the storage of the neurotransmitter was
manipulated. Reserpine, a compound that depletes cellular dopamine storage vesicles
(Schubert and Klier 1977), protected PC12 mitochondria from the damaging effects of
.
NO at high concentrations, as it was able to rescue mitochondrial respiration by ~
40%. Although long term incubations with reserpine is toxic, these results support the
idea that dopamine sensitizes mitochondria and therefore exacerbates the effects of
.
NO (Table II).
O
2
.-
formed during the oxidation of dopamine, in the presence of
.
NO can lead
to the formation of ONOO
-
. The potential role ONOO
-
plays in the synergistic
inhibition of dopamine and
.
NO on mitochondrial respiration was investigated using
isolated brain mitochondria. Inhibition of respiration by dopamine on isolated brain
mitochondria (30% inhibition) was more dramatic as compared to PC12 cell
mitochondria (1% inhibition). The different methods of dopamine exposure in both
models may explain the differences in the values for mitochondrial respiration.
Isolated brain mitochondria are in direct contact with dopamine and its oxidation
products (intracellular dopamine model), while a permeable cell membrane might act
73

TABLE II
Potentiation of nitric oxide-mediated damage by dopamine
in PC12 cells

Inhibition of Respiratory Control
Conditions (RC
control
– RC
sample
) / (RC
control
– 1)
(%)

+ dopamine 0.9 ± 0.5
+ NO (2.5 µM × s
–1
) 15.0 ± 5.7
+ NO (2.5 µM × s
–1
) + dopamine 70.9 ± 8.9
+ NO (4.0 µM × s
–1
) 90.1 ± 7.5
+ NO (4.0 µM × s
–1
) + reserpine 48.4 ± 8.5
Assay conditions: PC12 were exposed for 30 min at 37°C to exogenous dopamine
(1mM) or
.
NO (initial flux rate either 2.5 µM × s
–1
or 4.0 µM × s
–1
) or
.
NO plus
dopamine. Reserpine concentration was 1 µM. Cells were collected and mitochondrial
damage assessed as described in the Materials and Methods section. A flux rate of
.
NO
of 2.5 µM × s
–1
is at or near the threshold level; that of 4.0 µM × s
–1
is above the
threshold level and elicits maximal mitochondrial damage.
TABLE III

ONOO- induced brain mitochondrial dysfunction by nitric
oxide and dopamine autoxidation

Inhibition of Respiratory Control
Conditions (RC
control
– RC
sample
) / (RC
control
– 1)
(%)
+ dopamine 29.6 ± 2.6
+ NO (3.85 nM × s
–1
) 11.2 ± 3.2
+ NO + dopamine 72.3 ± 3.7
+ NO + dopamine + SOD 25.3 ± 1.8
Assay conditions: Isolated brain mitochondria were exposed for 3 minutes at 37°C to
exogenous dopamine (1 mM) or
.
NO (initial flux rate 3.85 nM × s
–1
) or
.
NO plus
dopamine. Superoxide dismutase (SOD at 20µM) was added to the latter treatment to
prevent ONOO- formation through the removal of superoxide.
74
to buffer PC12 mitochondria from direct exposure to dopamine and/or its oxidation
products (extracellular dopamine model). Treatment of isolated brain mitochondria
with dopamine and
.
NO also resulted in a synergistic inhibition of mitochondrial
respiration (72% inhibition). Addition of Cu,Zn-superoxide dismutase (Cu/Zn SOD)
attenuated mitochondrial respiration inhibition to 25% inhibition, suggesting that
ONOO- is the mediator of the synergistic inhibition afforded to dopamine and
.
NO
(Table III).

Discussion
According to the data shown here,
.
NO at low concentrations can cause minor
damage to the mitochondria of cells, thus cells have the ability to withstand some
increases of
.
NO. As the concentrations of
.
NO increase and pass a threshold, cells lose
the ability to protect its mitochondria. The presence of dopamine causes this threshold
to be much lower, therefore cells that synthesize and/or store dopamine are sensitive to
small alterations in
.
NO concentrations. This concept is particularly relevant in the
development of PD as the neurons within the substantia nigra pars compacta
synthesize and store dopamine and are also exposed to
.
NO during an inflammatory
response. It can be surmised that neurons, in general, will be able to buffer the
increase in
.
NO concentrations during inflammation, while only the neurons that
synthesize dopamine will be affected. Because neurotransmission, via dopamine, is
only afforded to specific areas such as the substantia nigra pars compacta, selective
neuronal degeneration can occur, a hallmark in PD.
75
The synergistic damage of dopamine and
.
NO to mitochondria was mediated
in part by ONOO
-
, a potent oxidant that can react fairly quickly. One way to deal with
the deleterious effects of ONOO
-
is to prevent the diffusion rate limited reaction of
.
NO and O
2
.-
to form ONOO
-
. This strategy was employed in this experiment by using
SOD, which was able to rescue mitochondrial respiration. In the case of PD studies
have shown the upregulation of SOD (Saggu, Cooksey et al. 1989), which further
supports this proposed mechanism of ONOO
-
induced death, in explaining the
selective death of the substantia nigra pars compacta in PD. As stated previously
ONOO
-
production has been proposed and nitration of proteins, a ONOO- finger print,
has been detected in the substantia nigra of PD patients, but the actual source remain
unclear. We show a novel and an abundant source of ONOO- from the oxidation of
dopamine in the presence of
.
NO, which causes the synergistic inhibition of
mitochondrial respiration. PD cybrids (engineered cell lines containing PD
mitochondria) have implicated mitochondria as a major player in the selective death of
dopaminergic neurons, as these cells are more susceptible to methyl
phenylpyridinium, a toxin to induce PD development (Hodaie, Neimat et al. 2007).
This increased susceptibility may be due to mitochondria that have been damaged
from the constant bombardment of ROS and RNS generated from dopamine oxidation
and inflammation.
Another potential mechanism in which dopamine oxidation compromises
mitochondria, that is rarely discussed or even considered, is the consumption of
oxygen to generate O
2
.-
. In humans the brain makes up a minuscule portion (2%) of
the total body weight, but is responsible for 20% of basal oxygen consumption. This
76
small portion of the body devours a significant amount of oxygen to sustain ATP
levels that will allow neurons to maintain ion gradients for action potential
propagation (Halliwell and Gutteridge 1989). Thus a depletion of either oxygen or
ATP can have negative effects on neurons, with respect to function and health.
Dopamine oxidation, as shown in Fig. 17, consumes oxygen very rapidly. This process
can create an anaerobic microenvironment within the substantia nigra pars compacta
that can lead to a compromised neuron. Couple this setting with the production of
.
NO
via inflammation and a scenario is created in which mitochondria can be seriously
damaged.
.
NO has the ability to bind and inhibit cytochrome oxidase of complex IV
within the electron transport chain. The extent of inhibition is dependent on the
[oxygen]/[
.
NO] ratio. As the ratio decreases the degree of binding and inhibition
increases (Boveris, Costa et al. 1999), thus preventing mitochondrial respiration and
ATP synthesis.
GSH depletion is another hallmark of PD. This loss of GSH is not responsible
for the selective death, as inhibition of GSH synthesis in the brain did not reduce the
number of dopaminergic neurons (Toffa, Kunikowska et al. 1997). Though GSH may
not directly play a role in the death of dopaminergic neurons, its indirect role is crucial
as it has the ability to prevent dopamine oxidation and the subsequent production of
O
2
.-
, which can lead to ONOO
-
formation. The mechanism in which GSH can prevent
the ONOO
-
formation is through: (i) reduction of dopamine semiquinone back to
dopamine (prevention of dopamine oxidation, Fig 5A-C), (ii) covalent binding to
dopamine quinine (prevention of dopamine oxidation) (Spencer, Jenner et al. 1998),
and (iii) the direct reaction with ONOO
-
(Srisook, Kim et al. 2005). GSH was able to
77
prevent dopamine oxidation via a one-electron reduction of dopamine semiquinone
back to dopamine as long as the antioxidant was present the oxidation products of
dopamine were not generated as depicted in the initial 18 minutes of Fig. 18B&C.
Once depleted the oxidation of dopamine preceded and with it the generation of O
2
.-
,
quinones and neuromelanin. GSH depletion in PD will not only allow dopamine
concentrations, a necessary neurotransmitter, to decrease, but also the generation of its
toxic oxidation byproducts (ONOO
-
in the presence of
.
NO) and a shift in the redox
status of the cell to a more oxidized environment allowing redox sensitive post-
translational modifications, such as nitration (Good, Hsu et al. 1998), to occur. Studies
support the relevance GSH depletion as a prerequisite of dopaminergic cell death.
Postmortem brain tissues (Dexter, Sian et al. 1994) and in vitro studies using a
dopaminergic cells report the a GSH deficit precedes the loss of complex I activity via
a peroxynitrite mediated event (Chinta and Andersen 2006).
Although SOD allowed respiration in the presence of
.
NO and dopamine, and
insinuated ONOO
-
as the key product, the concentrations used may or may not be
physiologically relevant. Upregulation of SOD in PD patients must be at least 10 fold
higher than
.
NO in order to have a protective effect. The reaction rate of
.
NO and O
2
.-

is k
2
= 1.9 x 10
10
M
-1
s
-
and takes precedent over the disproportionation of O
2
.-
by
superoxide dismutase, which occurs at ~10-fold slower rate (2.3 x 10
9
M
-1
s
-1
). SOD
concentration used in this experiment was 20 fold higher than that of
.
NO. Though the
relevance of SOD in preventing ONOO
-
formation in vivo is in question, the role GSH
plays is not. If indeed SOD cannot out-compete
.
NO for O
2
.-
, the significance of GSH
78
depletion increases as it maybe the only antioxidant defense that can prevent the
sensitization of dopaminergic neurons to
.
NO (Fig 21).
ONOO
-
toxicity, which is responsible for the synergistic inhibitory effects of
dopamine and
.
NO, maybe viewed as either direct or indirect. ONOO
-
can directly
react with protein tyrosine residues to form a stable post-translational modification.
One such protein that has critical tyrosines is aconitase. Aside from its tyrosine
content, aconitase displayed hypersensitivity to ONOO
-
as compared to other
mitochondrial proteins. Complex I, complex IV, and fumarase activity, compared to
control, decreased by ~5-15% when exposed to ONOO
-
, while aconitase lost 85% of
its activity. ONOO
-
indirect toxicity is mediated through GSH oxidation to GSSG. The
effects of increasing concentrations of GSSG were assessed by incubations with
aconitase. Section one of the dissertation delved into the results (glutathionylation) of
brain mitochondria exposed to an oxidized (-DTT) extra-mitochondrial environment.
Glutathionylation of the ATP synthase represents a major finding as this protein
complex is responsible for the generation of ATP. Section 3 investigates the
consequences of the direct and indirect toxicity of ONOO
-
, with respect to post-
translational modifications and altered function.

79

O
O
NH
2
Inflammation
Transition metals, Enzymes, NO
.
, O
2
O
2
O
2
.-
O
2
O
2
O
2
.-
O
2
.-
H
2
O
2
Covalent Binding
1. GSH
2. Protein Thiols
Dopaminochrome
precursor to neuromelanin
HO
HO
NH
2
Dopamine

NH
2
.
O
-
O
Dopamine
Semiquinone
GSH
GS
.
ONOO-
GSH
Dopamine
Quinone
N
O
HO
Fig. 21. GSH depletion associated with dopamine oxidation
80

Chapter 4: Critical post-translational protein modifications

Introduction
Aconitases (citrate (isocitrate) hydrolyase, EC 4.2.1.4) are iron-sulfur cluster-
containing proteins present both in mitochondria and cytosol of cells; the enzymes
catalyze the stereospecific conversion of citrate to isocitrate via the intermediate
formation of cis-aconitate (Beinert and Kennedy 1993; Lauble and Stout 1995). The
cubane [4Fe-4S]
2+
cluster in the active site is essential for catalytic activity, but it also
renders aconitase highly vulnerable to reactive oxygen- and nitrogen species (Gardner
and Fridowich 1991; Castro, Rodriguez et al. 1994). Three of the iron atoms in the
cubane structure are bound to cysteine residues of the protein backbone; the fourth
iron (Fe
α
) is ligated to inorganic sulfurs and participates in the binding of substrates to
the active site (Beinert and Kennedy 1993; Lauble and Stout 1995; Bulteau, Ikeda-
Saito et al. 2003). Superoxide anion (O
2
.–
), hydrogen peroxide (H
2
O
2
), and
peroxynitrite (ONOO
–
) have been shown to inactivate mitochondrial aconitase
through modifications of the [4Fe-4S]
2+
cluster (Gardner and Fridovich 1992; Castro,
Rodriguez et al. 1994; Nulton-Persson and Szweda 2001; Bulteau, Ikeda-Saito et al.
2003). ONOO
–
is believed to disrupt the Fe-S cluster by causing a loss of labile Fe
from the cluster resulting in an inactive [3Fe-4S]
1+
form (Castro, Rodriguez et al.
1994). However, direct measurements confirming the nature of ONOO
–
-mediated
modifications of the Fe-S cluster have not been obtained yet.
81
ONOO
–
may also modulate aconitase activity through modifications of amino
acids, such as cysteine and tyrosine. Aconitase contains 12 cysteine residues, with
three contributing ligands to the [Fe-S] cluster and one cysteine residing in the active
site (Kennedy, Spoto et al. 1988; Lauble and Stout 1995). Although the latter is not
essential for aconitase activity, its binding to various agents (e.g., NEM, N-
ethylmaleimide; DTNB, 5,5’-dithiobis(2-nitrobenzoic acid)) results in decrease
aconitase activity, probably by inhibiting citrate entry to the active site (Kennedy,
Spoto et al. 1988; Lauble and Stout 1995). Because thiol residues can be oxidized by
ONOO
–
and H
2
O
2
(Radi, Beckman et al. 1991; Bulteau, Ikeda-Saito et al. 2003), they
remain potential targets of oxidative attack. The reaction of ONOO
–
with thiols
generates primarily a thiyl radical (reaction 1), which may decay to a sulfenic acid
(reaction 3) via a nitrated sulfur intermediate (reaction 2) (Carballal, Radi et al. 2003).
RSH + [HO
.

…
NO
2
.
] → RS
.
+ NO
2
.
+ H
2
O

[1]
RS
.
+ NO
2
.
→ RSNO
2
[2]
RSNO
2
+ OH
–
→ RSOH + NO
2
-
[3]
Tyrosine residues react with ONOO
–
to form 3-nitrotyrosine as the major product
(reactions 4-5 show one possible mechanism) (Radi, Cassina et al. 2002). Aconitase
contains 22 tyrosine residues, and whether nitrotyrosine formation in some of these
residues may affect enzyme activity remains to be determined.
R-TyrH + [HO
.

…
NO
2
.
] → RTyr
.
+ NO
2
.
+ H
2
O [4]
RTyr
.
+ NO
2
.
→ R-Tyr-NO
2
[5]
Appreciation of the mechanisms inherent in aconitase inactivation by ONOO
–
in a
cellular setting requires consideration of enzyme substrate availability and the
82
pathways of ONOO
–
formation in mitochondria. Cysteine and tyrosine residues of
aconitase may be important targets of oxidant attack, for the [Fe-S]-containing active
site of aconitase may be protected by citrate in a cellular setting. It has been shown
that the presence of citrate can protect the thiol residues in the active site from thiol
alkylating agents, such as NEM (Kennedy, Spoto et al. 1988), and shield the Fe-S
cluster from inactivation by O
2
.–
(Hausladen and Fridovich 1996). Whether citrate may
sterically block the Fe-S cluster from ONOO
–
in a similar fashion, thereby rendering
amino acid residues vulnerable to oxidation, has not been explored.
Mitochondria are important sources of ONOO
–
, for the mitochondrial respiratory
chain (Cadenas and Boveris 1980; Han, Antunes et al. 2002) and the inner membrane-
associated mitochondrial nitric oxide synthase (Elfering, Sarkela et al. 2002; Riobo,
Melani et al. 2002; Ghafourifar and Cadenas 2005) furnish both reactants, O
2
.–
and
.
NO, required for ONOO
–
formation (O
2
.–
+
.
NO → ONOO
–
), a reaction that proceeds
at diffusion-controlled rates. In addition,
.
NO regulates both mitochondrial respiration
and O
2
.–
production by its reversible binding to cytochrome oxidase (Torres, Darley-
Usmar et al. 1995; Brown 1997; Antunes, Boveris et al. 2004) and inhibition of
electron transfer at the bc
1
segment (Poderoso, Carreras et al. 1996; Cadenas,
Poderoso et al. 2001), respectively. ONOO
–
, thus formed, has been reported to inhibit
complex I activity (Schopfer, Riobo et al. 2000; Riobó, Clementi et al. 2001),
stimulate inner membrane proton permeability (Brookes, Levonen et al. 2002), trigger
cytochrome c release to regulate apoptosis (Packer and Murphy 1994), and it appears
to contribute to the pathology of many diseases including Alzheimer’s disease,
myocardial ischemia, and amyotrophic lateral sclerosis (Beckman and Koppenol
83
1996). Similarly, O
2
.–
diffusing from mitochondria to cytosol via voltage-dependent
anion channels (Han, Antunes et al. 2003) can combine with
.
NO to yield ONOO
–
in
cytosol, thus widening the spectrum of protein targets (among them cytosolic
aconitase) of this powerful oxidant.
In this study, the sites and mechanisms of aconitase inactivation by ONOO
–
were
examined using a porcine aconitase preparation: the role of citrate in ONOO
–
-
mediated damage of aconitase iron-sulfur cluster and amino acids was investigated
and specific targets for ONOO
–
-mediated oxidation and nitration were identified. The
significance of these mechanisms within the context of the mitochondrial
thiol/disulfide status was assessed in terms of the sensitivity of aconitase to
glutathionylation. ATP synthase was identified in section 1 as another glutathionylated
protein. Using substrate supplementation as a means of de-glutathionylation, the
consequence of this modification on ATP synthase was determined.

Materials and methods
Chemicals – 4-(2-Hydroxyethyl)piperazine-1-ethanesulfonic acid (HEPES),
ethylenediaminetetraacetic acid (EDTA), bovine serum albumin, dithiothreitol (DTT),
ascorbic acid, NADP
+
, 5,5-dimethyl-1-pyrolline-N-oxide (DMPO), 3-morpholino-
sydnonimine (SIN-1), and DTNB (5,5’-dithiobis(2-nitrobenzoic acid)) were from
Sigma Chemical Co. (St Louis, MO, USA). Supelco P-10 sephadex G-25 columns
were from Amersham Pharmacia Biotech (Piscataway, NJ). Peroxynitrite (ONOO
–
)
was obtained from Upstate (Waltham, MA, USA). All other reagents were of
analytical grade. A water solution of DMPO containing 0.1 mM DETAPAC
84
(diethyenetriamine-pentaacetic acid) was purified several times with activated
charcoal; DMPO concentration was calculated spectrophotometrically (ε
232
= 7700 M
–
1
cm
–1
); the stock solution was kept under He at –20°C. ONOO
–
concentration was
verified on the day of the experiment by UV absorption spectrometry (ε
302
= 1670 M
–
1
cm
–1
).
Porcine aconitase preparations –obtained from Sigma Chemical Co. (St Louis,
MO, USA)– had been similarly used in a previous study involving ONOO
–
(Castro,
Rodriguez et al. 1994). Analysis of the aconitase preparation using SDS gel and mass
spectrometry revealed the fraction to contain other proteins, particularly porcine serum
albumin used to stabilize aconitase. Therefore mass spectrometry was used to confirm
the identity and purity of the aconitase band used in western blot analysis.
Aconitase assay – Porcine heart aconitase (15 mg) was activated in a reaction
containing 0.5 mM of ferrous ammonium sulfate and 5 mM of dithiothreitol buffered
with 100 mM Tris-HCl, pH 7.5, and incubated under He at 0°C for 30 min. Activated
aconitase was rapidly separated on Sephadex G-25 pre-equilibrated with He-saturated
100 mM Tris/HCl, pH 7.5 (Gardner, Costantino et al. 1997). Aliquots of activated
aconitase were stored under He at –80°C. Aconitase activity was assayed on a reaction
mixture consisting of 50 mM Tris-HCl, pH 7.4, 30 mM sodium citrate, 0.6 mM
MnCl
2
, 0.2 mM NADP
+
, and 1 U/ml of isocitrate dehydrogenase; the reaction was
followed at 340 nm (ε
340
= 6.22 mM
–1
cm
–1
) for 10 min at room temperature. An
aconitase activity of 1 mU corresponded to 1 nmol NADPH formed per min (Drapier
and Hibbs 1996).
85
Aconitase (2 mU/ml) in 50 mM Tris-HCl, pH 7.4, was treated with varying amounts
of ONOO
–
. During the addition of ONOO
–
, the aconitase-containing solution was
rapidly vortexed in order to ensure complete mixing before significant decomposition
of ONOO
–
occurred. Aconitase activity was measured immediately after mixing.
For experiments involving SIN-1, aconitase samples were incubated at room
temperature with varying amounts of SIN-1 for 10 minutes. To remove SIN-1 from
aconitase, treated samples were spun in micro bio-spin desalting column (Biorad,
Hercules, CA) for 4 minutes at 4°C. The desalted samples were immediately analyzed
for aconitase activity. Aconitase samples treated with GSSG were pre-incubated for 10
min with GSSG before aconitase activity was assessed. Control experiments were
performed to verify that GSSG did not affect isocitrate dehydrogenase activity. In
samples treated with DTT, aconitase was incubated with citrate and GSSG followed
by a 10-min treatment with 10 mM DTT. The presence of citrate was required when
working with DTT, for DTT directly affected the active site of aconitase.
Immunoblotting for protein thiols, glutathionylation, and nitrotyrosine –Aconitase
preparations (4 mU/ml) were treated with either ONOO
–
or GSSG in 50 mM Tris-
HCl, pH 7.4 and run on an 8% SDS-Page non-reducing gel. Western blot analysis for
nitrotyrosine in aconitase was determined with an anti-nitrotyrosine antibody (Upstate,
Charlottesville, VA). Glutathionylation of aconitase was assessed by western blotting
using an anti-glutathione monoclonal antibody (Virogen, Watertown, MA) (Wang,
Boja et al. 2001) after incubation with various concentrations of GSSG for 10 min.
Changes in aconitase protein thiols were determined by labeling the enzyme with
maleimide PEO
2
-biotin (20 mM; Pierce, Rockford, IL) for 1 hr. Western blot analysis
86
of biotinylated proteins was subsequently performed using a streptavidin antibody
(Pierce, Rockford. IL). Band densities were estimated with a Versadoc Image System
(BioRad, Hercules, CA). The band corresponding to aconitase in all western blots was
confirmed by LC/MS/MS analysis.
LC/MS/MS – (a) In-gel tryptic digest – Protein spots from SDS-PAGE were
excised from the gels using biopsy punches (Acuderm) or biopsy. In-gel tryptic digest
was carried out using trypsin that was reductively methylated to reduce autolysis
(Promega, Madison, WI). Prior to digestion, samples were neither reduced with DTT
nor alkylated with iodoacetamide in order to keep potential cysteine modifications
stable. The digestion reaction was carried out overnight at 37°C. Digestion products
were extracted from the gel with a 5% formic acid/50% acetonitrile solution (2X) and
one acetonitrile extraction followed by evaporation using an APD SpeedVac
(ThermoSavant). (b) Analysis of tryptic peptide sequence tags by tandem mass
spectrometry – The dried tryptic digest samples were cleaned with ZipTip and
resuspended in 10 µL 60% formic acid. Chromatographic separation of the tryptic
peptides was achieved using a ThermoFinnigan Surveyor MS-Pump in conjunction
with a BioBasic-18 100 mm X 0.18 mm reverse phase capillary column
(ThermoFinnigan, San Jose, CA). Mass analysis was done using a ThermoFinnigan
LCQ Deca XP Plus ion trap mass spectrometer equipped with a nanospray ion source
(ThermoFinnigan) employing a 4.5 cm long metal needle (Hamilton, 950-00954)
using data-dependent acquisition mode. Electrical contact and voltage application to
the probe tip took place via the nanoprobe assembly. Spray voltage of the mass
spectrometer was set to 2.9 kV and heated capillary temperature at 190°C. The
87
column equilibrated for 5 min at 1.5 µL/min with 95% solution A, 5% solution B
(A, 0.1% formic acid in water; B, 0.1%formic acid in acetonitrile) prior to sample
injection. A linear gradient was initiated 5 min after sample injection ramping to 35%
A, 65% B after 50 min and 20% A, 80% B after 60 min. Mass spectra were acquired
in the m/z 400-1800 range. (c) Protein Identification – Protein identification was
carried out with the MS/MS search software Mascot 1.9 (Matrix Science) with
confirmatory or complementary analyses with Turbo Sequest as implemented in the
Bioworks Browser 3.2, build 41 (ThermoFinnigan). NCBI Sus scrofa protein
sequences were used as the primary search database and searches were complemented
with the NCBI non-redundant protein database. Each MS/MS spectrum was analyzed
for methionine and cysteine oxidation (+16), tryptophan and tyrosine (+45), cysteine
and tyrosine for dioxidation and trioxidation (+32 and +48) as specific differential
modifications. Aconitase amino acid numbering is based upon the Sus scrofa reference
sequence NP 999119; for comparison with the amino acid numbering in the reported
the crystal structure of aconitase (Robbins and Stout 1989), it should be considered
that the first amino acid in the crystal structure is number 28 in the reference sequence
at NCBI. The aconitase structure shown in Fig. 7 was generated with the WebLab
Viewer Pro 3.7 (Molecular Simulations Inc., San Diego) from the protein data bank
(Structure explorer - 6ACN) based on the aconitase crystal structure (Robbins and
Stout 1989).
Electron paramagnetic resonance (EPR) – (a) DMPO/cysteinyl radical-protein
spin adduct measurements: EPR spectra were recorded with a Brucker ECS 106
spectrometer, equipped with a TM
110
room temperature cavity. Spectra acquisition
88
began 2 min after the rapid mixing of ONOO
–
(0.75 mM) into a mixture containing
50 mU/ml aconitase and 65 mM DMPO in 100 mM Tris-HCl, pH 7.5. Instrument
settings: microwave frequency, 9.77 GHz; microwave power, 20 mW; field
modulation frequency, 100 kHz; field modulation amplitude, 2 G; receiver gain, 8 x
10
5
; time constant, 328 ms; scan rate, 0.9 G s
–1
; number of scans accumulated, 5.
Experiments were performed at room temperature. Computer simulations of spectra
were performed using the WinSIM program (EPR calculations for MS-Windows NT
95, version: 0.96, from P.E.S.T. – Public EPR Software Tools) (Duling 1994). (b)
Glutathionyl radical (GS
.
) measurements: Spectra were recorded 1 min after the rapid
mixing of ONOO
–
(0.7 mM) in 100 mM Tris HCl, pH 7.5, containing 18 mM GSH,
52 mM DMPO, and different concentrations of aconitase. The EPR signal intensities
were calculated by double integration of the 3470 G line. Instrument settings:
microwave frequency, 9.77 GHz; microwave power, 20 mW; field modulation
frequency, 100 kHz; field modulation amplitude, 1 G; receiver gain, 8 x 10
5
; time
constant, 82 ms; scan rate, 1.9 G s
-1
. Experiments were performed at room
temperature. (c) Aconitase [3Fe-4S]
1+
measurements: The active form of aconitase
([4Fe-4S]
2+
) is EPR silent, whereas the inactive aconitase form ([3Fe-4S]
1+
) shows an
EPR signature at g ~ 2.02 at low temperatures (Kennedy, Antholine et al. 1997). EPR
spectra were recorded at 10K. Aconitase suspensions (50 mU/ml) were rapidly mixed
with various concentrations of ONOO
–
in 100 mM Tris-HCl, pH 7.5 and frozen in
liquid nitrogen before measurements. Instrument settings: microwave frequency, 9.177
GHz; microwave power, 100 mW; field modulation frequency, 100 kHz; modulation
amplitude, 0.5 mT; number of scans accumulated, 5.
89
ATPase Activity – ATPase activity of mitochondrial complex V was assayed by
coupling its activity to pyruvate kinase and lactate dehydrogenase as previously
described(Darley-Usmar 1987). The hydrolysis of ATP by ATPase was charged to the
conversion of phosphoenolpyruvate to pyruvate by pyruvate kinase. Pyruvate was then
converted to lactate by lactate dehydrogenase with the consumption of NADH.
Therefore, the consumption of NADH, monitored by spectrophotometer at 340 nm,
was indicative of ATPase activity. Basic reaction buffer consisted of 50 mM HEPES
pH 8.0-KOH, 5 mM MgSO
4
, and 250 mM sucrose. Right before the experiment, 5 µl
of 0.5 M sodium phosphoenolpyruvate, 5 µl of 0.4 µg/µl antimycin A, 5 µl of
PK/LDH enzyme mixture, and 10 µl (~50 µg) of broken mitochondria were added into
970 µl of basic reaction buffer in the glass cuvette. The absorbance was blanked at this
moment, and the experiment was initiated by the addition of 5 µl of 0.5 M ATP.
Thirty seconds after the initiation of experiment, 5 µl of 70 mM NADH was added and
the activity of ATPase was recorded as the rate of NADH consumption at 340 nm (ε
340

= 6.22 mM
-1
cm
-1
).

RESULTS
Aconitase inactivation by ONOO
–
: Protective effect of citrate
Treatment of the porcine heart aconitase preparation with ONOO
–
resulted in a
dose-dependent loss of aconitase activity (Fig. 22A). In the absence of citrate, the
enzyme substrate, half-maximal inhibition was observed with ∼3 µM ONOO
–
. Citrate
significantly protected aconitase against inactivation by ONOO
–
, with ∼66-fold higher
90
levels of the oxidant (200 µM) being required to elicit a 50% inhibition of enzyme
activity. The treatment of aconitase with SIN-1, which generates a continuous flow of
ONOO
–
, similarly inactivated aconitase, with citrate having a protective effect (Fig.
22B). The difference in dose-response curves observed for ONOO
–
and SIN-1 may be
explained as a lack of linear dependence between ONOO
–
produced and SIN-1
concentration (Haddad, Crow et al. 1994). All subsequent experiments were
performed with known concentrations of ONOO
–
.
The difference in ONOO
–
concentration needed to inactivate aconitase in the absence
and presence of citrate suggests that two different sites on aconitase can be affected by
ONOO
–
: (a) the Fe-S cluster in the active site, which is sensitive to low levels of
ONOO
–
(Castro, Rodriguez et al. 1994) and is protected by citrate and (b) amino acid
residues, which are less sensitive to ONOO
–
and cannot be protected by citrate. In
order to determine if the doses of citrate required to protect aconitase from ONOO
–

were associated with the binding of citrate to the active site, a dose-response curve
was performed (Fig. 22C). The K
M
of citrate for aconitase was estimated as 33 µM
(Gardner and Fridowich 1991). The percent of the aconitase active site occupied with
citrate (%
aconitase-citrate
) is given by the equation:
%
aconitase-citrate
= [citrate]/[citrate] + K
M

91

Fig. 22. ONOO
–
-mediated inactivation of aconitase: Effect of citrate
(A) ONOO
–
-mediated inactivation of aconitase. Porcine aconitase (2 mU/ml) in 50 mM
Tris-HCl, pH 7.4, was rapidly mixed with varying amounts of ONOO
–
in the absence (λ) or
presence (µ) of 2.5 mM citrate. (B) SIN-1 mediated inactivation of aconitase. Porcine aconitase
(2 mU/ml) in 50 mM Tris-HCl, pH 7.4, was incubated with varying concentrations of SIN-1 for
10 minutes in the absence (λ) or presence (µ) of 2.5 mM citrate. To remove SIN-1, samples were
spun down in a micro bio-spin desalting column for 4 minutes at 4°C and immediately analyzed
for aconitase activity. (C) Dose response curve of citrate protection against ONOO
–
-mediated
inactivation of aconitase. Porcine aconitase (2 mU/ml) was rapidly mixed with varying amounts
of citrate and ONOO
–
(200 µM). (µ) Measured percent protection by citrate against ONOO
–
. (λ)
Estimated % of aconitase active site occupied by citrate. The percent of citrate binding to
aconitase was calculated using the equation in the text. The K
M
of aconitase for citrate was
estimated as 33 µM.
100
50
0
0.1 0.2
[ONOO
–
] (mM)
A
A
c
o
n
i
t
a
s
e

A
c
t
i
v
i
t
y

(
%
)
Aconitase Activity (%)
[Citrate] (mM)
0.25 0.5
0
100
50
%

P
r
o
t
e
c
t
i
o
n
C
% Protection
100
50
0
5 10
[SIN-1] (mM)
%
B
Aconitase Activity (%)
92

A
3250 3300 3400 3350
Magnetic Field (G)
C
D
E
B
3250 3300 3350 3400
Magnetic Field (G)
A
B
C
D
E

Fig. 23. Effect ONOO
–
on the Fe-S cluster in the active site of aconitase: EPR analysis
Active aconitase contains an EPR-silent [4Fe-4S]
2+
cluster that becomes EPR-
detectable when converted to the inactive [3Fe-4S]
1+
form. Aconitase (50 mU/ml) was
rapidly mixed with various concentrations of ONOO
–
in 100 mM Tris-HCl buffer, pH
7.5, and frozen under liquid nitrogen before measurements. Measurements were
performed at 10 K with instrument settings as described in the Materials and Methods
section. (A) Aconitase, untreated. (B) Aconitase treated with 30 µM ONOO
–
. (C)
Aconitase treated with 500 µM ONOO
–
(D) Aconitase treated with 1 mM ONOO
–
. (E)
Aconitase treated with 500 µM ONOO
–
plus citrate (50 mM).
93
The protective effects of citrate tallied its binding to the active site of aconitase
(Fig. 22C). These results confirm that citrate protects aconitase by sterically blocking
ONOO
–
access to the Fe-S cluster-containing active site.
Modification of the Fe-S cluster by ONOO
–

The modification of the Fe-S cluster in the active site of aconitase by low doses of
ONOO
–
was confirmed by EPR spectroscopy. The active [4Fe-4S]
2+
cluster of
aconitase is EPR silent, but when converted to the inactive [3Fe-4S]
1+
form, it
becomes detectable by EPR at low temperatures (Kennedy, Antholine et al. 1997).
Untreated aconitase contained a small EPR signal (Fig. 23A). Both low (30 µM) (Fig.
23B) and high (500 µM) (Fig. 23C) levels of ONOO
–
yielded an EPR signal
characteristic of aconitase in the [3Fe-4S]
1+
form. Higher ONOO
–
concentrations (> 1
mM) caused a loss of the EPR signal (Fig. 23D), thus indicating the further
degradation of the [3Fe-4S]
1+
cluster to an EPR-silent form. Because high
concentrations of aconitase (50 mU/ml) are necessary for EPR analysis, corresponding
high levels of citrate and ONOO
–
were used. Citrate protected aconitase from ONOO
–

inactivation (Fig. 23E). These results support the notion that low levels of ONOO
–

inactivated the enzyme upon modification of the [Fe-S] cluster at the active site ([4Fe-
4S]
2+
→ [3Fe-4S]
1+
) and that citrate protected the Fe-S cluster from damage. In the
presence of citrate, however, the inactivation of aconitase by ONOO
–
(Fig. 22 A,B) is
likely due to its reaction at a site other than the Fe-S cluster-containing active site.
Spin trapping of an immobilized protein thiyl radical
To assess if ONOO
–
inactivation of aconitase in the presence of citrate was due to
modification of cysteine residues, EPR in conjunction with the spin trap DMPO was
94
3440 3480 3520
D
C
B
E
A
F
Magnetic Field (G)
Fig. 24. Detection of an immobilized DMPO/cysteinyl radical-protein radical adduct after
ONOO
–
treatment
Aconitase (50 mU/ml) was rapidly mixed with various concentrations of ONOO
–
in
50 mM Tris/HCl, pH 7.5, containing 65 mM DMPO. (A) Aconitase treated with 25 µM
ONOO
–
. (B) Aconitase treated with 500 µM ONOO
–
. (C) Computer simulation of (B) (r =
0.802) assuming a
N
= 14.5 G; a
β
H
= 15.6 G; line width = 7.7 G. (D) Aconitase treated with 500
µM ONOO
–
plus 200 mM citrate. (E) 500 µM ONOO
–
in the absence of aconitase. (F)
Aconitase pretreated with 2 mM NEM (2 mM) prior to treatment of 500 µM ONOO
–
.
95
used. The interaction of thiols (RSH) with ONOO
–
(or its protonated form, which
disassociates within a cage) yields mainly thiyl radicals (RS
.
) (reaction 1). EPR
analysis with DMPO was employed to determine the formation of protein thiyl
radicals during ONOO
–
treatment. The addition of a low concentration of ONOO
–
(25
µM) did not generate an EPR signal (Fig. 24A), whereas the addition of a high
concentration of ONOO
–
(500 µM) resulted in formation of a DMPO/cysteinyl radical
spin adduct (Fig. 24B) consisting of a broad, anisotropic EPR signal: the broad lines in
the spectrum are characteristic of a slow tumbling, protein-derived radical. Fig. 24C
shows a computer-simulated spectrum of the DMPO/protein cysteinyl radical spin
adduct (a
N
= 14.5 G; a
β
H
= 15.6 G; line width = 7.7G), which matches that of the
DMPO/protein cysteinyl radical adduct of hemoglobin and myoglobin previously
reported (Maples, Jordan et al. 1988; Augusto, Menezes et al. 2002). Because the
aconitase preparation contained some other proteins, particularly albumin to stabilize
aconitase, the DMPO/cysteinyl radical spin adduct could not be solely attributed to
aconitase. However, Western blot analysis with maleimide PEO
2
-biotin demonstrated
that aconitase was one of the few thiol-containing proteins (data not shown), thus
implying that aconitase contributed significantly to the formation of the
DMPO/protein thiyl radical spin adduct. The formation of such an adduct is consistent
with a mechanism entailing oxidation of –SH moieties by ONOO
–
to yield a thiyl
radical (reaction 1). The EPR signal was not affected by high concentrations of citrate
(200 µM; Fig. 24D). In the absence of aconitase, ONOO
–
yielded a low intensity
signal probably corresponding to the spin adduct of hydroxyl radical (Fig. 24E). The
protein cysteinyl radical adduct signal was abolished upon alkylation of protein thiols
96
by NEM (Fig. 24F). Because NEM binds to all thiol residues, the loss of the EPR
signal upon NEM treatment confirms the EPR spectrum originated form a thiol
residue in the protein.
Reaction of ONOO
–
with cysteine and tyrosine residues in aconitase
Thiyl radicals generated from ONOO
–
(reaction 1) or other oxidants can decay to
stable oxidation products, such as disulfides and sulfenic acids (reaction 3); the latter
have reactive properties different from thiols. Protein thiols can be labeled using
maleimide (such as maleimide-PEO
2
-biotin; i.e., biotinylation), while oxidized
cysteine products do not bind to maleimides (i.e., cannot be biotinylated). To assess
protein thiols levels after ONOO
–
treatment, Western blot analysis was performed
using maleimide-PEO
2
-biotin. Fig. 25A shows that thiols in aconitase are lost in a
dose-dependent manner upon ONOO
–
treatment. Aconitase preparations that had not
been labeled with maleimide PEO
2
-biotin failed to show a band; this demonstrated
that no endogenous biotin group(s) existed within aconitase. The presence of citrate
did not prevent loss of thiols upon ONOO
–
treatment (data not shown), thus
strengthening the notion that cysteine oxidation occurred at sites other than the active
site.
Besides cysteine, tyrosine residues represent a major target for ONOO
–
-induced
modification (reactions 4-5). Western blot analysis using anti-nitrotyrosine antibodies
showed a dose-dependent increase in nitrotyrosine formation in aconitase with
increasing ONOO
–
levels (Fig. 25B), thus suggesting that some of the 22 tyrosyl
residues in aconitase were nitrated upon treatment with ONOO
–
. As with the protein
thiols mentioned above, citrate did not prevent protein tyrosine nitration (data not
97

020 50 100 200 500
12
6
0
A
Optical Density/mm
2
020 50 100 200
4
2
0
B
Optical Density/mm
2
[ONOO
–
] (mM)

Fig. 25. Loss of aconitase thiols and nitrotyrosine formation following ONOO
–

treatment
(A) Loss of aconitase protein thiols following ONOO
–
treatment. Porcine
aconitase (4 mU/ml) was rapidly mixed with varying amounts of ONOO
–
in 50 mM
Tris-HCl buffer, pH 7.4. The aconitase preparation was then treated with maleimide-
PEO
2
-biotin as described in the Materials and Methods section. Samples were run on an
8% SDS-Page non-reducing gel and LC/MS-MS was used to confirm the aconitase
band. Upper panel: Western blot analysis of biotinylated proteins (performed using a
streptavidin antibody); lower panel: semi-quantitative analysis (densitometry) of
Western blots. (B) Porcine aconitase (4 mU/ml) was rapidly mixed with varying
amounts of ONOO
–
in 50 mM Tris-HCl buffer, pH 7.4; samples were run on a SDS-
Page (8%) non-reducing gel. LC/MS-MS was used to confirm the aconitase band.
Upper panel: Western blot analysis for nitrotyrosine (performed using a polyclonal anti-
nitrotyrosine antibody); lower panel: semi-quantitative analysis (densitometry) of
Western blots.
98

Table IV

LC/MS/MS Analysis of Aconitase Amino Acid Modifications

Tryptic Peptide Residue Charge MSc XC ∆cn

3* 33 2.7 0.45
VAVPSTIHCDHLIEAQLGGEK Cys
126
3 30 2.3 0.50
3 25 2.1 0.34

2* 83 4.9 0.54
2 81 4.8 0.56
DINQEVYNFLATAGAK Tyr
151
2 73 4.0 0.65
2 59 4.3 0.65
2 13 2.4 0.39

2* 64 4.6 0.71
VGLIGSCTNSSYEDMGR Cys
385
2 57 4.0 0.59
2 54 4.3 0.64
2 43 4.3 0.66

2* 33 2.5 0.54
NTIVTSYNR Tyr
472
2 33 2.4 0.62
2 27 2.0 0.24

Aconitase amino acids modified upon treatment with ONOO
–
. Experiments were carried out with
4 mU/ml aconitase supplemented with 300 µM ONOO
–
. Included are the tryptic peptide
fragment sequence, peptide charge, Mascot Ions Score (MSc), Sequest XC, and ∆cn value. ∆cn
is the difference in cross correlation score between the top two candidate peptides or proteins for
a given input data file. Charge and score values are presented for each instance of identification
over the course of at least six proteomic analyses. Residues were determined from the spectra in
Fig. 5. *Indicates spectra shown in Fig. 5.
99
shown), thus suggesting that the latter occurs even when the active site of the
enzyme is occupied by its substrate and that citrate does not react with ONOO
–
.
LC/MS/MS analysis of aconitase modified by ONOO
–
treatment
LC/MS/MS analysis of aconitase following ONOO
–
treatment revealed four amino
acid modifications: oxidation of cysteines 126 and 385 to sulfonic or cysteic acid and
nitration of tyrosines 151 and 472. These results confirm the Western blot analysis that
revealed loss of cysteine residues (Fig. 25A) and nitrotyrosine formation (Fig. 25B) as
major consequences of ONOO
–
treatment. Cysteine 385 is one of three cysteine
residues that bind the Fe-S cluster in the active site of aconitase. This suggests that
aconitase may be partly inactivated by disrupting the binding of the Fe-S cluster to
cysteine 385.
Sequence coverage for aconitase in both control and ONOO
–
-treated samples was
approximately 40% in samples subjected to LC/MS/MS analysis. The four amino acid
modifications were observed over the course of analysis of at least six separate distinct
sample preparations. In each of the four cases, tryptic peptides containing the
identified modification were observed by both Mascot and Sequest analysis in at least
two or more separate sample preparations and LC/MS/MS analyses. In no instance
were these oxidative and nitrative modifications observed in control samples (not
treated with ONOO
–
). Table IV summarizes the amino acid residues modified to
cysteic acid or nitrotyrosine over the course of multiple sample analyses by Mascot
and Sequest search software.
Tyrosine 151 – Nitration of Tyrosine 151 was observed in the same doubly
charged 16 amino acid tryptic peptide five times in four separate LC/MS/MS analyses.
100

Fig. 26. MS/MS spectra for aconitase tryptic peptides carrying an oxidative/nitrative
modification
MS/MS spectra for peptides modified at (A) Cys
126
; (B) Tyr
151
; (C) Cys
385
; (D)
Tyr
472
. Charge state and identification scores are listed in Table I. Above each spectra is the
peptide sequence with observed b and y ions indicated by their fragmentation number above (y
ions) or below (b ions) the sequence. ++ or +++ indicate doubly or triply charged ions. Single
charged ions are not designated with a +. The ion annotation is based upon results presented in
Sequest's "display ions view" window that were corroborated by dta file. Y* indicates a nitro-
tyrosine, C* indicates cysteine sulfonic acid or cysteic acid.
A

B
C
D
101
Sequest (cross correlation (XC) values ranged from 2.4 to 4.9 and ∆cn values from
0.39 to 0.65 (Table IV). Retention time for the nitro-tyrosine modified peptide was
consistent among separate LC/MS/MS runs and non-modified peptide sequences were
also observed with equal or higher XC values compared to modified peptide (data not
shown). A representative MS/MS spectra (XC = 4.8) is presented in Fig. 26A. The
precursor ion was doubly charged with a mass of 900.1 Da corresponding to a 16-
residue tryptic peptide, comprising amino acids 145 – 160, with one nitration
modification. Fragment b and y ions were observed for nearly all-observable single
charged fragments and four each of doubly charged b and y ions were assigned as
well.
Cysteine 385 – Cysteine 385, one of the three cysteine residues to bind the Fe-S
cluster was oxidized to cysteic acid following ONOO
–
treatment in four separate
sample analyses by LC/MS/MS. Each identification was based on the same doubly
charged 17 amino acid tryptic peptide with consistent retention times among each
LC/MS/MS run (data not shown). Sequest XC scores ranged from 4.0 to 4.6 with ∆cn
values of 0.59 to 0.66 (Table IV). A representative tandem MS/MS spectra (XC =
4.6) is shown in Fig. 27B. Cysteic acid at position 385 observed in the doubly charged
tryptic peptide corresponded to 17 amino acids, 379 – 395, of porcine aconitase. The
precursor ion mass of 927.7 Da correlated with the cysteic acid modification and also
a single oxidation at methionine 393, a commonly observed modification in tandem
MS/MS analyses. The MS/MS collision-induced fragmentation pattern (Fig. 26B)
allowed near complete assignment of all observable single charged b and y fragment
ions and also included instances of doubly charged b and y ions.
102
A
Tyr
151
Tyr
472
Cys
385
Cys
126
Cys
451
Cys
448
Citrate
Iron
Sulfur
B

Fig. 27. Structure of aconitase and amino acid modifications elicited by ONOO
–

(A) The full aconitase structure is illustrated in ribbon format. Amino acid
residues that compose the active site are shown in stick format in red. Key molecules are
illustrated in space-filling format: components of the Fe-S cluster (Fe shown in metallic
green and sulfur shown in yellow), cysteine residues ligated to the Fe-S cluster (red), and
citrate (orange). Cysteine residues oxidized to sulfonic acid (blue) (including modified
cysteine 385 that binds the Fe-S cluster), and tyrosine residues converted to 3-
nitrotyrosine (purple) by ONOO
–
treatment are also shown in space filling format. (B)
Close up view of the active site with same illustrative scheme as in (A) except that citrate
is displayed in stick configuration. The position of the view differs from that of (A) for
the sake of clarity.
103
Tyrosine 472 – Nitration of tyrosine 472 was observed in three doubly charged
nine amino acid tryptic peptides in two separate sample analyses by LC/MS/MS.
Sequest XC values for the double charged peptide ranged from 2.0 to 2.5 with ∆cn
values from 0.24 to 0.62 (Table IV). Retention times for the peptide were consistent
among different LC/MS/MS runs. A representative MS/MS spectra is shown in Fig.
26C (XC = 2.5). Precursor mass corresponding to the nitro-tyrosine modified double
charged tryptic fragment comprising aconitase residues 466 – 474 was 557.2 Da. The
collision induced ion fragmentation pattern included all observable single charged b
and y ions; two double charged b and three double charged y ions were also observed.
Cysteine 126 – Modification of cysteine to cysteic acid at position 126 was
observed three times, each in a separate LC/MS/MS analysis, in a triple charged 21
amino acid length tryptic peptide that corresponded to aconitase residues 118 – 138.
The Sequest XC- and ∆cn values ranged from 2.1 to 2.7 and 0.34 to 0.50, respectively.
Retention times were consistent among different LC/MS/MS runs. The XC values are
relatively low, with an XC value of 3 generally being considered the cut-off for
assigning an identification of a triple charged peptide. A representative MS/MS
spectra is presented in Fig 26D (XC = 2.7). Precursor ion for the triple charged peptide
was 756.2 Da. The collision induced fragmentation pattern shows a highly biased
fragmentation where two major peaks dominate the spectra that correspond to double
and triple charged y18 with the next two highest peaks corresponding to the double
and triple charged y19. This biased fragmentation pattern where the double and triple
charged y18 ions were by far the largest peaks with the y19 double and triple ions
being the next highest set was consistently observed in several LC/MS/MS runs.
104
Despite the abundance of the y18 and y19 ions, almost all of the observable y ions
were assigned in either single or double charged state (sometimes both) and nine of 21
total possible b ions were observed in either double or triple charged state. The
cysteine modification was observed in every possible double charged y ion that would
carry the modified cysteine (i.e. y11 – y20).
Modulation of aconitase activity by GSSG
Cysteine and tyrosine residues in aconitase were affected by ONOO
–
and these
modifications were associated with a decrease of aconitase activity. It may be
hypothesized that cysteine thiol modifications other than oxidation to sulfonic acid
(Fig. 26) may similarly decrease aconitase activity. An important physiological thiol
modification is glutathionylation of proteins thiols by GSSG through disulfide-thiol
exchange (Gilbert 1982; Thomas, Poland et al. 1995) (reaction 6).
Protein-SH + GSSG → Protein-SSG + GSH [6]
Incubation of aconitase with GSSG in the presence of citrate led to a loss in
enzyme activity (Fig. 28A). Maximal inhibition (35%) was observed with ~200 µM
GSSG. The loss of aconitase activity caused by GSSG corresponded with a decrease
in protein free thiols (as measured by Western blot analysis after maleimide PEO
2
-
biotin labeling (Fig. 28B), as expected upon protein thiol glutathionylation. The
glutathionylation of aconitase was confirmed by Western blot analysis using a
monoclonal antiglutathione antibody (Fig. 28C), thus showing that GSSG
glutathionylates aconitase in a dose-dependent manner. DTT, which reduces disulfide
bonds to the corresponding thiols, fully reversed the loss of aconitase activity brought
about by GSSG (Fig. 28A).
105

[GSSG] (mM)
0.2 0 0.1 0.5 1.0
0
3
2
1
Optical Density/mm
2
0.1 0 0.05 0.5 1.0 0.2
0
6
12
Optical Density/mm
2
[GSSG] (mM)
BC
2
0.1
0
0.2
50
100
% Activity
A
[GSSG] (mM)
Fig. 28. Effect of GSSG on aconitase activity and aconitase protein thiols. Glutathionylation
of aconitase
(A) Effect of GSSG on aconitase activity. Various concentrations of GSSG were
mixed with aconitase (2 mU/ml) in 50 mM Tris-HCl, pH 7.4 in the presence of 2.5 mM
citrate for 10 min. (µ) Aconitase plus GSSG; (λ) Aconitase plus GSSG plus DTT. In the
DTT-treated samples, aconitase was incubated with GSSG and citrate followed by a 10-min
treatment with DTT (10 mM). (B) Western blots showing the loss of aconitase protein thiols
following treatment with GSSG and the corresponding semi-quantitative analysis
(densitometry). (C) Porcine aconitase (4 mU/ml) was rapidly mixed with varying amounts of
GSSG in 50 mM Tris-HCl, pH 7.4, for 10 min. Western blots showing glutathionylation of
aconitase, which was determined using an anti-glutathione monoclonal antibody as
described in the Experimental Procedures section, and the corresponding semi-quantitative
analysis (densitometry).
106
These results demonstrate that aconitase activity can be modulated by
glutathionylation and suggest that modification of cysteine residues outside the active
site of aconitase can result in an impairment of enzyme activity.
Glutathionylation of ATP synthase results in the loss of activity
In section 1, immunoprecipitation followed by LC/MS/MS was used to
identify the mitochondrial proteins that were glutathionylated. The 70 kd band was
identified as a protein similar to 3-oxoacid CoA transferase (gi 34854196, 68.260kd),
which plays a role in ketone body metabolism. The 60 kd band was identified as ATP
synthase, H+ transporting, mitochondrial F1 complex, α subunit, isoform 1 (gi
40538742, 59.831 kd) (Fig 8B). The purified F1 sector is made up of five different
subunits with a stoichiometry of α
3
β
3
γδε (Amzel, McKinney et al. 1982; Bianchet,
Ysern et al. 1991). One of the cysteine residues in the α subunit is close both spatially
and in sequence to the glycine-rich loop, which is thought to be involved in binding
one or more of the ATP phosphates (Bianchet, Ysern et al. 1991).
To determine the functional consequence of glutathionylation ATP synthase
(complex V), its activity was determined by uncoupling mitochondria and measuring
its ATPase activity. ATPase activity of complex V was assayed by coupling its
activity to pyruvate kinase and lactate dehydrogenase. The hydrolysis of ATP by
ATPase was charged to the conversion of phosphoenolpyruvate to pyruvate by
pyruvate kinase. Pyruvate was then converted to lactate by lactate dehydrogenase with
the consumption of NADH. Therefore, the consumption of NADH, monitored by
spectrophotometer at absorbance 340 nm, can be used as an index of ATPase activity.
DPG isolated mitochondria were incubated at 37
o
C for 10 minutes with and without
107
substrates in order to glutathionylate or de-glutathionylate the alpha subunit of the
ATP synthase protein. Upon completion of the appropriate treatment and incubation,
mitochondria were either broken or converted into SMPs, in order to expose the
enzyme, and monitored for the ATPase activity.
The ATPase activity from broken mitochondria and SMPs were 0.9 + 0.2
(glutathionylated) and 2.3 + 0.1 (de-glutathionylated) nmol/mg/s and 9.7 + 0.8
(glutathionylated) and 31.1 + 0.2 (de-glutathionylated) nmol/mg/s of NADH
consumption respectively. De-glutathionylation of ATP synthase had a profound
effect, as the ATPase activity increased by 174% and 221% in broken mitochondria
and SMPs (Table V).
Table V

ATPase Activity: NADH consumption (nmols/mg/s)

Mitochondria Activity % Change
Broken
Control 0.9 + 0.2
Substrates 2.3 + 0.1 174

Sub-Mitochondrial Particles
Control 9.7 + 0.8
Substrates 31.1 + 0.2 221
Freshly isolated intact rat brain mitochondria were treated with or without substrates
(2.4 mM glutamate, 2.4 mM malate, and 0.7 mM ADP). Mitochondria were either
broken or converted into SMPs. SMPs were made by sonicating intact mitochondria
6 times for a total of 1 minute (10 seconds per sonication). ATPase activity of
mitochondrial complex V was assayed by coupling its activity to pyruvate kinase and
lactate dehydrogenase and monitoring NADH consumption.
108
Discussion
Aconitase activity is frequently viewed as a marker of oxidative stress in
biological systems because the enzyme [4Fe-4S]
2+
cluster is readily inactivated by O
2
.–
, H
2
O
2
, and ONOO
–
. This study puts forward two mechanisms by which the Fe-S
cluster in the active site could be modified by low levels of ONOO
–
: (a) conversion
the active Fe-S cluster from the [4Fe-4S]
2+
form to the inactive [3Fe-4S]
1+
form with
loss of labile iron (as confirmed by low temperature EPR analyses); (b) disrupting the
binding of the Fe-S cluster to cysteine residues (cysteine 385) of the protein through
oxidation of cysteine to cysteic acid (as shown by LC/MS/MS analyses). However,
whether the disruption in Fe-S binding to cysteine causes Fe-S degradation or the
breakdown of the Fe-S cluster into the inactivate [3Fe-4S]
1+
form remains to be
elucidated.
Binding of citrate to the aconitase active site selectively protected against
modifications of the Fe-S cluster by ONOO
–
, but this protection was not extended to
amino acid residues (distal from- or in close proximity to the active site). The
mechanism by which citrate protects the Fe-S cluster from ONOO
–
probably involves
inhibition of the entry of ONOO
–
to the Fe-S cluster upon citrate binding. In this
regard, recent X-ray crystallography studies have shown that binding of citrate induces
a conformational change of the aconitase active site (Lauble and Stout 1995). Hence,
citrate may shield the Fe-S cluster against ONOO
–
attack by (a) sterically hindering
access of reactive molecules to Fe-S cluster or (b) causing a conformational shift of
the active site that makes the Fe-S cluster less accessible to reactive molecules. It is
likely that a combination of both mechanisms contributes to the effectiveness of citrate
109
protection of the Fe-S cluster from oxidant attack. Whichever the mechanism
underlying the protection by citrate, it may be surmised that citrate levels in the
mitochondrial matrix will determine the site of ONOO
–
attack on aconitase by shifting
ONOO
–
reactions from Fe-S clusters to cysteine, tyrosine, and other amino acid
residues of aconitase. Physiological levels of citrate in the matrix are determined by
the metabolic state of mitochondria, but in bacteria citrate levels have been estimated
in the millimolar range (Lowry, Carter et al. 1971). If similar levels were to apply to
mammalian mitochondria, the Fe-S cluster of aconitase may not be as vulnerable to
oxidants as previously reported.
In the presence of citrate, the inhibitory effect of ONOO
–
suggested the occurrence
of sensitive sites other than the Fe-S cluster-containing active site in the regulation of
aconitase activity. LC/MS/MS analysis revealed that cysteine 385 (one of three
cysteine residues that bind the Fe-S cluster in the active site of aconitase) was oxidized
to cysteic acid and that three other amino acids (tyrosine 151 and 472, and cysteine
126) outside the active site that were modified upon ONOO
–
treatment. The other two
cysteines involved in cluster ligation are cysteines 448 and 451 (41). The crystal
structure of porcine aconitase (Robbins and Stout 1989; Lauble, Kennedy et al. 1992)
indicates that all three modified amino acids (tyrosine 151 and 472, and cysteine 126)
were adjacent to amino acids comprising the active site (Fig. 7A and B). Tyrosine 472,
for example, lies as close as 3.86 Å to cysteine 451, which binds the Fe-S cluster (Fig.
7B). Nitration of tyrosine 472 can greatly crowd the active site, thus the catalytic
activity of aconitase. Thus, modifications to tyrosine 151 and 472, and cysteine 126,
because of their close proximity to the active site, may cause conformational changes
110
that destabilize the active site. The oxidative reactions by ONOO
–
can take place
by both one- and two-electron mechanisms (sequences in reaction 7 and 8,
respectively) (Koppenol, Moreno et al. 1992; Goldstein and Czapski 1995). The
oxidation of cysteine residues (R-SH) in aconitase to sulfonic acid (R-SO
3
H) involves
oxidation steps beyond the initial formation of sulfenic acid (R-SOH) (reactions 1-3):
these steps are likely to encompass another stable product, such as sulfinic (R-SO
2
H)
(reaction 7). Alternatively, the thiyl radical (R-S
.
) formed in reaction 1, can undergo
further ONOO
–
-mediated one-electron oxidations and reactions with molecular
oxygen leading to cysteine sulfenic acid formation (sequence in reaction 8) (Harman,
Mottley et al. 1984) followed by similar reactions to sulfinic and sulfonic products.
R-SH → R-SOH → R-SO
2
H → R-SO
3
H [7]
R-SH → R-S
.
→ R-SOH → R-S
.
O → R-SO
2
H → R-S
.
O
2
→ R-SO
3
H [8]
These conformational changes may account for the ONOO
–
-mediated decrease in
aconitase activity observed in the presence of citrate. Nonetheless, it remains to be
determined whether there is one critical modification involved in the modulation of
aconitase activity or whether all modifications contribute to loss of aconitase activity.
However, based on the sensitivity of aconitase to glutathionylation, cysteine
modifications appear to be more critical than tyrosine modifications in modulating
aconitase activity. Loss of aconitase after GSSG treatment suggests glutathionylation
of one or more cysteine residues may cause conformational changes that affect
enzyme activity and thiols may be required to stabilize a protein structure associated
with function.
111
This study provides the first evidence that aconitase and ATP synthase are
susceptible to glutathionylation and that this process may modulate their activities.
These findings suggest that glutathionylation may be an important means of
modulating enzyme activity under oxidative and nitrosative stress. The mitochondrial
matrix contains high levels of GSH, which is believed to scavenge ONOO
–
, and
protect sensitive enzymes such as aconitase and ATP synthase. Yet, GSSG is the
major end product (Radi, Beckman et al. 1991) originating from the reaction between
GSH and ONOO
–
(reaction 9-11), thus providing a reactant for protein mixed
disulfide formation (reaction 6).
GSH + [HO
.

…
NO
2
.
] → GS
.
+
.
NO
2
+ OH
–
[9]
GS
–
+ GS
.
→ GSSG
.–
[10]
GSSG
.–
+ O
2
→ GSSG + O
2
.–
[11]
ONOO
–
formation in the mitochondrial matrix may therefore modulate aconitase
activity in two ways: (a) directly, upon modifications of the Fe-S cluster and/or critical
amino acids residues, and/or (b) indirectly, through oxidation of mitochondrial GSH to
GSSG, which in turn can lead to glutathionylation of exposed/active cysteines.
Because of the high levels of GSH in mitochondria, glutathionylation would appear a
major physiological consequence of ONOO
–
formation in the mitochondrial matrix.
Mitochondrial GSH may have other functions, for it was reported that the recovery of
aconitase activity ([3Fe-4S]
1+
→ [4Fe-4S]
2+
) was accomplished in vivo by a process
involving GSH (Gardner and Fridovich 1993) or the iron chaperone protein frataxin
(Bulteau, O'Neill et al. 2004).
112
It may be surmised that [GSH]:[GSSG] ratios in the mitochondrial matrix are
important factors that determine aconitase and ATP synthase functionality. GSSG
levels are tightly regulated in mitochondria and GSSG is rapidly reduced to GSH by
glutathione reductase. This indicates that aconitase and ATP synthase activity is finely
regulated by the mitochondrial metabolic and redox states, which determine citrate
and NADPH levels, the latter closely tied to GSSG level regulation. Reactive oxygen-
and nitrogen species levels, citrate concentration, and the [GSH]:[GSSG] ratio may be
intrinsically involved in the regulation of aconitase and ATP synthase activity in
mitochondria and the sites and mechanisms of inactivation will be determined by the
interaction of these factors. Thus, the metabolic and thiol redox status of the
mitochondrial matrix may play an important role in the physiological regulation of
aconitase and ATP synthase activity.

113
Chapter 5: Conclusion
The relationship between cellular viability and redox status is such that when a
cell becomes oxidized, the cell progresses from proliferation, differentiation, apoptosis
and finally to necrosis. Though this concept is well established, the actual mechanisms
through which the redox status control cellular and mitochondrial function is unclear.
In this work we examined the hypothesis that changes in the cell redox status –
elicited by
.
NO overproduction and dopamine autoxidation – affect mitochondrial and
cellular function through specific and critical protein modifications leading to
apoptotic cell death. This hypothesis was tested through three experimental
approaches directed at (1) evaluating the mitochondrial redox status as a function of
mitochondrial GSH content and metabolism, (2) assessing the effects of
.
NO and
dopamine, alone or synergistically on the cell and mitochondrial respiration, and (3)
identifying the protein post-translational modifications arising from changes in the
redox status and its significance for mitochondrial and cell function. The above
hypothesis is tenable in view of the following evidence in this dissertation
1. Mitochondrial Redox Status – Mitochondria maintain a distinct pool of GSH
and the mechanism of GSH modulation within the mitochondria remains
unclear. Changes in the extra-mitochondrial redox environment can be
modeled, through different isolation methods and buffer constituents. Exposure
to severe oxidative stress promoted glutathionylation of mitochondrial
proteins. This post-translational modification was removed upon complex I
substrate supplementation and correlated with an increase in GSH content in
isolated mitochondria. This increase could not be accounted solely from the
114
reduction of GSSG by the glutathione reductase enzyme into GSH. This
suggests that the mitochondrial redox environment and its energy status are
closely associated. The released GSH by substrate supplementation has
physiological significance as it increased the ability of mitochondria to buffer
against different doses of H
2
O
2
. Two novel concepts are presented from the
data collected in this section (i) that mitochondria can respond/ adapt to
changes in their extra mitochondrial environment by regulating its own GSH
pool or vice versa and (ii) mitochondrial energy production is closely
associated with the redox environment of the mitochondria. Placed within a
cellular context, the latter finding is of significance as two important factors
that define cell viability, energy status and redox environment, modulate one
another.
2. Dopamine,
.
NO, and Mitochondrial Dysfunction – The results presented
demonstrate that dopamine oxidation in the presence of
.
NO generated ONOO
-
and caused cellular and mitochondrial dysfunction. Dopamine and
.
NO are
modifiers of the redox environment in Parkinson’s disease. Their interactions
resulted in the generation ONOO
-
, a potent oxidant. GSH, an antioxidant,
prevented the oxidation of dopamine by reducing dopamine semiquinone back
to dopamine. GSH has many protective functions, the data presented in this
dissertation demonstrates other potential protective effects in PD which can be
achieved through these three mechanisms; (i) maintenance of dopamine
concentrations, (ii) preventing the formation of toxic oxidation products
(dopamine quinone and neuromelanin) and (iii) prevention ONOO
-
formation,
115
as O
2
._
generation was decreased. This protection of GSH in PD, not only
highlights the significance of this molecule but also provides a possible
mechanism for GSH depletion in dopaminergic neurons resulting in an
oxidized cellular redox environment and susceptibility to cell death
(apoptosis). Dopamine and
.
NO synergistically inhibited in situ PC12 cell
respiration and mitochondrial respiration in isolated brain mitochondria by
~72%. The synergism was in part mediated by ONOO
-
as SOD was able to
partially rescue mitochondrial respiration. This study, for the first time, show
evidence that the synergistic effects of dopamine and
.
NO is mediated through
the formation of their reaction product ONOO
-
. Increased reactive oxygen and
nitrogen species are not only determinants of cellular and mitochondrial redox
status (GSH:GSSG couple) and respiration, but also critical in modulating
chemical post-translational protein modifications.
3. Protein Post-Translational Modifications – Nitration, cysteine oxidation, and
glutathionylation are protein post-translational modifications that resulted from
the direct or indirect interaction with amino acid residues and ONOO
-
. These
modifications are the link between an oxidized cellular redox environment,
dysfunction of cellular/mitochondrial respiration, and cell death. ONOO
-
can
induce tyrosine nitration and cysteine oxidation directly to the protein or
glutathionylation indirectly by decreasing the GSH:GSSG ratio, which results
in an increase in GSSG and mixed disulfide formation. ONOO
-
dose-
dependently decreased aconitase activity, and resulted in the nitration of
tyrosine151 and 472 and oxidation to sulfonic acid of cysteine 126 and 385. In
116
the presence of citrate, a substrate for aconitase, a 66-fold higher
concentration of ONOO
-
was required for half maximal inhibition. When
exposed to an oxidized redox environment (high GSSG content), to test the
indirect interactions of ONOO
-
, aconitase and ATP synthase were post-
translationally modified by glutathionylation. Glutathionylation had a similar
effect on aconitase, as it too inhibited protein activity. ATP synthase was also
susceptible to glutathionylation. Protein sulfhydryl recovery (de-
glutathionylation) resulted in a 221% increase in protein activity. Thus taken
together, reactive oxygen- and nitrogen species levels, citrate concentration,
and the [GSH]:[GSSG] ratio may be intrinsically involved in the regulation of
the cells energy status through the modulation of aconitase and ATP synthase
activity. These proteins are key in energy production; a compromise in their
activity through these modifications will cause an energy crisis. In order for a
healthy neuron to function properly, it must consume large quantities of ATP
as neurotransmission is dependent on constant maintenance of an
electrochemical gradient by the Na
+
/K
+
ATP dependent transport pumps. A
compromise in aconitase and ATP synthase activity may thus cause neuronal
dysfunction and eventually apoptosis.

117
Chapter 6: Future Perspectives
The data presented in this dissertation are centered on the biochemistry of
redox changes in mitochondria, the chemistry of
.
NO and dopamine, and the post-
translational modifications of redox sensitive mitochondrial proteins. The data
amassed through these three specific aims provide a good foundation for further
studies into the Parkinson’s disease model and other neurodegenerative diseases,
where impairment of mitochondrial function is an early event. It would be interesting
to further investigate the effects of dopamine and NO in a PC12 model with respect to
changes in the cytosolic as well mitochondria GSH, GSSG concentrations and
quantify the changes in their respective Nernst potential. The models utilized in this
proposal adopt transient changes or alterations in the extra mitochondrial and cellular
redox environment. A cellular based model will enable us to investigate these redox
and energy changes under chronic oxidative stress. This can be achieved using a co-
culture method already established in our laboratory where PC12 cells can be co-
cultured with microglia, a physiological source of
.
NO. This model would be an
exciting tool as it is more physiological and would enable us to study redox and
energy changes that occur over extended periods of time (48hrs).
Moving into a cell-based model will enable us to assess redox changes in the
presence of
.
NO and dopamine on a whole cell model. It would be important to
identify potential cytosolic or nuclear proteins that might be glutathionylated or
nitrated in the presence of
.
NO and dopamine. It has been recently demonstrated that
.
NO itself can lead to selective glutathionylation of proteins (West, Hill et al. 2006).
In the presence of
.
NO, cysteinyl groups in proteins typically become nitrosylated.
118
However, in many instances where S-nitrosylation has been described, the protein
cysteines are also oxidized resulting in the subsequent formation of disulfide bonds (S-
thiolation). In vivo, the most abundant protein thiol is glutathione and exists in
milimolar concentration in the cytoplasm. Hence, S-nitrosylation is likely to promote
S-glutathionylation (Martinez-Ruiz and Lamas 2004). Identifying additional proteins
that might be glutathionylated or nitrated might provide additional information of
mitochondrial-cytosol or mitochondrial-nucleus crosstalk that is modulated by discrete
redox sensitive signaling pathways. Additionally, it would be interesting to assess the
therapeutic effects of GSH repleating agents such as N-Acetyl cysteines or redox
modulators such as alpha-lipoic acid in a PD model.

119
References
Amzel, L. M., M. McKinney, et al. (1982). "Structure of the mitochondrial F1 ATPase
at 9-A resolution." Proc Natl Acad Sci U S A 79(19): 5852-6.

Anderson, M. F. and N. R. Sims (2000). "Improved recovery of highly enriched
mitochondrial fractions from small brain tissue samples." Brain Res Brain Res
Protoc 5(1): 95-101.

Antunes, F., A. Boveris, et al. (2004). "On the mechanism and biology of cytochrome
oxidase inhibition by nitric oxide." Proc. Natl. Acad. Sci. USA 101: 16774-
16779.

Antunes, F. and E. Cadenas (2001). "Cellular titration of apoptosis with steady state
concentrations of H(2)O(2): submicromolar levels of H(2)O(2) induce
apoptosis through Fenton chemistry independent of the cellular thiol state."
Free Radic Biol Med 30(9): 1008-18.

Augusto, O., L. d. Menezes, et al. (2002). "EPR detection of glutathionyl and
hemoglobin-cysteinyl radicals during the interaction of peroxynitrite with
human erythrocytes." Biochemistry 41: 14323-8.

Bains, J. S. and C. A. Shaw (1997). "Neurodegenerative disorders in humans: the role
of glutathione in oxidative stress-mediated neuronal death." Brain Res Brain
Res Rev 25(3): 335-58.

Barker, J. E., J. P. Bolanos, et al. (1996). "Glutathione protects astrocytes from
peroxynitrite-mediated mitochondrial damage: implications for
neuronal/astrocytic trafficking and neurodegeneration." Dev Neurosci 18(5-6):
391-6.

Beckman, J. and W. H. Koppenol (1996). "Nitric oxide, superoxide, and peroxynitrite:
The good, the bad, and the ugly." Am. J. Physiol. 261: H590-H597.

Beer, S. M., E. R. Taylor, et al. (2004). "Glutaredoxin 2 catalyzes the reversible
oxidation and glutathionylation of mitochondrial membrane thiol proteins:
implications for mitochondrial redox regulation and antioxidant DEFENSE." J
Biol Chem 279(46): 47939-51.

Beinert, H. and M. C. Kennedy (1993). "Aconitase, a two-faced protein: enzyme and
iron regulatory factor." Faseb J. 7(15): 1442-9.

Bharath, S., M. Hsu, et al. (2002). "Glutathione, iron and Parkinson's disease."
Biochem Pharmacol 64(5-6): 1037-48.

120

Bianchet, M., X. Ysern, et al. (1991). "Mitochondrial ATP synthase. Quaternary
structure of the F1 moiety at 3.6 A determined by x-ray diffraction analysis." J
Biol Chem 266(31): 21197-201.

Bizouarn, T., O. Fjellstrom, et al. (2000). "Proton translocating nicotinamide
nucleotide transhydrogenase from E. coli. Mechanism of action deduced from
its structural and catalytic properties." Biochim Biophys Acta 1457(3): 211-28.

Bolanos, J. P., S. J. Heales, et al. (1996). "Nitric oxide-mediated mitochondrial
damage: a potential neuroprotective role for glutathione." Free Radic Biol Med
21(7): 995-1001.

Boveris, A. and E. Cadenas (1975). "Mitochondrial production of superoxide anions
and its relationship to the antimycin insensitive respiration." FEBS Lett 54(3):
311-4.

Boveris, A., E. Cadenas, et al. (1976). "Role of ubiquinone in the mitochondrial
generation of hydrogen peroxide." Biochem J 156(2): 435-44.

Boveris, A. and B. Chance (1973). "The mitochondrial generation of hydrogen
peroxide. General properties and effect of hyperbaric oxygen." Biochem J
134(3): 707-16.

Boveris, A., L. E. Costa, et al. (1999). "Regulation of mitochondrial respiration by
adenosine diphosphate, oxygen, and nitric oxide." Methods Enzymol 301: 188-
98.

Boveris, A., N. Oshino, et al. (1972). "The cellular production of hydrogen peroxide."
Biochem J 128(3): 617-30.

Brookes, P. S., A.-L. Levonen, et al. (2002). "Mitochondria: Regulators of signal
transduction by reactive oxygen and nitrogen species." Free Radic. Biol. Med.
33: 755-764.

Brown, G. (2000). "Energy, life, and death." The Biochemist 22: 11-15.

Brown, G. C. (1997). "Nitric oxide inhibition of cytochrome oxidase and
mitochondrial respiration: implications for inflammatory, neurodegenerative
and ischaemic pathologies." Mol. Cell Biochem. 174: 189-192.

Bulteau, A. L., M. Ikeda-Saito, et al. (2003). "Redox-dependent modulation of
aconitase activity in intact mitochondria." Biochemistry 42: 14846-14855.

121
Bulteau, A. L., H. A. O'Neill, et al. (2004). "Frataxin acts as an iron chaperone
protein to modulate mitochondrial aconitase activity." Science 305: 242-245.

Cadenas, E. (2004). "Mitochondrial free radical production and cell signaling." Mol
Aspects Med 25(1-2): 17-26.

Cadenas, E. and A. Boveris (1980). "Enhancement of hydrogen peroxide formation by
protophores and ionophores in antimycin-supplemented mitochondria."
Biochem. J. 188: 31-37.

Cadenas, E., A. Boveris, et al. (1977). "Production of superoxide radicals and
hydrogen peroxide by NADH-ubiquinone reductase and ubiquinol-cytochrome
c reductase from beef-heart mitochondria." Arch Biochem Biophys 180(2):
248-57.

Cadenas, E., J. J. Poderoso, et al. (2001). "Analysis of the Pathways of Nitric Oxide
Utilization in Mitochondria." Free Radic. Res. 33(6): 747-756.

Carballal, S., R. Radi, et al. (2003). "Sulfenic acid formation in human serum albumin
by hydrogen peroxide and peroxynitrite." Biochemistry 42: 9906-14.

Castro, L., M. Rodriguez, et al. (1994). "Aconitase is readily inactivated by
peroxynitrite, but not by its precursor, nitric oxide." J Biol Chem 269(47):
29409-15.

Castro, L., M. Rodriguez, et al. (1994). "Aconitase is readily inactivated by
peroxynitrite, but not by its precursor, nitric oxide." J. Biol. Chem. 269: 29409-
29415.

Chinta, S. J. and J. K. Andersen (2006). "Reversible inhibition of mitochondrial
complex I activity following chronic dopaminergic glutathione depletion in
vitro: implications for Parkinson's disease." Free Radic Biol Med 41(9): 1442-
8.

Colell, A., O. Coll, et al. (2001). "Tauroursodeoxycholic acid protects hepatocytes
from ethanol-fed rats against tumor necrosis factor-induced cell death by
replenishing mitochondrial glutathione." Hepatology 34(5): 964-71.

Colell, A., C. Garcia-Ruiz, et al. (1998). "Selective glutathione depletion of
mitochondria by ethanol sensitizes hepatocytes to tumor necrosis factor."
Gastroenterology 115(6): 1541-51.

Coll, O., A. Colell, et al. (2003). "Sensitivity of the 2-oxoglutarate carrier to alcohol
intake contributes to mitochondrial glutathione depletion." Hepatology 38(3):
692-702.
122

Cummings, B. S., R. Angeles, et al. (2000). "Role of voltage-dependent anion
channels in glutathione transport into yeast mitochondria." Biochem Biophys
Res Commun 276(3): 940-4.

Darley-Usmar, V. M., Rickwood, D., and Wilson, M.T. (1987). Mitochondria:a
practical approach: 109-113.

Dehmer, T., J. Lindenau, et al. (2000). "Deficiency of inducible nitric oxide synthase
protects against MPTP toxicity in vivo." J Neurochem 74(5): 2213-6.

DeLeve, L. D. and N. Kaplowitz (1991). "Glutathione metabolism and its role in
hepatotoxicity." Pharmacol Ther 52(3): 287-305.

Dexter, D. T., J. Sian, et al. (1994). "Indices of oxidative stress and mitochondrial
function in individuals with incidental Lewy body disease." Ann Neurol 35(1):
38-44.

Dhanbhoora, C. M. and J. R. Babson (1992). "Thiol depletion induces lethal cell
injury in cultured cardiomyocytes." Arch Biochem Biophys 293(1): 130-9.

Dionisi, O., T. Galeotti, et al. (1975). "Superoxide radicals and hydrogen peroxide
formation in mitochondria from normal and neoplastic tissues." Biochim
Biophys Acta 403(2): 292-300.

Drapier, J. C. and J. B. Hibbs (1996). "Aconitases: a class of metalloproteins highly
sensitive to nitric oxide synthesis." Methods Enzymol. 269: 26-36.

Duling, D. R. (1994). "Simulation of multiple isotropic spin-trap EPR spectra." J.
Magnetic Resonance Series B 104: 105-110.

Ehrhart, J., M. Gluck, et al. (2002). "Functional glutaredoxin (thioltransferase) activity
in rat brain and liver mitochondria." Parkinsonism Relat Disord 8(6): 395-400.

Elfering, S. L., T. M. Sarkela, et al. (2002). "Biochemistry of mitochondrial nitric-
oxide synthase." J. Biol. Chem. 277: 38079-38086.

Ellman, G. L. (1959). "Tissue sulfhydryl groups." Arch Biochem Biophys 82(1): 70-7.

Fernandez-Checa, J. C., C. Garcia-Ruiz, et al. (1991). "Impaired uptake of glutathione
by hepatic mitochondria from chronic ethanol-fed rats. Tracer kinetic studies in
vitro and in vivo and susceptibility to oxidant stress." J Clin Invest 87(2): 397-
405.
123

Fernando, M. R., J. M. Lechner, et al. (2006). "Mitochondrial thioltransferase
(glutaredoxin 2) has GSH-dependent and thioredoxin reductase-dependent
peroxidase activities in vitro and in lens epithelial cells." Faseb J 20(14): 2645-
7.

Finkel, T. and N. J. Holbrook (2000). "Oxidants, oxidative stress and the biology of
ageing." Nature 408(6809): 239-47.

Forno, L. S. (1996). "Neuropathology of Parkinson's disease." J Neuropathol Exp
Neurol 55(3): 259-72.

Foster, M. W. and J. S. Stamler (2004). "New insights into protein S-nitrosylation.
Mitochondria as a model system." J Biol Chem 279(24): 25891-7.

Garcia-Ruiz, C., A. Colell, et al. (1995). "Role of oxidative stress generated from the
mitochondrial electron transport chain and mitochondrial glutathione status in
loss of mitochondrial function and activation of transcription factor nuclear
factor-kappa B: studies with isolated mitochondria and rat hepatocytes." Mol
Pharmacol 48(5): 825-34.

Garcia-Ruiz, C., A. Morales, et al. (1995). "Evidence that the rat hepatic
mitochondrial carrier is distinct from the sinusoidal and canalicular
transporters for reduced glutathione. Expression studies in Xenopus laevis
oocytes." J Biol Chem 270(27): 15946-9.

Gardner, P., G. Costantino, et al. (1997). " Nitric oxide sensitivity of the aconitase." J.
Biol. Chem. 272: 25071-25076.

Gardner, P. and I. Fridowich (1991). "Superoxide sensitivity of the Escherichia coli
aconitase." J. Biol. Chem. 266(29): 19328-19333.

Gardner, P. R. and I. Fridovich (1992). "Inactivation-reactivation of aconitase in
Escherichia coli. A sensitive measure of superoxide radical." J. Bio. Chem.
267(13): 8757-63.

Gardner, P. R. and I. Fridovich (1993). "Effect of glutathione on aconitase in
Escherichia coli." Arch. Biochem. Biophys. 301(1): 98-102.

Ghafourifar, P. and E. Cadenas (2005). " Mitochondrial nitric oxide synthase." Trends
Pharmacol. Sci. 26: 190-195.

124

Gilbert, H. F. (1982). "Biological disulfides: the third messenger? Modulation of
phosphofructokinase activity by thiol/disulfide exchange." J. Biol. Chem. 257:
12086-91.

Gilbert, H. F. (1984). "Redox control of enzyme activities by thiol/disulfide
exchange." Methods Enzymol 107: 330-51.

Gladyshev, V. N., A. Liu, et al. (2001). "Identification and characterization of a new
mammalian glutaredoxin (thioltransferase), Grx2." J Biol Chem 276(32):
30374-80.

Goldstein, S. and G. Czapski (1995). "Direct and indirect oxidation by peroxynitrite."
Inorg. Chem. 34: 4401-4048.

Good, P. F., A. Hsu, et al. (1998). "Protein nitration in Parkinson's disease." J
Neuropathol Exp Neurol 57(4): 338-42.

Greenamyre, J. T., R. Betarbet, et al. (2003). "The rotenone model of Parkinson's
disease: genes, environment and mitochondria." Parkinsonism Relat Disord 9
Suppl 2: S59-64.

Greenamyre, J. T., T. B. Sherer, et al. (2001). "Complex I and Parkinson's disease."
IUBMB Life 52(3-5): 135-41.

Griffith, O. W. and A. Meister (1985). "Origin and turnover of mitochondrial
glutathione." Proc Natl Acad Sci U S A 82(14): 4668-72.

Haddad, I. Y., J. P. Crow, et al. (1994). "Concurrent generation of nitric oxide and
superoxide damages surfactant protein A." Am. J. Physiol. 267: L242-L249.

Halliwell, B. and J. M. C. Gutteridge (1989). Free radicals in biology and medicine.
Oxford New York, Clarendon Press ;Oxford University Press.

Hammond, C. L., T. K. Lee, et al. (2001). "Novel roles for glutathione in gene
expression, cell death, and membrane transport of organic solutes." J Hepatol
34(6): 946-54.

Han, D., F. Antunes, et al. (2003). "Voltage-dependent anion channels control the
release of the superoxide anion from mitochondria to cytosol." J Biol Chem
278(8): 5557-63.

Han, D., F. Antunes, et al. (2003). "Voltage-dependent anion channels control the
release of the superoxide anion from mitochondria to cytosol." J. Biol. Chem.
278: 5557-5563.
125

Han, D., F. Antunes, et al. (2002). "Mitochondrial superoxide anion production and
release into intermembrane space." Meth. Enzymol. 349: 271-280.

Han, D., R. Canali, et al. (2005). "Sites and mechanisms of aconitase inactivation by
peroxynitrite: modulation by citrate and glutathione." Biochemistry 44(36):
11986-96.

Han, D., R. Canali, et al. (2003). "Effect of glutathione depletion on sites and topology
of superoxide and hydrogen peroxide production in mitochondria." Mol
Pharmacol 64(5): 1136-44.

Han, D., N. Hanawa, et al. (2006). "Mechanisms of liver injury. III. Role of
glutathione redox status in liver injury." Am J Physiol Gastrointest Liver
Physiol 291(1): G1-7.

Han, D., K. Matsumaru, et al. (2004). "Usnic acid-induced necrosis of cultured mouse
hepatocytes: inhibition of mitochondrial function and oxidative stress."
Biochem Pharmacol 67(3): 439-51.

Han, D., E. Williams, et al. (2001). "Mitochondrial respiratory chain-dependent
generation of superoxide anion and its release into the intermembrane space."
Biochem J 353(Pt 2): 411-6.

Hansen, J. M., Y. M. Go, et al. (2006). "Nuclear and mitochondrial compartmentation
of oxidative stress and redox signaling." Annu Rev Pharmacol Toxicol 46:
215-34.

Harman, L. S., C. Mottley, et al. (1984). "Free radical metabolites of L-cysteine
oxidation." J. Biol. Chem. 259: 5606-5611.

Harvey, P. R., R. G. Ilson, et al. (1989). "The simultaneous determination of oxidized
and reduced glutathiones in liver tissue by ion pairing reverse phase high
performance liquid chromatography with a coulometric electrochemical
detector." Clin Chim Acta 180(3): 203-12.

Hausladen, A. and I. Fridovich (1996). "Measuring nitric oxide and superoxide: rate
constant for aconitase reactivity." Methods Enzymol. 269: 37-41.

Herlinger, E., R. F. Jameson, et al. (1995). "Spontaneous Autoxidation of Dopamine."
Journal of the Chemical Society-Perkin Transactions 2(2): 259-263.

Hodaie, M., J. S. Neimat, et al. (2007). "The dopaminergic nigrostriatal system and
Parkinson's disease: molecular events in development, disease, and cell death,
and new therapeutic strategies." Neurosurgery 60(1): 17-28; discussion 28-30.
126

Hurd, T. R., N. J. Costa, et al. (2005). "Glutathionylation of mitochondrial proteins."
Antioxid Redox Signal 7(7-8): 999-1010.

Iravani, M. M., K. Kashefi, et al. (2002). "Involvement of inducible nitric oxide
synthase in inflammation-induced dopaminergic neurodegeneration."
Neuroscience 110(1): 49-58.

Jain, A., J. Martensson, et al. (1991). "Glutathione deficiency leads to mitochondrial
damage in brain." Proc Natl Acad Sci U S A 88(5): 1913-7.

Jenner, P. (1998). "Oxidative mechanisms in nigral cell death in Parkinson's disease."
Mov Disord 13 Suppl 1: 24-34.

Jo, S. H., M. K. Son, et al. (2001). "Control of mitochondrial redox balance and
cellular defense against oxidative damage by mitochondrial NADP+-
dependent isocitrate dehydrogenase." J Biol Chem 276(19): 16168-76.

Johansson, C., C. H. Lillig, et al. (2004). "Human mitochondrial glutaredoxin reduces
S-glutathionylated proteins with high affinity accepting electrons from either
glutathione or thioredoxin reductase." J Biol Chem 279(9): 7537-43.

Jones, D. P. (2006). "Redefining oxidative stress." Antioxid Redox Signal 8(9-10):
1865-79.

Jung, C. H. and J. A. Thomas (1996). "S-glutathiolated hepatocyte proteins and insulin
disulfides as substrates for reduction by glutaredoxin, thioredoxin, protein
disulfide isomerase, and glutathione." Arch Biochem Biophys 335(1): 61-72.

Kennedy, M. C., W. E. Antholine, et al. (1997). "An EPR investigation of the products
of the reaction of cytosolic and mitochondrial aconitase with nitric oxide." J.
Biol. Chem. 272: 20340-7.

Kennedy, M. C., G. Spoto, et al. (1988). "The active site sulfhydryl of aconitase is not
required for catalytic activity." J. Biol. Chem. 263: 8190-8193.

Klatt, P. and S. Lamas (2000). "Regulation of protein function by S-glutathiolation in
response to oxidative and nitrosative stress." Eur J Biochem 267(16): 4928-44.

Koppenol, W., J. Moreno, et al. (1992). "Peroxynitrite, a cloaked oxidant formed by
nitric oxide and superoxide." Chem. Res. Toxicol. 5: 834-842.

Kwong, L. K. and R. S. Sohal (1998). "Substrate and site specificity of hydrogen
peroxide generation in mouse mitochondria." Arch Biochem Biophys 350(1):
118-26.
127

Lauble, H., M. C. Kennedy, et al. (1992). "Crystal structures of aconitase with
isocitrate and nitroisocitrate bound." Biochemistry 31: 2735-2748.

Lauble, H. and C. D. Stout (1995). "Steric and conformational features of the
aconitase mechanism." Proteins 22: 1-11.

Liberatore, G. T., V. Jackson-Lewis, et al. (1999). "Inducible nitric oxide synthase
stimulates dopaminergic neurodegeneration in the MPTP model of Parkinson
disease." Nat Med 5(12): 1403-9.

Lluis, J. M., A. Colell, et al. (2003). "Acetaldehyde impairs mitochondrial glutathione
transport in HepG2 cells through endoplasmic reticulum stress."
Gastroenterology 124(3): 708-24.

Lowry, O. H., J. Carter, et al. (1971). "The effect of carbon and nitrogen sources on
the level of metabolic intermediates in Escherichia coli." J. Biol. Chem. 246:
6511-21.

Lundberg, M., C. Johansson, et al. (2001). "Cloning and expression of a novel human
glutaredoxin (Grx2) with mitochondrial and nuclear isoforms." J Biol Chem
276(28): 26269-75.

Maples, K. R., S. J. Jordan, et al. (1988). "In vivo rat hemoglobin thiyl free radical
formation following phenylhydrazine administration." Mol. Pharmacol. 33:
344-350.

Martensson, J., J. C. Lai, et al. (1990). "High-affinity transport of glutathione is part of
a multicomponent system essential for mitochondrial function." Proc Natl
Acad Sci U S A 87(18): 7185-9.

Martinez-Ruiz, A. and S. Lamas (2004). "S-nitrosylation: a potential new paradigm in
signal transduction." Cardiovascular Research 62(1): 43-52.

Martinez-Ruiz, A. and S. Lamas (2004). "S-nitrosylation: a potential new paradigm in
signal transduction." Cardiovasc Res 62(1): 43-52.

Mcgeer, P. L., S. Itagaki, et al. (1988). "Reactive Microglia Are Positive for Hla-Dr in
the Substantia Nigra of Parkinsons and Alzheimers-Disease Brains."
Neurology 38(8): 1285-1291.

Nulton-Persson, A. C. and L. I. Szweda (2001). "Modulation of mitochondrial
function by hydrogen peroxide." J. Biol. Chem. 276: 23357-23361.

128

Olanow, C. W. and W. G. Tatton (1999). "Etiology and pathogenesis of Parkinson's
disease." Annu Rev Neurosci 22: 123-44.

Packer, M. A. and M. P. Murphy (1994). "Peroxynitrite causes calcium efflux from
mitochondria which is prevented by cyclosporin A." FEBS Lett. 345: 237-240.

Perry, T. L., D. V. Godin, et al. (1982). "Parkinson's disease: a disorder due to nigral
glutathione deficiency?" Neurosci Lett 33(3): 305-10.

Perry, T. L. and V. W. Yong (1986). "Idiopathic Parkinson's disease, progressive
supranuclear palsy and glutathione metabolism in the substantia nigra of
patients." Neurosci Lett 67(3): 269-74.

Poderoso, J. J., M. C. Carreras, et al. (1996). "Nitric oxide inhibits electron transfer
and increases superoxide radical production in rat heart mitochondria and
submitochondrial particles." Arch. Biochem. Biophys. 328(1): 85-92.

Poderoso, J. J., C. Lisdero, et al. (1999). "The regulation of mitochondrial oxygen
uptake by redox reactions involving nitric oxide and ubiquinol." J Biol Chem
274(53): 37709-16.

Radi, R., J. S. Beckman, et al. (1991). "Peroxynitrite oxidation of sulfhydryls. The
cytotoxic potential of superoxide and nitric oxide." J. Biol. Chem. 266: 4244-
4250.

Radi, R., J. S. Beckman, et al. (1991). "Peroxynitrite oxidation of sulfhydryls." J. Biol.
Chem. 266(7): 4244-4250.

Radi, R., A. Cassina, et al. (2002). "Peroxynitrite reactions and formation in
mitochondria." Free Radic. Biol. Med. 33: 1451-1464.

Rettori, D., Y. Tang, et al. (2002). "Pathways of dopamine oxidation mediated by
nitric oxide." Free Radic Biol Med 33(5): 685-90.

Richards, E. M., R. E. Rosenthal, et al. (2006). "Postischemic hyperoxia reduces
hippocampal pyruvate dehydrogenase activity." Free Radic Biol Med 40(11):
1960-70.

Riobó, N. A., E. Clementi, et al. (2001). "Nitric oxide inhibits mitochondrial NADH-
ubiquinone reductase activity through the formation of peroxynitrite."
Biochem. J. 359: 139-145.

129

Riobo, N. A., M. Melani, et al. (2002). "The modulation of mitochondrial nitric-oxide
synthase activity in rat brain development." Journal of Biological Chemistry
277(45): 42447-42455.

Robbins, A. H. and C. D. Stout (1989). "Structure of activated aconitase: Formation of
the [4Fe-4S] cluster in the crystal." Proc. Natl. Acad. Sci. USA 86: 3639-3643.

Romero, F. J. and H. Sies (1984). "Subcellular glutathione contents in isolated
hepatocytes treated with L-buthionine sulfoximine." Biochem Biophys Res
Commun 123(3): 1116-21.

Sabadie-Pialoux, N. and D. Gautheron (1971). "Free--SH variations during ATP
synthesis by oxidative phosphorylation in heart muscle mitochondria."
Biochim Biophys Acta 234(1): 9-15.

Saggu, H., J. Cooksey, et al. (1989). "A selective increase in particulate superoxide
dismutase activity in parkinsonian substantia nigra." J Neurochem 53(3): 692-
7.

Scarlett, J. L., M. A. Packer, et al. (1996). "Alterations to glutathione and nicotinamide
nucleotides during the mitochondrial permeability transition induced by
peroxynitrite." Biochem Pharmacol 52(7): 1047-55.

Schafer, F. Q. and G. R. Buettner (2001). "Redox environment of the cell as viewed
through the redox state of the glutathione disulfide/glutathione couple." Free
Radic Biol Med 30(11): 1191-212.

Schapira, A. H. (1998). "Human complex I defects in neurodegenerative diseases."
Biochim Biophys Acta 1364(2): 261-70.

Schopfer, F., N. A. Riobo, et al. (2000). "Oxidation of ubiquinol by peroxynitrite:
implications for protection of mitochondria against nitrosative damage."
Biochem. J. 349: 35-42.

Schroeter, H., C. S. Boyd, et al. (2003). "c-Jun N-terminal kinase (JNK)-mediated
modulation of brain mitochondria function: new target proteins for JNK
signaling in mitochondrion-dependent apoptosis." Biochem J 372(Pt 2): 359-
69.

Schubert, D. and F. G. Klier (1977). "Storage and release of acetylcholine by a clonal
cell line." Proc Natl Acad Sci U S A 74(11): 5184-8.

130

Sciamanna, M. A. and C. P. Lee (1993). "Ischemia/reperfusion-induced injury of
forebrain mitochondria and protection by ascorbate." Arch Biochem Biophys
305(2): 215-24.

Shan, X., D. P. Jones, et al. (1993). "Selective depletion of mitochondrial glutathione
concentrations by (R,S)-3-hydroxy-4-pentenoate potentiates oxidative cell
death." Chem Res Toxicol 6(1): 75-81.

Sian, J., D. T. Dexter, et al. (1994). "Alterations in glutathione levels in Parkinson's
disease and other neurodegenerative disorders affecting basal ganglia." Ann
Neurol 36(3): 348-55.

Spencer, J. P., P. Jenner, et al. (1998). "Conjugates of catecholamines with cysteine
and GSH in Parkinson's disease: possible mechanisms of formation involving
reactive oxygen species." J Neurochem 71(5): 2112-22.

Srisook, K., C. Kim, et al. (2005). "Cytotoxic and cytoprotective actions of O2- and
NO (ONOO-) are determined both by cellular GSH level and HO activity in
macrophages." Methods Enzymol 396: 414-24.

Stokes, A. H., T. G. Hastings, et al. (1999). "Cytotoxic and genotoxic potential of
dopamine." J Neurosci Res 55(6): 659-65.

Tatton, W. G. and C. W. Olanow (1999). "Apoptosis in neurodegenerative diseases:
the role of mitochondria." Biochim Biophys Acta 1410(2): 195-213.

Taylor, E. R., F. Hurrell, et al. (2003). "Reversible glutathionylation of complex I
increases mitochondrial superoxide formation." J Biol Chem 278(22): 19603-
10.

Thomas, J. A., B. Poland, et al. (1995). "Protein sulfhydryls and their role in the
antioxidant function of protein S-thiolation." Arch Biochem Biophys 319(1):
1-9.

Thomas, J. A., B. Poland, et al. (1995). "Protein sulfhydryls and their role in the
antioxidant function of protein S-thiolation." Arch. Biochem. Biophys. 319(1):
1-9.

Toffa, S., G. M. Kunikowska, et al. (1997). "Glutathione depletion in rat brain does
not cause nigrostriatal pathway degeneration." J Neural Transm 104(1): 67-75.

Torreilles, F., S. Salman-Tabcheh, et al. (1999). "Neurodegenerative disorders: the
role of peroxynitrite." Brain Res Brain Res Rev 30(2): 153-63.

131

Torres, J., V. Darley-Usmar, et al. (1995). "Inhibition of cytochrome c oxidase in
turnover by nitric oxide: mechanism and implications for control of
respiration." Biochem. J. 312: 169-173.

Turrens, J. F. and A. Boveris (1980). "Generation of superoxide anion by the NADH
dehydrogenase of bovine heart mitochondria." Biochem J 191(2): 421-7.

Uhlig, S. and A. Wendel (1992). "The physiological consequences of glutathione
variations." Life Sci 51(14): 1083-94.

Vizi, E. S. (2000). "Role of high-affinity receptors and membrane transporters in
nonsynaptic communication and drug action in the central nervous system."
Pharmacol Rev 52(1): 63-89.

Wallin, C., M. Puka-Sundvall, et al. (2000). "Alterations in glutathione and amino acid
concentrations after hypoxia-ischemia in the immature rat brain." Brain Res
Dev Brain Res 125(1-2): 51-60.

Wang, J., E. S. Boja, et al. (2001). "Reversible glutathionylation regulates actin
polymerization in A431 cells." J. Biol. Chem. 276: 47763-6.

West, M. B., B. G. Hill, et al. (2006). "Protein glutathiolation by nitric oxide: an
intracellular mechanism regulating redox protein modification." Faseb J
20(10): 1715-7.

Whitton, P. S. (2007). "Inflammation as a causative factor in the etiology of
Parkinson's disease." Br J Pharmacol.

Wullner, U., J. Seyfried, et al. (1999). "Glutathione depletion and neuronal cell death:
the role of reactive oxygen intermediates and mitochondrial function." Brain
Res 826(1): 53-62.

Zeevalk, G. D., L. Manzino, et al. (2007). "Characterization of intracellular elevation
of glutathione (GSH) with glutathione monoethyl ester and GSH in brain and
neuronal cultures: relevance to Parkinson's disease." Exp Neurol 203(2): 512-
20.

Zhang, L., V. L. Dawson, et al. (2006). "Role of nitric oxide in Parkinson's disease."
Pharmacol Ther 109(1-2): 33-41.

Abstract (if available)

Abstract The increased generation of reactive oxygen and nitrogen species and glutathione depletion creates profound changes in the redox status of the cell and has been implicated in the progression of neurodegenerative diseases such as Parkinson's disease. Redox regulation has become exceedingly important in understanding the cellular adaptation to oxidative stress. However, it remains unclear how changes in the cellular and mitochondrial redox environment affect redox sensitive protein function during exposure to NO and dopamine and the consequences of such changes with respect to mitochondrial function and redox signaling (achieved through post-translational modifications).

Linked assets

University of Southern California Dissertations and Theses

Conceptually similar

PDF

NO mediated neurotoxicity: redox changes and energy failure in a neuroinflammatory model

PDF

The mitochondrial energy – redox axis in aging and caloric restriction: role of nicotinamide nucleotide transhydrogenase

PDF

Modulation of the redox status of isolated mitochondria by energy-linked substrates: quantification by high performance liquid chromatography; and "Splicing up" drug discovery, cell-based express...

PDF

Post-translational modification crosstalk regulates KAP1 co-repressor functions in response to DNA damages

PDF

PI3K/AKT signaling and the regulation of the mitochondrial energy-redox axis

PDF

LDL protein nitration: implication for protein unfolding and mitochondrial function by p-JNK-2

PDF

Role of neuronal nitric oxide synthase in aging and neurodegeneration

PDF

C-jun N-terminal Kinase (JNK) mediated inhibition of Pyruvate Dehydrogenase (PDH) activity and its effect on mitochondrial metabolism during brain aging

PDF

Metabolic shift in lung alveolar cell mitochondria after exposure to environmental toxicants

PDF

Mitochondrial dynamics regulate Leydig cell health and integrity

PDF

The differential effects of genistein on cellular effects in T47D tumorigenic and MCF10A nontumorigenic breast epithelial cells: role of metabolism

PDF

Uncovering the protective role of protein glycosylation in Parkinson's disease utilizing protein semi-synthesis

PDF

Novel synthesis of β-glycosides for SPPS of GLCNAC glycoproteins and study of their site-specific biochemical and biophysical consequences

PDF

Energy metabolism and inflammation in brain aging: significance of age-dependent astrocyte metabolic-redox profile

PDF

Synthesis, characterization and application of chemical tools for investigating the role of O-GlcNAc modification in the development and survival of mammalian cells

PDF

Developing and exploiting small molecules to study O-GlcNAc modification

PDF

The structure and function of membrane curving proteins on different membrane shapes and their regulation by post-translational modifications

PDF

Impacts of post-translational modifications on interactions between G9a and its N-terminus binding partners

PDF

Using chemistry to reveal the consequences of post translational modifications in cancer

PDF

Optimization of chemical reporters of O-GlcNAc for improved specificity and metabolic mapping

Asset Metadata

Creator Garcia, Jerome Vincent (author)

Core Title Protein post-translational modifications in the cell's redox and energy states

School School of Pharmacy

Degree Doctor of Philosophy

Degree Program Molecular Pharmacology

Publication Date 05/03/2007

Defense Date 03/27/2007

Publisher University of Southern California (original), University of Southern California. Libraries (digital)

Tag mitochondria,OAI-PMH Harvest,protein post-translational modifications,redox

Language English

Advisor Cadenas, Enrique (committee chair), Alkana, Ronald (committee member), Hsiai, Tzung K. (committee member), Sevanian, Alex (committee member)

Creator Email jvg@usc.edu

Permanent Link (DOI) https://doi.org/10.25549/usctheses-m480

Unique identifier UC1296093

Identifier etd-Garcia-20070503 (filename),usctheses-m40 (legacy collection record id),usctheses-c127-492263 (legacy record id),usctheses-m480 (legacy record id)

Legacy Identifier etd-Garcia-20070503.pdf

Dmrecord 492263

Document Type Dissertation

Rights Garcia, Jerome Vincent

Type texts

Source University of Southern California (contributing entity), University of Southern California Dissertations and Theses (collection)

Repository Name Libraries, University of Southern California

Repository Location Los Angeles, California

Repository Email cisadmin@lib.usc.edu