Mitochondrial Electron Transport: Oxidative Phosphorylation, Mitochondrial Oxidant Production, and Methods of Measurement

Abstract

The mitochondrial electron transport chain utilizes a series of electron transfer reactions to generate cellular ATP through oxidative phosphorylation. A consequence of electron transfer is the generation of reactive oxygen species (ROS), which contributes to both homeostatic signaling as well as oxidative stress during pathology. In this graphical review we provide an overview of oxidative phosphorylation and its inter-relationship with ROS production by the electron transport chain. We also outline traditional and novel translational methodology for assessing mitochondrial energetics in health and disease.

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Mitochondrial Electron Transport: Oxidative Phosphorylation, Mitochondrial Oxidant
Production, and Methods of Measurement
Deirdre Nolfi-Donegan, Andrea Braganza, Sruti Shiva
PII: S2213-2317(20)30879-X
DOI: https://doi.org/10.1016/j.redox.2020.101674
Reference: REDOX 101674
To appear in: Redox Biology
Received Date: 1 April 2020
Revised Date: 24 July 2020
Accepted Date: 31 July 2020
Please cite this article as: D. Nolfi-Donegan, A. Braganza, S. Shiva, Mitochondrial Electron Transport:
Oxidative Phosphorylation, Mitochondrial Oxidant Production, and Methods of Measurement, Redox
Biology, https://doi.org/10.1016/j.redox.2020.101674.
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© 2020 The Author(s). Published by Elsevier B.V.

Mitochondrial Electron Transport: Oxidative Phosphorylation,
Mitochondrial Oxidant Production, and Methods of Measurement
Deirdre Nolfi-Donegan
1,2#
, Andrea Braganza
1#
, and Sruti Shiva
1,3
*
1
Heart, Lung, Blood Vascular Medicine Institute, University of Pittsburgh, Pittsburgh, PA 15261
2
Department of Hematology, Children’s Hospital of Pittsburgh, Pittsburgh, PA 15224
3
Department of Pharmacology & Chemical Biology, University of Pittsburgh, Pittsburgh, PA
15261
#
These authors contributed equally
*Corresponding Author:
Sruti Shiva, PhD
200 Lothrop Street BST E1240
University of Pittsburgh
Pittsburgh, PA 15213
Phone: (412)383-5854
Fax: (412) 648-5980
Email: sss43@pitt.edu
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Abstract:
The mitochondrial electron transport chain utilizes a series of electron transfer reactions to generate
cellular ATP through oxidative phosphorylation. A consequence of electron transfer is the generation of
reactive oxygen species (ROS), which contributes to both homeostatic signaling as well as oxidative
stress during pathology. In this graphical review we provide an overview of oxidative phosphorylation
and its inter-relationship with ROS production by the electron transport chain. We also outline traditional
and novel translational methodology for assessing mitochondrial energetics in health and disease.
Journal Pre-proof

Introduction:
In 1957, Peter Siekevitz branded the mitochondrion the “powerhouse” of the cell [1]. Less than a
decade later came the first reports that the organelle generated reactive oxygen species (ROS) as a
byproduct of cellular respiration [2]. Since then, it has become apparent that mitochondria are highly
dynamic organelles that contribute to cellular homeostasis not only through maintaining adenosine
triphosphate (ATP) levels, but also through the generation of low levels of ROS for cell signaling, and
that dysfunction in either of these processes can propagate pathology. Notably, the importance of each
of these functions varies by cell type. For example, cardiomyocytes rely on mitochondria to supply
>95% of the energy required for their function [3]. In contrast, endothelial cells rely more heavily on
glycolysis than mitochondria for ATP, but mitochondrial ROS production is essential for endothelial
homeostatic signaling [4].
Despite this variation between cell types, mitochondrial ATP generation and ROS production are
intimately linked through function of the electron transport chain (ETC), and thus efficient measurement
of ETC function can provide insight into mechanisms of physiology and disease pathogenesis. In this
review, we will provide an overview of the function of the ETC, focusing on oxidative phosphorylation
and its relationship to ROS production. Further, we will compare current methodologies to measure
bioenergetic function and mitochondrial ROS production.
Overview of oxidative phosphorylation
Cellular metabolism comprises the utilization of carbohydrates, fats, and proteins, to synthesize energy.
The processes for the catabolism of glucose (via glycolysis and subsequent pyruvate oxidation), fatty
acids (via fatty acid β-oxidation), and amino acids (via the oxidative deamination and transamination)
are reviewed in detail elsewhere [5]. However, the molecules derived from these processes are used in
the tricarboxylic acid (TCA) cycle to generate substrates that enter the ETC for oxidative
phosphorylation. Here we summarize the reactions that occur in the ETC to produce energy, but for a
more detailed review, refer to Zhao et al. [6].
The ETC is embedded within the extensive inner membrane of the mitochondrion, in close proximity to
the mitochondrial matrix in which the TCA cycle is localized (Figure 1). NADH and FADH
2
generated by
the TCA cycle donate electrons to the ETC at either Complex I (NADH:ubiquinone oxidoreductase) or
Complex II (succinate dehydrogenase), respectively (Figure 1). The electrons from NADH are passed
to ubiquinone (CoQ) through a chain of co-factors including a flavin mononucleotide (FMN) followed by
seven low to high potential iron-sulfur (FeS) clusters in Complex I to enter the Q cycle, where CoQ is
reduced to ubiquinol (QH
2
). This electron transfer induces the pumping of protons by complex I from
the matrix into the intermembrane space. Though the mechanism linking electron transfer to proton
pumping remains unclear, one hypothesis speculates an indirect pumping of two protons via a
conformation-coupled manner and the direct pumping of the other two protons via the ubiquinone redox
reaction, while another hypothesis suggests that changes in the conformation and density of water in
Complex I dictates the proton translocation [6]. Regardless of the exact mechanism, the transfer of two
electrons from NADH results in the pumping of four protons.
Electrons also enter the ETC through Complex II, which is a component of both the TCA cycle and the
ETC. Electrons donated from FADH
2
are transferred sequentially to CoQ via the FeS cluster of
Complex II, in a similar manner as at Complex I. Unlike Complex I, electron transport at Complex II is
not accompanied by proton translocation from the matrix to the intermembrane space [6].
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Once in the Q-cycle, electrons are transferred to Complex III (coenzyme Q:cytochrome c reductase)
and then to cytochrome c. First, QH
2
binds to the cytoplasmic side of Complex III and releases two
protons into the intermembrane space. One electron is transferred from QH
2
to the catalytically active
and high-potential iron-sulfur cluster in the Rieske center of Complex III, while the second electron is
transferred to cytochrome b within the complex. This electron is then transferred from cytochrome b to
a second molecule of Q that is bound to the matrix side of the complex to generate ubisemiquinone (Q·
-
). Concomitantly, the electron at the 2Fe-2S cluster is transferred to cytochrome c
1
, from where it is
transferred to, and reduces the mobile carrier, cytochrome c. At this point, a second QH
2
molecule
binds to the membrane side of the complex and undergoes the same oxidation process to reduce Q·
-
back to QH
2
, complete the Q-cycle, and pump two more protons into the intermembrane space [6].
Once the mobile electron carrier cytochrome c is reduced, it ferries single electrons from Complex III to
Complex IV (cytochrome c oxidase), where molecular oxygen binds and is reduced to water. Complex
IV consists of three core subunits, namely, the central subunit I that contains the redox-active metal
centers heme a (Fe
a
) and a binuclear center composed of heme a
3
(Fe
a3
) and Cu
B
, and flanking subunit
II, that contains the other redox-active metal center, namely, Cu
A
and subunit III that is involved in
proton pumping. Reduced cytochrome c can simultaneously interact with subunit II of Complex IV, and
transfer electrons to the Cu
A
site of subunit II to then pass these electrons from heme a to the binuclear
center of subunit I, where O
2
binds and is reduced to H
2
O. Therefore, at Complex IV, a total of eight
protons are pumped from the matrix, of which, four are used to form two water molecules, and the other
four are transferred into the intermembrane space. This process of oxygen consumption is known as
mitochondrial respiration [6].
In response to electron transport, a total of ten protons (H
+
) (two from Complex III, and four from each
Complex I and Complex IV) are pumped from the matrix into the intermembrane space, where they
accumulate to generate an electrochemical proton gradient known as the mitochondrial membrane
potential (ΔΨ). ΔΨ combined with proton concentration (pH) generates a protonmotive force (Δp) which
is an essential component in the process of energy storage during OXPHOS since it couples electron
transport (complexes I-IV) (and oxygen consumption) to the activity of Complex V (ATP synthase),
where protons re-enter the matrix to dissipate the proton gradient. Complex V is a multi-subunit
complex comprised of two distinct domains, extra-membranous (termed F
1
) and transmembrane
(termed F
O
), and functions under a rotational motor mechanism to allow for ATP production. Proton
movement through F
O
from the intermembrane space is coupled to the rotation that results in the
addition of a phosphate to adenosine diphosphate (ADP) to synthesize adenosine triphosphate (ATP)
at sites in F
1
(Figure 1) [6].
Generation of Superoxide from the ETC
It is now well established that the mitochondrion is a significant source of cellular ROS (Figure 2).
While multiple sites of ROS production have been identified in the organelle and reviewed in detail
elsewhere [7], this section will focus on generation of ROS by the complexes of the ETC and its
interplay with cellular respiration and energetics. The predominant route of ROS production by the ETC
is the premature leak of electrons from complexes I, II, and III to mediate the one electron reduction of
oxygen to superoxide (O
2
•
−
), which can then be dismutated to hydrogen peroxide (H
2
O
2
) [6]. The rate of
O
2
•
−
production from the ETC
is dependent on the concentration of the one-electron donor at a
particular site and the rate at which this redox active donor reacts with oxygen O
2
[8]. Importantly,
factors such as local O
2
tension, Δp, electron flux, and ATPase activity can change these factors to
dynamically modulate O
2
•
−
from the ETC, and these factors will be considered along with a brief
explanation of the mechanisms of ETC O
2
•
−
generation at each complex.
Two sites of O
2
•
−
generation have been identified at Complex I - 1) the FMN cofactor which accepts
electrons from NADH and 2) the Q binding site at which two electrons are transferred the terminal Fe-S
to Q. With forward electron transport, electrons from NADH fully reduce the FMN center, which reacts
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with oxygen to generate O
2
•
−
[8]. Thus, O
2
•
−
production at this center is regulated by the ratio of NADH
to NAD+ and in physiological conditions is relatively low. However, conditions leading to the
accumulation of NADH, such as low ATP demand resulting in decreased respiration or damage to the
ETC, lead to greater reduction of the FMN and increased O
2
•
−
production at this site. Notably, the
Complex I inhibitor rotenone, which binds to the Q binding site, stimulates O
2
•
−
production by
potentiating electron accumulation and reduction of the FMN site. A second mechanism by which
Complex I generates O
2
•
−
is dependent on reverse electron transport (RET) [9]. RET occurs when the
Q pool is highly reduced and Δp is high. In these conditions, the energy is high enough to drive electron
transport against the redox potential gradient of the ETC and electrons are driven back from QH
2
into
Complex I. Superoxide generated by RET is thought to occur at the Q binding site of Complex I. This is
supported by the fact that RET-dependent O
2
•
−
production is abolished by rotenone. Further, this
mechanism of ROS production is highly sensitive to minimal changes in pH. Given that proton pumping
is linked with the Q binding site, it is likely that Q•
−
is formed during proton pumping and RET, leading to
the O
2
•
−
generation observed. While RET-induced ROS production was once thought to be a strictly in
vitro phenomenon, it is now known to contribute to the pathogenesis of bacterial sepsis, as well as
physiological hypoxia-sensing in the carotid body [10].
It is important to note that in physiological conditions, when respiration and ATP production are high
(and Δp is low), Complex I ROS production is low. While early studies suggested that physiological
ROS production by Complex I was high, it is now recognized that much of the ROS measured in those
studies may have originated from flavin-dependent dehydrogenases that utilize the NADH/NAD
+
pool
and operate at a similar redox potential as Complex I. These enzymes, which include α-ketoglutarate
dehydrogenase, pyruvate dehydrogenase, and branched chain 2-oxoacid dehydrogenase are now
known to be significant generators of ROS and generate O
2
•
−
in the matrix of the mitochondrion
(similarly to both sites of the Complex I) (Figure 2) [11]. However, under conditions of high
NADH/NAD
+
or high QH
2
/Q and elevated Δp, O
2
•
−
production is significantly enhanced by FMN
reduction and RET respectively at complex I, significantly out-producing these dehydrogenase
complexes.
Complex III, specifically the site of QH
2
oxidation, is also a generator of O
2
•
−
within the ETC. As
described above, the two electrons carried by QH
2
when it binds complex III are transferred from QH
2
to
the Rieske Fe-S center and then to cytochrome c in sequence. The transfer of one electron with the
other remaining on Q favors the formation of the unstable Q•
−
species, which can react with oxygen to
produce O
2
•
−
. The rate of ROS production at this site is physiologically low, and favored when the site
is only partially reduced, which occurs when substrate is at submaximal levels. Alternatively, inhibition
of the complex downstream of Q•
−
also potentiates ROS production [11]. For example, treatment with
the Complex III inhibitor Antimycin A binds the Qi site of the complex, stabilizing the semiquinone
radical in the Q
o
site. Addition of Q
o
inhibitors such as myxathiazol completely inhibit ROS production by
complex III [12]. Similarly, to dehydrogenases that contribute to the NADH/NAD
+
pool, several enzymes
responsible for substrate catabolism and operate at the same redox potential as Complex III are
generators of O
2
•
−.
These enzymes include mitochondrial glycerol-3-phosphate dehydrogenase
(mGPDH), the electron transferring flavoprotein/ ETF:ubiquinone oxidoreductase system, and
dihydroorotate dehydrogenase (DHODH), all of which donate electrons to the CoQ pool. Of note, ROS
generated by these enzymes as well as complex III are released on both the matrix and intermembrane
side of the mitochondrion (Figure 2) [11].
While Complex II is not generally considered a major source of ROS in comparison to Complexes I and
III, several reports describe O
2
•
−
production by the complex. While there is some debate surrounding
the exact site of ROS production in the complex, Brand and colleagues have demonstrated that the
flavin site, at which FAD binds the active site of the enzyme, is responsible for the production. These
studies demonstrate that in isolated mitochondria ROS generation by Complex II is highly regulated by
succinate concentration, with a bell-shaped response in which ROS production is optimized when
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succinate concentration is not too high or too low. Moreover, the Q pool must be highly reduced such
as what is observed with Complex III inhibition [11]. While the exact mechanism by which Complex II
generates ROS is being elucidated, its contribution to overall mitochondrial ROS production in
physiological conditions remains unknown.
Early studies estimated that 1-2% of electrons entering the ETC contributed to O
2
•
−
production. It is
now appreciated that the rate of mitochondrial ROS generation is likely lower than this value in vivo [13]
and fluctuates depending on ETC function. As outlined above, each site of mitochondrial ROS
production is differentially sensitive to factors such as substrate supply, rate of ATP production and Δp
depending on how these conditions affect the reduction of particular site of ROS production.
Importantly, discussion of the impact of these factors assumes high efficiency of oxidative
phosphorylation in which respiration is “coupled” to ATP generation by Δp and the membrane potential
(Δψ
m
) is maintained within a homeostatic range (140-160mV) [14] (Figure 2A). Physiologically,
oxidative phosphorylation is not completely coupled, and low levels of proton leak dissipate Δp and
decrease Δψ, which can attenuate ROS production. The level of basal proton leak is determined by
inner membrane structure and composition as well as proteins such as the adenine nucleotide
translocase (ANT), which regulates the Δψ
m
– driven exchange of ATP for ADP across the inner
membrane, but also is responsible for over half of basal proton leak [6]. In addition to ANT, the
mitochondrial ATP-sensitive potassium channel (mitoK
ATP
) also mediates uncoupling in response to
high concentrations of superoxide and H
2
O
2
.
Free fatty acids can also act as uncouplers, though this
function appears to be distinct from their ability to act as substrate to fuel respiration [15]. Beyond these
proteins and substrates that are constitutive to the mitochondrion, expression of specific uncoupling
proteins (UCP) within the inner mitochondrial membrane facilitate the leak of protons from the
intermembrane space back to the matrix without involvement of the ATP synthase. Expression of these
UCP proteins is induced by many stimuli, including inner membrane hyperpolarization and high levels
of ROS. Expression of UCP1 was first identified in brown adipose tissue, in which uncoupling of
oxidative phosphorylation generates heat [16]. However, it is now known that at least five different UCP
isoforms exist and expression of these proteins are implicated in a variety of physiological signaling
pathways ranging from T-cell maturation [17] and neuroprotection [18], to mediating the protective
effects of caloric restriction [19]. For an in-depth review of UCP function see [15].
A final consideration in the factors regulating O
2
•
−
production from the ETC is the availability of oxygen
as an electron acceptor. While decreased availability of oxygen should attenuate O
2
•
−
production, a
number of studies in cell systems demonstrate that mitochondrial ROS production is paradoxically
increased in hypoxic conditions [12, 20]. Several hypotheses have been proposed to explain this effect
including the notion that oxygen limitation at Complex IV causes accumulation of electrons and
increased reduction of Complexes I-III. However, given the low K
m
of Complex IV for O
2
(<1µM),
limitation at Complex IV should also limit oxygen available for reduction to O
2
•
−
[21]. More recent
studies have suggested that hypoxia induces a conformational change in Complex III such that Q•
−
is
stabilized or that there is more access of O
2
to the Q•
−
[12, 20]. Notably, Brookes and colleagues
showed in controlled experiments utilizing isolated mitochondria and fixed substrate concentration that
unlike in cells, ROS
production by the ETC decreased as O
2
concentrations were lowered below ~1µM
O
2
[21]. These data demonstrate a discrepancy between the effect of hypoxia in isolated mitochondria
and cell systems, and suggest that cellular factors outside the mitochondrion may regulate ETC ROS
production during hypoxia in cells. These factors may include nitric oxide (NO) production, which
regulates the ETC at Complex V [22], changes in substrate entry into the ETC or utilization, or cellular
kinase activity that potentially regulates ETC activity through phosphorylation [23].
Mitochondrial Antioxidant Systems and ROS-mediated signaling
While the mitochondrion is a site for ROS production, antioxidant systems contribute to the regulation of
the concentration and redox species in the organelle. Though the proximal ROS generated by the ETC
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is O
2
•
−
, this species is rapidly dismutated to H
2
O
2
by manganese superoxide dismutase (MnSOD)
localized in the mitochondrial matrix and low concentrations of copper/zinc SOD located in the
intermembrane space [6]. Notably, unlike O
2
•
−
, H
2
O
2
is stable and uncharged, and thus able to leave
the mitochondrion to mediate cytosolic cell signaling. This signaling occurs predominantly through the
oxidation of either metal cofactors or reduced thiols on cytosolic proteins, changing their function. It is
now well recognized that mitochondrial H
2
O
2
regulates a number of signaling pathways by this
mechanism. While discussion of each of these specific pathways is outside the scope of this article, we
direct the reader to reviews of the essential role of mitochondrial ROS signaling in hypoxic adaptation
[20, 24], apoptosis [25], regulation of phosphorylation signaling [26, 27], and cell growth and
differentiation [28, 29].
The concentration of H
2
O
2
that leaves the mitochondrion is regulated by two main antioxidant systems
in the mitochondrial matrix: the glutathione and thioredoxin/peroxiredoxin systems (Figure 2).
Glutathione (GSH) is oxidized by H
2
O
2
to form the glutathione disulfide (GSSG), a reaction catalyzed by
glutathione peroxidase (GPx). GSSG is then reduced back to GSH by Glutathione Reductase (GR).
Similarly, H
2
O
2
oxidizes a cysteine residue in the catalytic site of peroxiredoxin (PRx), which forms a
disulfide bridge with a neighboring cysteine. The reductive power of thioredoxin (TRx) reduces PRx
through a disulfide exchange reaction and TRx is then reduced by Thioredoxin reductase (TR).
Importantly, both GR and TR require NADPH for their reductive activity. The pool of reduced NADPH is
maintained by several enzymes in the mitochondrial matrix including malic enzyme (ME), glutamate
dehydrogenase (GDH), and isocitrate dehydrogenase (IDH2) [30]. Additionally, membrane-associated
nicotinamide nucleotide transhydrogenase (NNT) is particularly important in maintaining NADPH pools
through its function of pumping protons into the matrix to regenerate NADPH by coupling oxidation of
NADH to the reduction of NADP
+
. Importantly, the ability of NNT to renew the pool of available NADPH
is dependent on ΔΨ. States of low ΔΨ decrease the amount of available reduced NADPH which then
decreases the amount of reduced glutathione and thioredoxin to buffer H
2
O
2
.
Measurement of mitochondrial bioenergetics in human disease
In the previous sections we have outlined the mechanisms by which the ETC functions to generate ATP
and produce ROS, highlighting the interplay between these two processes. While physiological function
of the ETC generates ROS at levels required for homeostatic signaling, ETC dysfunction, particularly
that which decreases mitochondrial energy production and enhances ROS generation, has been linked
to the onset and development of a number of biological changes including obesity and aging, and
pathology in all organ systems including cardiovascular diseases. Thus, accurate and efficient
assessment of energetic function and mitochondrial ROS production in humans could potentially enable
the diagnosis of disease and the mechanistic understanding of pathogenesis. In this section, we focus
on a translational approach to the measurement of ETC function. Currently, no standard methodologies
exist to directly measure mitochondrial ROS production in humans. Thus, we will give an overview of
methodology to assess mitochondrial energetic function in humans.
A number of non- and minimally-invasive technologies such as near-infrared spectroscopy (NIRS),
magnetic resonance spectroscopy (MRS), and positron emission tomography (PET) can be utilized to
indirectly assess mitochondrial function in humans (Figure 3). Near-infrared spectroscopy capitalizes
on the principle that near-infrared light is absorbed at differential wavelengths by the oxygenated and
deoxygenated heme groups of hemoglobin and myoglobin, allowing the calculation of tissue oxygen
consumption, which is proportional to mitochondrial respiration. While NIRS is relatively inexpensive,
one major disadvantage of this method is that near infrared light applied non-invasively cannot reach
solid organs such as the heart, and thus is best utilized for exercising large muscles closer to the skin
[31]. Magnetic resonance spectroscopy is a non-invasive and ionizing-radiation-free method that can be
used in the heart and other solid organs to measure the presence of metabolites such as choline-
containing compounds, creatine, glucose, alanine and lactate by measuring the resonance of MR-
visible isotopes [32].
31
Phosphorous (
31
P) is most frequently utilized to visualize concentrations of ATP,
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phosphocreatine, and inorganic phosphate, which can provide a dynamic measure of ATP production in
the tissue.
13
Carbon (
13
C) and proton NMR (
1
H) can also be used to measure intermediates in the TCA
cycle [33, 34]. In contrast to NMR, PET imaging employs the detection of injected positron-emitting
radionuclide tracers to measure the accumulation or consumption of metabolic intermediates [35, 36].
While PET and MRS provide a solid assessment of energetics, both techniques require costly
equipment and a high degree of training for data acquisition and analysis. In addition, PET has the
added obstacle of radiopharmaceutical production and the exposure of patients to radiation. Further,
these technologies are highly specialized and not suited for high throughput clinical application, limiting
their mainstream use.
The gold standard for the direct assessment of mitochondrial ETC function in humans is the
measurement of mitochondrial function in small biopsies of tissue [37, 38]. Once obtained, biopsies are
most frequently subjected to measurement of mitochondrial oxygen consumption, which provides
information on multiple facets of ETC function. The Clark-type oxygen electrode, developed in the
1960’s, enabled the polarographic measurement of oxygen consumption rate (OCR) by isolated
mitochondria or cells in a sealed chamber. This technology was later utilized by the Oroboros O2k
system to provide amperometric measurements of O
2
consumption with maximal sensitivity and
precision. In addition, the Oroboros O2k contains closed, air-tight reaction chambers that minimize O
2
back-diffusion, thus limiting over estimations of OCR. Further, it allows the measurement of OCR at
fixed oxygen tensions and the potential for simultaneous measurement of multiple parameters including
pH, Δψ
m
, [Ca
2+
], and NO in the same chamber. More recently, development of the Seahorse
extracellular flux (XF) analyzer, has enabled the measurement of OCR in an intact monolayer of cells in
a high throughput multi-well format. The Seahorse analyzer simultaneously measures extracellular pH
to enable the estimation of glycolytic rate. Despite key differences between the two systems, such as a
greater sample size required for the Oroboros and the need for a monolayer for the Seahorse versus
sample in suspension for the Oroboros, both systems can be used with a series of pharmacologic
mitochondrial modulators to provide a profile of ETC function (Figure 4). In a typical bioenergetic
profile, basal OCR is measured to determine the current turnover of the ETC. This is followed by the
addition of oligomycin A, an inhibitor of the ATP synthase, in order to measure OCR that is not
contributing to ATP production (proton leak). A protonophore, such as trifluoro-methoxy -carbonyl
cyanide-4-phenylhydrazone (FCCP) is then added to uncouple respiration, yielding the maximal rate of
OCR. Finally, a mitochondrial inhibitor is added such that the rate of non-mitochondrial OCR is revealed
[39]. Notably, if intact cells are being utilized, pharmacologic inhibitors of glucose, fatty acid, or amino
acid oxidation/entry into the mitochondrion can be utilized (as shown in Figure 1) to determine the
source of substrate for the ETC.
While the respirometric techniques described above can be applied to any cell type, traditionally for
human studies, skeletal muscle biopsies or cells cultured from skin grafts have been most frequently
used as a source of viable tissue/mitochondria. However, the invasiveness of obtaining these samples
is prohibitive of their widespread use for routine mitochondrial measurement. Recently, a number of
labs have advanced the concept of performing respirometric measures in circulating blood cells,
including platelets and peripheral blood mononuclear cells, as an assessment of systemic bioenergetic
function [40-42]. Blood cells contain fully functional mitochondria, and are abundant, self-renewing, and
minimally invasive to obtain. Our group and others have successfully employed Seahorse XF and/or
Oroboros to circulating cells to demonstrate that this methodology is feasible [43, 44], reproducible in
human populations [40], and that bioenergetics of circulating cells reflect bioenergetics in solid tissues
such as the heart [42], skeletal muscle [42, 45], brain [46], and lungs [47]. Further, these studies
demonstrate alterations in blood cell bioenergetics in a number of natural biological conditions (e.g
aging) or pathologies, and correlate with physical or clinical parameters of these conditions [45, 47-49].
Altered platelet bioenergetics have been measured in neurologic [46], metabolic [46], hematologic [44],
cardiopulmonary [47, 49, 50], and infectious diseases [51] (Figure 5).
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One example of platelet alterations in cardiovascular disease is in sickle cell disease in which a
bioenergetic “screen” of platelets from a cohort of sickle cell patients demonstrated that basal OCR was
decreased but maximal uncoupled OCR was unchanged compared to age and race matched healthy
subjects [44]. This pattern of alteration potentially suggested that while the capacity for respiration was
unchanged in sickle cell patients, Δψ
m
was higher (leading to decreased basal OCR). Direct
measurement of Δψ
m
confirmed this hypothesis, and further measurement of individual complex
activities demonstrated the increased Δψ
m
was due to Complex V inhibition and resulted in increased
O
2
•
−
production. This production of mitochondrial O
2
•
−
is linked to platelet activation in sickle cell
patients (REF 44) and the measurement of Complex V activity and markers of ROS in platelets from
these patients can now be utilized to not only elucidate the pathogenesis of the disease but also test
the ability of potential therapeutics to attenuate this mitochondrial-driven oxidative pathogenesis [52].
These data demonstrate the potential for the measurement of ETC function to reveal changes in both
mitochondrial energetic and redox function and highlight the potential use of blood bioenergetic
measurements as a translational and clinical tool.
In summary, the mitochondrial ETC is a redox hub that regulates cellular homeostasis through the
production of ATP and the generation of ROS, two processes that are intimately linked. Given that
alterations in both of these processes have been associated with the pathogenesis of a myriad of
diseases spanning all organ systems, the ability to accurately and efficiently measure ETC function in
humans may provide useful diagnostic and mechanistic information.
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Funding sources: This work was supported by NIH RO1 HL133003-01A1 and funds from the
Hemophilia Center of Western Pennsylvania to SS and AHA 19POST34380956 to ACB.
Conflicts of interest: None
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Figure 1: (A) Substrate supply for mitochondrial respiration. Glucose, fatty acids, and amino acids
(glutamine shown here) undergo catabolism to feed into the tricarboxylic acid cycle (TCA cycle), which
generates substrates for the electron transport chain (ETC). Glucose is metabolized by glycolysis. Red
inhibitory signs denote pharmacologic agents utilized to inhibit specific sources of substrate in order to
delineate substrate supply to the ETC, particularly in in vitro respirometric assays as outlined in the text.
2-deoxy-D-glucose (2-DG) inhibits glycolysis. Etomoxir inhibits fatty acid entry into the mitochondria
through carnitine palmitoyltransferase 1 (CPT1). Bis-2-(5-phenylacetamido-1,3,4-thiadiazol-2-yl)-ethyl
sulfide (BPTES) inhibits glutaminase (GLS) to inhibit glutamine metabolism to glutamate. 2-Cyano-3-(1-
phenyl-1H-indol-3-yl)-2-propenoic acid (UK5099) inhibits the mitochondrial pyruvate carrier (MPC) to
prevent pyruvate entry into the organelle. Within the mitochondrial matrix, acetyl-CoA is produced from
pyruvate or from the beta-oxidation of fatty acids and serves as a point of entry to the TCA cycle.
Citrate synthase converts acetyl-CoA and oxaloacetate to citrate. Citrate is converted to isocitrate by
aconitase, and isocitrate is in turn oxidized into α-ketoglutarate, which reduces NAD
+
to NADH. Next, α-
ketoglutarate which is derived from the hydrolysis of extracellular amino acid glutamine to glutamate via
the glutaminase (GLS) I and II pathways, enters the TCA cycle. α-ketoglutarate undergoes oxidative
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decarboxylation with NAD
+
and CoA-SH (CoA not attached to an acyl group) to irreversibly form
succinyl-CoA, NADH, CO
2
and H
+
. The succinyl-CoA is then hydrolyzed to form succinate, CoA-SH and
energy in the form of GTP. Succinate is oxidized to fumarate by Complex II/succinate dehydrogenase
while converting FAD to FADH
2
. Hydration of fumarate by fumarase results in the formation of L-malate
that then becomes oxidized to form oxaloacetate, NADH and H
+
. At this point the cycle is complete and
the aldol condensation of oxaloacetate with acetyl CoA and water can restart the TCA cycle.
Throughout the TCA cycle, the reduction of NAD
+
to NADH is coupled to the release of CO
2
.
(B) Mitochondrial oxidative phosphorylation (OXPHOS). The mitochondrial electron transport chain
(ETC) consists of five protein complexes integrated into the inner mitochondrial membrane. The TCA
cycle in the mitochondrial matrix supplies NADH and FADH
2
to the ETC, each of which donates a pair
of electrons to the ETC via Complexes I and II respectively. The transfer of electrons from Complex I to
the Q cycle results in a net pumping of 4 protons across the inner membrane into the intermembrane
space (IMS). Of note, Complex II does not span the inner membrane and does not participate in proton
translocation. The electrons from either Complex I (2 electrons) or Complex II (2 electrons one at a
time) are donated to ubiquinone (Q) which is reduced to ubiquinol (QH
2
). Ubiquinol is oxidized by
Complex III allowing one electron at a time to continue the journey through cytochrome c (c). For every
electron transferred to cytochrome c, 2 protons (H
+
) are pumped into the IMS, resulting in 4H
+
pumped
into the IMS for every electron pair moved through the cycle. Cytochrome c transports electrons to
Complex IV where molecular oxygen acts as a terminal electron acceptor and is reduced to water. The
reduction of one molecule of O
2
requires 4 electrons. The reduction of O
2
to H
2
O results in the pumping
of 4 protons to the IMS, but 2 protons are consumed in the process, netting a total of 2 H
+
pumped into
the IMS at Complex IV. The movement of protons from the mitochondrial matrix into the intermembrane
space in response to electron transfer creates a protonmotive force (Δp), which is the proton
concentration (pH) combined with the electrochemical proton gradient known as the mitochondrial
membrane potential (ΔΨ). The membrane potential is dissipated by the re-entry of H
+
back into the
matrix through Complex V, which is coupled to the production of ATP from ADP. In contrast, uncoupled
respiration due to proton leak is facilitated by adenine nucleotide translocase (ANT) and uncoupling
proteins (UCP) and dissipates membrane potential at the expense of ATP production.
Abbreviations: protonmotive force, Δp; oxaloacetic acid, OAA, alpha-ketoglutarate, α-KG; carbon
dioxide, CO
2
; reduced flavin adenine dinucleotide reduced FADH
2
; glutaminase, GLS; carnitine palmityl
transferase, CPT; carnitine-acylcarnitine translocase, CACT; mitochondrial pyruvate carrier, MPT.
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Figure 2: Sites of mitochondrial ROS production and antioxidant systems. Seven major ETC sites
of ROS generation are shown here, with red lines indicating on which side of the membrane the ROS
are formed. Superoxide (O
2•–
) is the primary ROS generated by the ETC. O
2
•
−
is dismutated to H
2
O
2
by
MnSOD in the matrix and Cu/Zn SOD in the intermembrane space. Also within the matrix is the
glutathione peroxidase (GPx)/ glutathione reductase (GR) system and catalase, found in liver and
cardiac tissues. The peroxiredoxin (Prx)/ thioredoxin (TrxR) system overlaps between the cytosol and
the matrix. NADPH/NADP+ renews these antioxidant systems with its reducing potential. Matrix
enzymes (malic enzyme (ME), glutamate dehydrogenase (GDH), and isocitrate dehydrogenase (IDH2))
and the inner membrane-associated nicotinamide nucleotide transhydrogenase (NNT) regenerate the
pool of NADPH.
Abbreviations: mitochondrial glycerol-3-phosphate dehydrogenase, mGPDH; dihydroorotate
dehydrogenase, DHODH, electron transfer flavoprotein oxidoreductase, ETFQO; superoxide, O
2
•-
;
hydrogen peroxide, H
2
O
2
; water, H
2
O; glutathione, GSH; glutathione disulfide, GSSG; glutathione
peroxidase, GPx; GR; thioredoxin reductase,TR; ; reduced thioredoxin, Trx(SH)
2
; oxidized thioredoxin
Trx-S
2
; peroxiredoxin, Prx; reduced peroxiredoxin, Prx(SH)
2
; oxidized peroxiredoxin, Prx-S
2
; malic
enzyme, ME; glutamate dehydrogenase, GDH; NADP
+
-dependent isocitrate dehydrogenase, IDH2, and
nicotinamide nucleotide transhydrogenase, NNT.
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Figure 3: Minimally-invasive methods of human bioenergetic assessment. Near Infrared
Spectroscopy (NIRS) operates on the principle that near-infrared light (700–900 nm) penetrates tissue
with little scatter and is absorbed by heme-containing groups (e.g., hemoglobin, myoglobin, and the
heme-containing prosthetic groups within the mitochondrial electron chain complexes) in an oxygen-
dependent manner. Measurement of changes in absorbance can be used to reflect changes in tissue
oxygenation and mitochondrial oxidative capacity. Magnetic Resonance Spectroscopy (MRS) utilizes
magnetic resonant-visible isotopes (eg,
1
H,
31
P and
13
C), with each resonating at characteristic
frequencies within a magnetic field. The distinct resonance of known isotopes creates a signature for
identification of compounds within living tissues. MRS can distinguish products of glycolysis, creatine
metabolism, choline metabolism, and amino acid metabolism. Positron Emission Tomography (PET)
utilizes a radioactive isotope tracer such as glucose analogue 18-fluorodeoxyglucose ([
18
F]FDG) or
other radio-labeled metabolites (acetate, choline, methionine or glutamine). The tracer is administered
intravenously and becomes trapped within metabolically active cells. The nucleus of the isotope emits
positrons as it decays. The positrons make contact with electrons to generate high-energy photons
(gamma rays) that are detected by the PET camera and translated into an electrical signal to produce
an image. The image that is produced is dependent on the metabolism of the cells/tissues that take up
the isotope.
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Figure 4: Respirometric methods of bioenergetic assessment. Panel (A) shows the common
pharmacologic modulators of the respiratory chain utilized to generate a bioenergetic profile (along with
their chemical structures), the target of these modulators, the bioenergetic parameter measured in their
presence, and the significance of each parameter. (B) A typical bioenergetic profile generated in the
Oroboros system showing oxygen concentration in the chamber over time (blue trace) and the
calculated oxygen flux (red trace). (C) A typical bioenergetic profile generated by the Seahorse XF
analyzer in which oxygen consumption rate (OCR) over time is shown.
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Figure 5: Changes in platelet bioenergetics in human disease. The bioenergetic profiles of platelets
reflect alterations that occur at a systemic level during human disease states. Bioenergetic profiles
have been measured in platelets isolated from patients with a wide array of diagnosed diseases. This
figure demonstrates published platelet bioenergetic alterations in neurologic disease including
Alzheimer’s disease [53] and bipolar disorder [54], metabolic changes including type II diabetes [55]
and advanced age [45], cardiopulmonary diseases including pulmonary hypertension [50, 56] and
asthma [47, 57], sickle cell disease as a hematologic disease [44], and infectious disease such as
sepsis [58].
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CONFLICT OF INTEREST: None
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