Mitochondrial Dynamics in the Regulation
of Nutrient Utilization and Energy Expenditure
Marc Liesa1and Orian S. Shirihai1,2,*
Medicine, 650 Albany Street, Boston, MA 02118, USA
2Department of Clinical Biochemistry, School of Medicine, Ben Gurion University, Beer-Sheva 84103, Israel
quality control might not be the only task carried out by mitochondrial dynamics. Recent studies link mito-
chondrial architecture as a mechanism for bioenergetic adaptation to metabolic demands. By favoring either
connected or fragmented architectures, mitochondrial dynamics regulates bioenergetic efficiency and
energy expenditure. Placement of bioenergetic adaptation and quality control as competing tasks of mito-
chondrial dynamics might provide a new mechanism, linking excess nutrient environment to progressive
mitochondrial dysfunction, common to age-related diseases.
As our relationship with mitochondria evolves, we remain fasci-
conditions: aging and metabolic diseases. While aging involves
insufficiency of mitochondrial quality control and turnover mech-
anisms (such as autophagy), type 2 diabetes and obesity are
influenced by the ability of the organism to deal with excess
nutrient environment. The observation that both conditions are
impactedbythe durationof exposureto excessnutrient environ-
ment raises the question, are the tasks of handling nutrients in
excess and maintaining quality control ever in conflict? In this
review, we discuss evidence to support a hypothesis that adap-
tation to excess nutrient environment interferes with quality
control functions and, as a result, affects mitochondrial function
in a magnitude that reflects the duration to which the organism
was exposed to excess nutrient environment.
In response to changes in energy demand and supply, the
organism adapts by adjusting both its capacity and/or efficiency
of ATP production. Mitochondrial bioenergetic efficiency is
defined as the ATP produced in the mitochondria per molecule
of nutrient (Figure 1), and mitochondrial ATP synthesis capacity
is defined as the rate at which ATP is produced per unit of time.
As an adaptation to excess nutrients, the organism recruits
mechanisms to utilize nutrients first by storage and then by
waste (heat generation). While spending time at the gym may
be the appropriate way to waste energy and keep healthy,
reducing bioenergetic efficiency might enable energy waste in
tissues other than muscle and in individuals that are less
compatible with the gym.
Studies in the field of mitochondrial dynamics have identified
an intriguing link between energy demand and supply balance
and mitochondrial architecture. Cells exposed to a rich-nutrient
environment tend to keep their mitochondria in a separated
(fragmented) state, and mitochondria in cells under starvation
tend to remain for a longer duration in the connected (elongated)
state (Molina et al., 2009; Gomes et al., 2011). Thus, it appears
that bioenergetic adaptation that involves changes to bioener-
getic efficiency and mitochondrial ATP synthesis capacity also
implies remodeling of mitochondrial architecture.
However, bioenergetic adaptation is not the only mitochon-
drial task that involves changes to mitochondrial architecture.
A vital task that engages the fusion and fission machinery is
the mitochondrial life cycle (Twig et al., 2008a). The mitochon-
drial life cycle represents continuous changes to mitochondrial
architecture through fusion and fission events. These brief tran-
sitions between connected and separated mitochondria enable
tion of damaged material, thereby maintaining a healthy mito-
chondrial population. One can appreciate that the life cycle of
mitochondria would be compromised if mitochondrial fusion or
fore, under certain nutrient environments, bioenergetic adapta-
tion and quality control might represent conflicting tasks.
That mitochondrial quality control has evolved within the same
mechanism that controls for bioenergetic efficiency is not
surprising, given the understanding that a low-nutrient environ-
ment (caloric restriction) may support increased longevity.
Adaptation of bioenergetic efficiency and ATP synthesis
capacity to nutrient availability differs among tissues and is inti-
mately linked to their specific physiology. Thus, we will focus on
three paradigmatic tissues that show different bioenergetic effi-
ciencies and mechanisms of adaptation to nutrient availability:
(1) Brown adipose tissue: When stimulated, brown adipo-
cytes can go through an acute and robust transition
from high to low bioenergetic efficiency. Under these
stimulatory conditions, energy obtained from mitochon-
drial nutrient oxidation is almost entirely directed toward
heat production rather than ATP synthesis (reviewed in
Cannon and Nedergaard, 2004).
(2) Muscle: Muscle cells harbor higher bioenergetic effi-
ciency as compared to either beta cells (Affourtit and
Brand, 2006) or stimulated brown fat. In the contracting
red muscle, nutrient oxidation is primarily directed
Cell Metabolism 17, April 2, 2013 ª2013 Elsevier Inc.
towards production of ATP in the mitochondria (Chappell
and Perry, 1954) to support contraction. Thus, the oxida-
tive muscle is a good example of high mitochondrial ATP
synthesis capacity and likely high bioenergetic efficiency
(Marcinek et al., 2004).
(3) Beta cells: Mitochondria in pancreatic beta cells serve as
nutrient sensors and signal generators for insulin secre-
tion. Nutrients are ‘‘sensed’’ through their metabolism,
which involves nutrient oxidation mediated by beta cell
mitochondria (Ashcroft et al., 1984; reviewed in Deeney
et al., 2000). Therefore, bioenergetic efficiency is ex-
pected to be highly regulated to allow proper insulin
Although the mechanisms for tissue-specific differences in
bioenergetic efficiency are understood to a certain extent, less
is known about the contribution of mitochondrial dynamics to
tissue and diet-dependent bioenergetic efficiency and mito-
chondrial ATP synthesis capacity. Mitochondrial dynamics is
a concept that comprises mitochondrial architecture resulting
from movement, tethering, fusion, and fission events. Multiple
important for cell viability, senescence, mitochondria health,
bioenergetic function, quality control, and intracellular signaling
(reviewed in Liesa et al., 2009; reviewed in Twig et al., 2008b).
On the other hand, we are now beginning to understand how
nutrients and the cellular metabolic state are regulating mito-
chondrial dynamics in different tissues and vice versa, particu-
larly in the beta cell, brown adipose tissue, and muscle (Molina
et al., 2009; Quiro ´s et al., 2012; Sebastia ´n et al., 2012). Along
with this, the relevance of mitochondrial dynamics in the specific
physiology of different tissues has only been revealed recently,
mostly thanks to different mouse models harboring tissue-
specific deletions of core components regulating mitochondrial
dynamics (Chen et al., 2007, 2010; Chen et al., 2011; Ishihara
et al., 2009; Sebastia ´n et al., 2012; Wakabayashi et al., 2009;
Zhang et al., 2011).
In this context, the aim of this review is to summarize the
current understanding of mitochondrial bioenergetic function
and efficiency regulation by nutrient availability and energy
demand in health and disease. We will discuss how mitochon-
drial dynamics may be required for proper adaptation to the
diverse bioenergetic requirements. In the last section, we will
provide a model in which adaptation to sustained exposure to
nutrient excess results in prolonged changes to mitochondrial
dynamics. These changes can impact mitochondrial quality
control and thereby contribute to the mitochondrial dysfunction
characteristic of metabolic and other age-related diseases.
Regulation of Cellular Bioenergetics by Nutrients
How Can Bioenergetic Efficiency Affect Cellular
Functionality and Viability?
Intuitively, it is expected that conditions of limited nutrient avail-
ability will increase the ratio of ATP produced per nutrient
consumed, thereby reducing and optimizing the consumption
of nutrients. Mechanisms to increase energy efficiency are ex-
pected to diverse between tissues that are primarily relying on
‘‘anaerobic’’ glycolysis and those that are relying primarily on
oxidative metabolism for the production of ATP.
In this regard, recent studies performed in transformed cell
lines demonstrate that starvation increases mitochondrial ATP
synthesis capacity (ATP production per unit of time). This
increase involves the formation of ATP synthase dimers at the
cristae curvatures, which show higher activity (Gomes et al.,
2011). This result may represent a shift from ‘‘anaerobic’’
glycolysis (to lactate) toward mitochondrial respiration under
starvation, as respiration can produce more ATP per molecule
of glucose. In oxidative cell types, one would also expect the
activation of mechanisms that increase mitochondrial bioen-
ergetic efficiency to ensure survival under limited availability of
nutrients. Mechanisms enhancing mitochondrial bioenergetic
efficiency have not been described in detail under these condi-
tions. On the other hand, increased mitochondrial ATP synthesis
capacity reported in transformed cell lines (Gomes et al., 2011)
was associated with and dependent on changes in mitochon-
drialdynamics, whichwere presented asdecreased fissionrates
and mitochondrial elongation. This change in dynamics
suggests that elongation could be an active mechanism contrib-
uting to increased mitochondrial bioenergetic efficiency.
energy obtained from nutrient oxidation toward heat production,
most commonly by increased uncoupled respiration. Decreased
bioenergetic efficiency may serve as a protective mechanism
from the detrimental effects associated with nutrient overload.
This is achieved through the reduction of reactive oxygen
species (ROS) production and by the enhanced removal of
excess nutrients and their potentially cytotoxic metabolites
Balanced nutrient availability
Figure 1. Regulation of Cellular Bioenergetic Efficiency under
Conditions of Nutrient Excess
In the balanced statefuel/nutrient ‘‘supply’’is sufficientto sustain energy (ATP)
‘‘demand.’’ Under this condition, ‘‘waste’’ or inefficiency in the form of heat is
minor. Nutrient excess, characterized by ‘‘excessive supply’’ in the absence of
a parallel increase in ‘‘demand,’’ represents a situation in which the energy
required to satisfy ATP demand is lower than the available energy. This is
compensated for by addition of an energy sink that does not involve ATP
synthesis. This component is inefficiency/waste in the form of heat. The major
mechanism for inefficiency/waste in the form of heat is mitochondrial proton
‘‘leak.’’ This mechanism can slow down nutrient accumulation and prevent the
Cell Metabolism 17, April 2, 2013 ª2013 Elsevier Inc.
The flow of electron-mediated proton translocation in the
respiratory chain can be compared to a flow of water in a garden
hose (see the ‘‘Understanding Mechanisms of Bioenergetic Effi-
ciency and Changes in ATP Synthesis Capacity by Respiration
Studies’’ section for a more detailed bioenergetics description).
NADH, resulting from nutrient oxidation, feeds the hose inlet with
water, while ATP synthase controls the hose final outlet. The
pressure that the flow of water generates in the hose is the mito-
chondrialmembrane potential(Dcm). Theflowof waterandpres-
sure in the hose are determined by the rates of NADH production
and ATP synthesis. The minimum and maximum values of pres-
sure that the hose can hold are determined by the material and
integrity of the hose, not by the flow of water or the inlets and
outlets (i.e., the range of Dcmin mV is determined by thermody-
namics and theintegrity of the organelle). ATP synthesisis deter-
mined by ATP demand, meaning that the hose outlet is
sure, we would not have to be concerned with any parameter
beyond ATP demand. However, this is not the case. The hose,
as it turns out, has some cuts through which water can escape,
whenpressurebuilds up. Apressurevalve thatcan divert excess
waterthroughasafeconduit can reducethe pressureinthehose
and prevent water leakage through the ‘‘cuts’’ in the hose
(increasing or maintaining the flow of water). In our analogy,
the escape of water through the cuts represents the escape of
electrons to produce ROS. The pressure valve represents the
combination of inducible and inherent uncoupled respiration,
the latter being caused by the inherent proton leak of the inner
membrane. Inducible uncoupling can include uncoupling protein
1 (UCP1) activation in brown fat and the permeability transition
pore opening. Inherent, nonactivated, proton leak is directly
(but nonlinearly) correlated to the membrane potential and is
mediated, in part, by inner membrane proteins (such as adenine
nucleotide translocator or nonactivated UCP1 in brown fat)
(Parker et al., 2009). The balance between ATP demand and
nutrient supply determines both the rate of ATP synthesis and
the level of ROS produced by mitochondria.
Different tissues employ different mechanisms in their
response to nutrient overload. The selection of specific compen-
satory mechanisms allows each tissue to maintain its unique
primary function, while minimizing side effects related to ROS
production. In certain cell types, compensatory mechanisms
are placed upstream of the mitochondria, preventing their expo-
sure to high levels of fuel. However, in beta cells, brown adipose
tissue, and muscle, mounting evidence suggests that conditions
of nutrient excess that increase fuel availability to the mitochon-
dria might modulate bioenergetic efficiency and mitochondrial
ATP synthesis capacity (Koves et al., 2008; Bonnard et al.,
2008; Rothwell and Stock, 1979; Wikstrom et al., 2007).
Understanding Mechanisms of Bioenergetic Efficiency
and Changes in ATP Synthesis Capacity by Respiration
Mitochondria from any tissue can provide energy in the form of
ATP as a result of nutrient oxidation (Chance and Williams,
1955; Mitchell, 1961). Oxidation of nutrients will provide elec-
trons to the mitochondrial electron transport chain (constituted
tial transport of electrons from complex I or II to III and IV
extrudes protons from the matrix to the intermembrane space,
generating an electrochemical gradient (DmH+) resulting in
a difference in charge (Dc) and in proton concentration (DpH).
utor to DmH+(reviewed in Nicholls and Ferguson, 2002). In intact
mitochondria, maximal and minimal Dcmvalues are around 225
and 90 mV, respectively. This range in mV is dictated by the
thermodynamic stability of functional mitochondria and repre-
sents the balance between proton extrusion and re-entry.
Energy from proton re-entry through complex V is used for the
synthesis of ATP from ADP. The state at which isolated mito-
chondria are synthesizing ATP at maximal rates is named state
3 (Chance and Williams, 1955), and it occurs at intermediate
Dcmvalues (?140 mV). As such, this state is characterized by
a high rate of both proton extrusion and re-entry (reviewed in
Nicholls and Ferguson, 2002).
Proton re-entry through mechanisms that do not involve
complex V and ATP synthesis are referred to as uncoupled
respiration. Uncoupled respiration results in the generation of
heat and is not controlled by ATP turnover (reviewed in Nicholls
and Ferguson, 2002). It is important to distinguish between two
different types of respiratory states resulting from uncoupling.
These two respiratory states determined in isolated mitochon-
dria show major functional differences and might mimic respira-
tory states under different physiological conditions in vivo:
(1) Respiration controlled by inherent proton leak. This is
typically measured in vitro, in isolated mitochondria in
which ATP synthesis has been inhibited either by ADP
exhaustion (state 4) or by the use of complex V inhibitor
olygomycin. It is also referred to as respiration controlled
by basal proton conductance (Parker et al., 2009) and can
mimic physiological conditions of decreased mitochon-
drial ATP demand and high nutrient availability.
(2) Respiration controlled by inducible uncouplers. This type
of uncoupled respiration can be experimentally mimicked
molecules located in the inner mitochondrial membrane,
such as UCP1. The activation of these endogenous
uncouplers takes the control of respiration from ATP
controlled by the capacity of the respiratory chain and
by the availability of mitochondrial fuels. This type of
respiration is also characterized by decreases Dcm
values, due to increased proton re-entry. Itis also referred
to as inducible proton conductance (Parker et al., 2009).
A key difference between these two types of uncoupled respi-
ration is the membrane potential at which they are conducted.
Mitochondrial respiration controlled by inherent proton leak,
which occurs in coupled mitochondria under conditions of low
ATP synthesis and high nutrient availability, is associated with
The high Dcmvalues result from a combination of decreased
rates of proton re-entry through ATP synthase and low values
of proton conductance contributed by the inherent proton leak.
The combination of these effects maintains Dcmvalues within
the range dictated by thermodynamic stability of intact mito-
chondria. This state is associated with relatively higher ROS
generation, as a consequence of the increase in Dcm.
Cell Metabolism 17, April 2, 2013 ª2013 Elsevier Inc.
In marked contrast, mitochondria treated with uncouplers
(such as FCCP) have decreased Dc value, which causes an
increase in respiration rates to values higher or close to state
3. The concomitant increase in respiration maintains Dcmvalues
within therangeofthermodynamic stability(?90–120mV).Inthis
case, absolute values of calories from nutrients used for heat
generation will be higher in uncoupler-induced respiration
compared to inherent proton leak controlled respiration. There-
fore, respiration that is activated by uncouplers is characterized
by decreased bioenergetic efficiency and lower mitochondrial
ATP synthesis capacity, as it drives nutrient oxidation toward
heat generation. Furthermore, it is associated with lower ROS
production, as Dcmvalues are reduced. The description of these
basic differences between the two types of uncoupled respira-
tion is relevant to understanding the physiological conse-
quences of nutrient-mediated changes in respiration rates,
Dcmand mitochondrial dynamics described in the ‘‘Relationship
between Bioenergetic Efficiency and Mitochondrial Dynamics’’
Nutrient Availability Control of Mitochondrial
Mitochondrial respiration is controlled by three different pro-
cesses: (1) ATP turnover, determined by cellular ATP consump-
tion and matrix ADP levels; (2) substrate utilization, determined
by fuel availability inside the mitochondrial matrix and its oxida-
tion to generate NADH, FADH2; and (3) proton leak, determined
by the inherent permeability of the inner membrane to protons.
Understanding the contribution of each of these processes is
essential to predict under which physiological and mitochondrial
respiratory states, nutrient availability will be determining mito-
chondrial respiration and Dcm.
In isolated mitochondria under state 3, where maximal ATP
synthesis rates are induced, both nutrient utilization and ATP
turnover exert a similar control over respiration and thus over
Dcm. This control can be quantified as the control coefficient
over the mitochondrial respiratory flux. A value of 1 for this
coefficient represents an absolute control of a process over
respiration. Under state 3, ATP turnover was found to have
a control coefficient value of >0.5, while nutrient utilization has
a control coefficient of <0.4. (see Hafner etal., 1990). This finding
in isolated mitochondria supports the idea that ATP demand has
the main control over the rate of mitochondrial respiration in
intact cells under physiological conditions, while mitochondrial
nutrient availability and the inherent proton leak have relative
lower control over respiration rates.
However, in an intact cell, the metabolic processes providing
NADH/FADH2to the mitochondrial matrix, including glycolysis,
fatty acid oxidation, and TCA cycle, can control respiration with
a flux control coefficient over respiration between 0.15–0.3
under resting conditions (reviewed in Nicholls and Ferguson,
2002; Hafner et al., 1990). Therefore, although ATP turnover
has a major influence controlling respiration and membrane
potential (control coefficient value 0.5), under conditions of
high ATP demand, nutrient utilization and its availability can still
have a significant control over respiration and the exact mito-
chondrial Dc values in intact cells. Furthermore, nutrient avail-
ability will have even a greater control over respiration after
the induction of uncoupling with either pharmacological uncou-
plers or by stimulation of uncoupling mechanisms such as
UCP1 in intact cells and in isolated mitochondria, as ATP turn-
over will have a reduced control over mitochondrial respiration
under these conditions. Overall, the fact that the control coeffi-
cient of each process over mitochondrial respiratory flux can
vary suggests that under certain physiological scenarios mito-
chondrial nutrient availability may control the mitochondrial
Dc (within the range dictated by thermodynamics; around 90–
Of particular relevance for this review, in certain cell types,
including nutrient sensors such as the beta cell, nutrient avail-
ability has a higher flux control coefficient and greater control
over mitochondrial respiration and membrane potential than in
other cell types (i.e., muscle cells). Consistent with this, recent
evidence confirmed previous findings that mitochondrial hyper-
polarization is proportional to the increase in extracellular
nutrient concentration (glucose and pyruvate) in beta cell line
(Goehring et al., 2012; Wikstrom et al., 2007; Danial et al.,
2008; Heart et al., 2006).
Furthermore, uncoupling protein 1 in the brown adipocyte
conductance and thereby mitochondrial respiration can be acti-
vated by nutrients per se (Rial et al., 1983; Parker et al., 2009;
Shabalina et al., 2008). That brown adipose tissue evolved to
utilize fatty acids as a signal for nutrient wasting brings up
a potential general concept that nutrients with high caloric
content can activate thermogenesis and exert important control
mechanism could promote ‘‘nutrient wasting’’ in the form of heat
generation under conditions of increased nutrient supply
(Figure 1). Such regulatory pathways decreasing bioenergetic
efficiency could exist in other tissues, but likely through other
mediators and/or regulators. These regulatory pathways are ex-
pected to be relevant in nutrient sensors, which harbor high
nutrient permeability. These mechanisms could involve and/or
require changes in mitochondrial dynamics, as discussed in
the ‘‘Relationship between Bioenergetic Efficiency and Mito-
chondrial Dynamics’’ section.
In this regard, obesity and diabetes research have put forward
mitochondrial ‘‘nutrient wasting’’ in the form of heat as an impor-
tant concept in metabolic adaptation. This concept is based on
chondrial nutrient oxidation in certain tissues, including muscle,
brown adipose tissue, or beige adipocytes, could potentially
compensate for the deregulated energetic balance associated
with nutrient excess (Levine et al., 1999; Schutz et al., 1984;
Wu et al., 2012). Consequently, understanding how this mito-
chondrial ‘‘nutrient wasting’’ process is regulated in all cell types
and in a tissue-specific manner might prove useful for the treat-
ment of conditions associated with excess nutrients.
Cells that should be particularly susceptible to nutrient
supply and demand imbalance are those allowing nutrient
permeability regardless of their energy demand. Such cells
are the nutrient sensors, the regulators and the storage organs:
the beta cells, the hepatocytes, and the adipocytes. In the case
of white adipocytes, high nutrient permeability allows for
storage of nutrients in the form of triacylglicerides. However,
in the nutrient sensors (e.g., beta cells), nutrient oxidation and
ATP/ADP ratio serve as a sensing mechanism and a signal
generator for insulin secretion. This ability of the beta cell to
Cell Metabolism 17, April 2, 2013 ª2013 Elsevier Inc.
control and modulate its mitochondrial bioenergetics according
to nutrient supply is essential to maintain its function in nutrient
stimulated insulin secretion. It might also play a role in main-
taining beta cell viability through the removal of excess
nutrients that if left to accumulate may have a toxic effect
(reviewed in Prentki et al., 2002; reviewed in Muoio and New-
In the ‘‘Relationship between Bioenergetic Efficiency and
Mitochondrial Dynamics’’ section, we will discuss evidence for
the role of mitochondrial dynamics and morphology in regulating
energy efficiency and nutrient wasting.
Effects of Nutrient Excess on Mitochondrial
Bioenergetics in Brown Adipose Tissue, Muscle,
and the Beta Cell
Brown Adipose Tissue. Mitochondria from brown adipose
tissue harbor UCP1, activation of which generates heat through
dissipation of mitochondrial membrane potential and increased
respiratory rates (Aquila et al., 1985; Heaton et al., 1978; Nich-
olls, 1974; Nicholls et al., 1978). UCP1 is used as a specific
marker to detect brown adipocytes within other tissues. The
brown adipocyte represents a model in which a large shift in
bioenergetic efficiency can
hormonal stimulation. Activation of nonshivering thermogenesis
in human brown adipocytes by cold is achieved by the increase
in fatty acid availability to the mitochondria and their oxidation,
which is the result of norepinephrine (NE)- induced lipolysis (re-
viewed in Cannon and Nedergaard, 2004; Ouellet et al., 2012).
In the case of rodents, high fat diet (a form of nutrient excess)
increases brown adipose tissue (BAT) mass. This is mainly
thanks to the increase in brown fat proliferation and differentia-
tion, which result in the increase in UCP1 expression and the
expansion of mitochondrial mass per cell in rodent models
(Himms-Hagen et al., 1981; Rothwell and Stock, 1979). Whether
an increase in the activity of this diet-induced expanded BAT in
rodents contributes to what was defined as diet-induced ther-
mogenesis is controversial (reviewed in Kozak, 2010).
Mitochondrial expansion induced by high-fat-diet in rodent
brown fat shows that when ATP demand is not the main drive
for oxygen consumption (i.e., conditions characterized by
increased uncoupling such as in the activated brown fat),
nutrient excess and increased fuel availability to the mitochon-
dria does not impair bioenergetic function. This lack of toxicity
could be explained by the association between mitochondrial
membrane potential and escape of electrons from the electron
transport chain to generate ROS (Brand et al., 2004). Coupled
respiration normallyoccurs athighervalues ofmembranepoten-
tial as compared to uncoupled respiration, which generates heat
through UCP1 activation or other uncouplers. This means that
uncoupled mitochondria will potentially generate less ROS
when compared to coupled mitochondria under conditions of
nutrient excess. Following the metaphor of the hose, mitochon-
dria from brown fat would have a second valve, constituted by
UCP1, which would allow increasing water flow, while avoiding
high pressure and any damage to the hose. The lack of this
second valve with high capacity in muscle mitochondria might
explain why diets similar to the ones inducing mitochondrial
expansion in brown fat cause mitochondrial oxidative damage
and dysfunction in muscle (decreased citrate synthase activity
and decreased expression of complex IV subunits) (Bonnard
be acutelyinduced through
et al., 2008) (see the next section). Thus, nutrient excess in the
form of high-fat diet can expand mitochondrial capacity in
some tissues, whereas mitochondria from other tissues might
be damaged by the same diet.
Muscle. Current data suggest potential mechanisms by which
nutrient supply and demand imbalance might affect muscle
mitochondrial function. Nutrient excess in the form of long-
term high-fat diet results in the accumulation of toxic levels of
intermediates of fatty acid metabolism. Some of these interme-
diates were shown to be a result of incomplete mitochondrial
and todecreased glucose oxidation (Koves etal.,2008).Further-
of mitochondrial electron transport chain function reported in
skeletal muscle from type-2-diabetic patients (Kelley et al.,
2002). Other studies show that increased ROS generation,
caused by nutrient excess through long-term feeding of a high-
damage to the mitochondria and their dysfunction, the latter
taking place after the onset of insulin resistance (Bonnard
et al., 2008). Thus, excessive ROS production would be a major
contributor to insulin resistance. These mechanisms would
suggest that decreased mitochondrial function is not a regulated
process but rather caused by damaging effects caused by
Other studies suggest that decreased mitochondrial electron
transport chain (ETC) function reported in diabetic muscle might
venting insulin resistance, although sometimes not successfully.
These studies characterized two mouse models of a ‘‘primary’’
reduction in ETC complexes activity, which are muscle-specific
knockouts of the apoptosis-inducing factor (AIF) and the tran-
scription factor A mitochondrial (TFAM), respectively (Pospisilik
et al., 2007; Wredenberg et al., 2006). These knockout mice
showed improved insulin sensitivity (Wredenberg et al., 2006;
Pospisilik et al., 2007) and protection from high fat diet-induced
obesity (Pospisilik et al., 2007). These findings suggest that the
observed decrease in mitochondrial bioenergetic function in
type 2 diabetics could be preventing mitochondrial-mediated
toxicity associated with nutrient excess. This would favor the
hypothesis that inherited or induced transcriptional downregula-
tion of mitochondrial transcripts (Mootha et al., 2003; Patti et al.,
2003, Petersen et al., 2004) is a protective mechanism which
counteracts insulin resistance, rather than a pathogenic mecha-
nism contributing to insulin resistance.
A potential explanation for the beneficial effect of reduced
ETC activity is that reduction in the mass of coupled mitochon-
dria in the muscle exposed to nutrient excess and low ATP
demand might serve as a mechanism for avoiding ROS-
mediated insulin resistance.
Another mechanism that could cope with toxicity associated
with nutrient excess is muscle uncoupled respiration. Increase
of proton conductance can decrease mitochondrial ROS
production and can enhance the removal of toxic intermediates
by completing their oxidation (see the previous section). How-
ever, nutrient-overload-induced uncoupling and its relationship
to ROS production in muscle is still controversial, and the
conclusions are different depending on the study, diets, mouse
models, and even the mitochondrial population analyzed
Cell Metabolism 17, April 2, 2013 ª2013 Elsevier Inc.
(subsarcolemal versus intermyofibrillar mitochondria) (Asami
et al., 2008; Mollica et al., 2006; Almind et al., 2007; Fink et al.,
2007; Nabben et al., 2011a, 2011b).
These inconsistent findings might reflect the inability of oxida-
tive muscle to promote a large shift in bioenergetic efficiency. A
large increase in uncoupling capacity by nutrient excess, as in
brown fat, could severely compromise ATP synthesis and thus
oxidative muscle contractile function and calcium homeostasis.
Furthermore, muscle is a ‘‘nutrient-consuming organ,’’ and it has
a steady supply of nutrients in vivo. In addition to fatty acids,
these include glucose during the fed state, glycogen during the
initial phaseofstarvation,andketone bodiesduringintermediate
starvation. Therefore, it makes physiological sense that high-
caloric nutrients, such as fatty acids, do not by and large
increase uncoupling capacity in oxidative muscle (inducible
proton conductance) as in brown fat. On the other hand, it is of
relevance to study the regulation of the basal proton conduc-
tance or the inherent proton leak in muscle, as this tissue
accounts for the major part of nutrient oxidation and thus for
the overall organism metabolic efficiency. Thus, the study of
mechanisms controlling inherent proton leak in muscle might
reveal mechanisms coping with nutrient excess.
The Beta Cell. The beta cell gauges glucose, free fatty acid,
and amino acid availability in the bloodstream and secretes
insulin accordingly (reviewed in Deeney et al., 2000; reviewed
in Rutter, 2001). This gauging is performed through nutrient
oxidation and mitochondrial respiration. Mechanistically, the
main signal stimulating insulin secretion is increased cytosolic
ATP/ADP ratio, through glucose oxidation and likely increased
mitochondrial ATP synthesis. In addition, various studies show
that byproducts of nutrient oxidation in the mitochondria,
including Malonyl-CoA, ROS, and GTP, serve as mediators of
insulin secretion, also termed as ‘‘secretagogues’’ (Pi et al.,
2007; reviewed in Prentki et al., 1997; Kibbey et al., 2007; re-
viewed in Rutter, 2001). Some amino acids can stimulate insulin
secretion by providing Acetyl-CoA to the Krebs cycle and
increasing mitochondrial ATP synthesis (Floyd et al., 1966; re-
viewed in Poitout and Robertson, 2008). Along with this, there
are also additive effects on insulin secretion by simultaneous
presence of different nutrients. Fatty acids can modulate
glucose-stimulated insulin secretion, through their beta oxida-
tion, through the generation of monoglycerides and acyl-CoA,
or by direct interaction with plasma membrane receptors (re-
viewed in Poitout and Robertson, 2008). Since beta cells import
and metabolize nutrients based on availability, and not on
demand, mechanisms that handle excess nutrient availability
are of particular value.
How do beta cell mitochondria respond to nutrient excess?
Long-term exposureof betacellstohigh levelsof glucose,lipids,
or their combination has deleterious effects on beta cell mito-
chondrial function, physiology, and viability. The observation
that glucose synergizes with free fatty acids in producing the
toxic effects of nutrient excess suggests that the two converge
onto a common product (reviewed in Poitout and Robertson,
2008; reviewed in Prentki et al., 2002; reviewed in Deeney
et al., 2000). The usual suspect would be a situation of reductive
stress characterized by increase in NADH, which, in the absence
of increased ATP demand, generates mitochondrial hyperpolar-
ization and produces excess ROS (see the hose metaphor in the
‘‘How Can Bioenergetic Efficiency Affect Cellular Functionality
and Viability?’’ section).
Perhaps, since the ability to adapt to excess supply has rarely
if ever been selected for, beta cells are designed to be sensitive
to ROS as a mechanism for nutrient sensing. As such, the beta
cells have low antioxidant activity. ROS production mediated
by high nutrients is utilized in the beta cell to couple nutrient
oxidation to insulin secretion independently of changes in
mitochondrial ATP synthesis (Pi et al., 2007). Therefore, insulin
secretion could occur under conditions in which the ATP
demand in the beta cell is low. However, an abnormal situation
of permanent nutrient excess or continuous exposure to fat
(such as type 2 diabetes) would cause mitochondrial damage
or decreased function by sustained overproduction of ROS
combined with reduced antioxidant activity.
Given the importance of ATP production, ROS, and mitochon-
drial-derived coupling factors in insulin secretion, one would
expect that respiration would be very efficiently coupled to
ATP synthesis in beta cells. However, the case is exactly the
opposite. Beta cell mitochondria show higher levels of inherent
proton leak than do mitochondria from other tissues (e.g.,
muscle-derived cells) (Affourtit and Brand, 2006). Although it
might seem counterintuitive, uncoupled respiration allows
limiting ROS-mediated toxicity caused by nutrient excess. This
is consistent with the fact that beta cells require other mecha-
nisms to control ROS production, as they harbor low antioxidant
activity. Thus, mitochondrial uncoupling is one of the few antiox-
idant mechanisms described so far that maintains proportional-
ity between nutrient oxidation and insulin secretion through
ROS production. At the same time, uncoupling should be tightly
regulated in a relatively short period of time, as ATP/ADP ratio is
a signal for insulin secretion, which requires efficient and
coupled ATP synthesis.
We can conclude that mitochondria in the beta cell have some
bioenergetic properties that fall in between mitochondria from
muscle and brown fat, which permit executing their specific
physiological function related to nutrient sensing.
Relationship between Bioenergetic Efficiency and
In this section, we will summarize the changes observed in mito-
ergetic adaptation. This association raises different questions
that are essential to answer in order to understand the relevance
of this association:
(1) What comes first, changes in mitochondrial dynamics or
changes in bioenergetic efficiency? Which factor serves
the other? In this section, we will discuss evidence
showingthat changes in dynamics modulate bioenergetic
efficiency and vice versa. It is likely that the cell type and
the metabolic state are major determinants in this rela-
(2) If bioenergetic adaptation requires changes in mitochon-
drial dynamics, what are the consequences for mitochon-
drial quality control?
Regarding the first question, specific modulation of mitochon-
drial bioenergetics has been shown to cause profound changes
Cell Metabolism 17, April 2, 2013 ª2013 Elsevier Inc.
to mitochondrial dynamics. These changes were to a large
extent, interpreted in the context of quality control activation
(Twig et al., 2008a; reviewed in Twig et al., 2008b). However,
new evidence suggests that changes in mitochondrial structure
mediated by nutrients and their metabolites might represent an
adaptation to the changes in ATP demand and supply.
Summary of Proteins Regulating Mitochondrial
Mitochondrial architecture is determined by motility, fusion, and
fission events. Mitochondrial fusion in mammals is mediated by
mitofusins (Mfn1 and Mfn2, located in the outer mitochondrial
membrane) and optic atrophy gene 1 (Opa1, located in the inner
membrane) (reviewed in Liesa et al., 2009). These three proteins
require GTPase activity to mediate mitochondrial fusion. Proteo-
lytic processing of Opa1 controls its fusion activity but also an
Opa1 fusion-independent role, controlling cristae structure re-
modeling (reviewed in Liesa et al., 2009; Ishihara et al., 2006;
Frezza et al., 2006). On the other hand, mitochondrial fission is
mediated by fission 1 protein (Fis1, located in the outer mito-
chondrial membrane), mitochondrial fission factor (Mff, located
in the outer mitochondrial membrane), and dynamin-related
protein 1 (Drp1, which is mostly cytosolic and translocates to
the outer mitochondrial membrane during fission). Drp1 recruit-
ment to the outer mitochondrial membrane and GTP hydrolysis
are required for Drp1-mediated fission (reviewed in Liesa
et al., 2009). Mff and Fis1 do not harbor GTPase activity, and
different studies show that they mediate fission by recruiting
Drp1 (or other factors) to the mitochondria to a different extent.
MiD49 and MiD51 have been recently described to recruit
Drp1 to the mitochondria, although the role of MiD49- and
MiD51-mediated recruitment in mitochondrial fission is still
under investigation (Palmer et al., 2011; Loso ´n et al., 2013). Of
note, Drp1, Fis1, and Mff also control peroxisomal fission
(Schrader, 2006; Waterham et al., 2007; Gandre-Babbe and
van der Bliek, 2008).
Respiratory Capacity: Effects of Chemical Uncouplers
and Nutrient Excess
The addition of chemical uncouplers (i.e., FCCP or CCCP)
causes complete mitochondrial network fragmentation, Drp1
recruitment to the outer membrane, and OPA1 degradation (Du-
vezin-Caubet et al., 2006; Griparic et al., 2007; Ishihara et al.,
2006; Song et al., 2007; Legros et al., 2002; Cereghetti et al.,
2008). In addition, more-recent studies show that depolarization
by CCCP also triggers the proteasome-dependent degradation
of additional mitochondrial fusion proteins (Mfn1 and Mfn2)
and other outer-membrane proteins. However, this protea-
some-dependent degradation of Mfns requires the overexpres-
sion of the E3-ubiquitin-ligase Parkin (Tanaka et al., 2010; Ziviani
et al., 2010; Chan et al., 2011). These studies demonstrated that
mitochondrial fission is stimulated and fusion is inhibited in de-
and OPA1/Mfn degradation, respectively. This suggests the
possibility that fragmentation is advantageous for a system
working at maximal respiratory capacity or for effective un-
coupled respiration and depolarization.
Depolarization, decreased mitochondrial ATP synthesis effi-
ciency, or inhibition of fusion is not equivalent to mitochondrial
dysfunction. Consistent with this, the use of uncouplers can
mimic physiological conditions of nutrient excess and thus
increase nutrient oxidation and electron transport chain activity,
such as in the activated brown fat or in the beta cell.
Consistent with this idea, studies exposing beta cells to
nutrient excess (Molina et al., 2009) or to conditions that
uncouple mitochondria with a physiological stimulus show
increased respiration and robust fragmentation of the mitochon-
drial network (see Figure 2). Thus, it is likely that fragmentation is
also associated with both maximal respiratory rates and
increased proton conductance.
In this regard, there are some differences between the frag-
mentation observed under FCCP and the fragmentation
observed under a rich-nutrient environment or oxidative stress.
Treatment with uncouplers results in fragmentation and the
generation of doughnut (bagel)-shaped mitochondria (Liu and
Hajno ´czky, 2011). Nutrient-induced fragmentation in the beta
cell is accompanied by increase in mitochondrial diameter to
form ball-shapedinsteadof doughnut(bagel-shaped)
20 mM Glucose
5 mM Glucose
Mitochondrial Network Fragmentation
% cells with
Control Glu 20 Pal0.4
Figure 2. Nutrient Excess Induces Mitochondrial Fragmentation in
the Beta Cell
INS-1 cells treated for 4 hr with different concentrations of glucose and fatty
acids (palmitate conjugated to BSA). The upper panel shows representative
images of INS-1 cells cultured with physiological glucose concentrations
(5 mM glucose) and with high glucose and high fatty acid concentrations
(20 mM glucose + 0.4 mM palmitate BSA at 4:1 ratio) for 4 hr. Mitochondria are
shown in red and were labeled with DsRed targeted to the mitochondria. Cells
exposed to high levels of nutrients (20 mM glucose+ 0.4 mM palmitate) show
fragmentation and the formation of spherical mitochondria (ball shape),
whereas mitochondria with 5 mM glucose appear tubular. The bar graph
shows the percentage of cells with fragmented mitochondria after 4 hr incu-
bation with different concentrations of glucoseand palmitate (in mM). Note the
additive effect of glucose and fatty acids causing fragmentation. See Molina
et al. (2009) for more details.
Cell Metabolism 17, April 2, 2013 ª2013 Elsevier Inc.
mitochondria (Molina et al., 2009) (see Figure 2). The difference
between the two conditions might hint to the potential different
roles of the fragmentation and the increase in diameter. Frag-
mentation might support increased respiration, and the increase
in diameter might support increased inherent proton leak.
Indeed, these different morphologies can be explained by mito-
chondrial membrane potential values. FCCP causes massive
mitochondrial depolarization (see the ‘‘Regulation of Cellular
Bioenergetics by Nutrients’’ section; it can reach 90 mV),
whereas nutrient excess increases mitochondrial membrane
potential (Goehring et al., 2012; Wikstrom et al., 2007; Danial
et al., 2008; Heart et al., 2006). Indeed, oligomycin, which mark-
edly increases membrane potential (up to 220 mV in isolated
mitochondria; see the ‘‘Regulation of Cellular Bioenergetics by
Nutrients’’ section) was shown to cause fragmentation (Legros
et al., 2002). Therefore, the increase in mitochondrial diameter
with high nutrients could be a consequence of the increase in
inherent proton leak associated with high membrane potential
values. On the other hand, FCCP would artificially increase
proton conductance by itself (induced) and would not activate
the inherent proton leak.
The common denominator between uncoupler-induced respi-
ration and basal protonconductance increased by high nutrients
is higher respiration and a decrease in ATP synthesis efficiency
(less ATP per molecule of nutrient oxidized). The most conspic-
uous difference lies in the values of the mitochondrial membrane
The mechanism by which fragmentation may benefit a condi-
tion of maximal respiration under uncoupler is not yet under-
stood. Among other possibilities, fragmentation might represent
a change in cristae structure that allows increased nutrient
import. This would also be consistent with the dual role of
OPA1 in mitochondrial fusion and cristae remodeling (Frezza
et al., 2006). Thus, OPA1 processing/degradation could be one
of the molecular mechanisms behind changes in cristae struc-
ture induced by uncouplers, facilitating nutrient import and/or in-
hibiting mitochondrial ATP synthase dimerization.
Since fragmentation is associated with increased proton
conductance, one might consider the possibility that mitochon-
drial fission proteins, such as Drp1, might facilitate it. At least in
tion might promote proton conductance through the perme-
ability transition pore due to increased recruitment of Bax (Mon-
tessuit et al., 2010). In other systems, Drp1 recruitment to the
outer mitochondrial membrane triggered cristae remodeling
(Germain et al., 2005) and FCCP promoted Drp1 recruitment
(Cereghetti et al., 2008). However, this does not mean that all
forms of fragmentation facilitate proton conductance. Neverthe-
less, it raises the potential role of fragmentation as a first step in
the conversion of a cell into a high proton conductance and high
Mitochondrial Elongation and Bioenergetic Function:
Changes in Dynamics Associated with Situations
Requiring Increased ATP Synthesis Capacity
The opposite condition to nutrient excess, starvation, causes an
acute inhibition of mitochondrial fission, by inhibiting Drp1
recruitment to the mitochondria, and mitochondrial elongation
due to unopposed fusion (Gomes et al., 2011; Rambold et al.,
2011). These studies show that elongation prevented the
removal of mitochondria by the starvation-induced autophagy.
In addition, it causes an increase in mitochondrial cristae
number, which is associated with the dimerization of the ATP
synthase and thus higher ATP synthesis activity (Gomes et al.,
2011). Therefore, starvation would elongate mitochondria in
order to increase ATP synthesis capacity and thus sustain the
ATP demand required during periods of limited nutrient avail-
ability. Furthermore, one could expect mitochondria from oxida-
tive cell types under starvation to be more coupled and to
produce ATP more efficiently, as increased ATP synthesis
capacity alone would deplete the limited amount of nutrients
In a similar manner, mitochondrial elongation occurs during
G1/S phase of the cell cycle, which is characterized by a large
increase in ATP demand to support biogenic processes. Conse-
quently, mitochondrial elongation during G1/S phase could
permit high ATP synthesis rates that can sustain cell duplication
(Mitra et al., 2009). These observations are consistent with respi-
rometry studies, in which it was demonstrated that cells at G1
phase have increased levels of coupled respiration and
membrane potential (Schieke et al., 2008).
Consistent with the notion that mitochondrial elongation
promotes increased mitochondrial ATP synthesis capacity is
the association of elongation with cell senescence (Lee et al.,
2007; Yoon et al., 2006). Senescence involves a decreased
capacity of proliferation, homeostatic imbalance, and thus
decreased capacity of mitochondrial biogenesis. Under this
efficiency serves as an adaptation to reduced mitochondrial
biogenesis. Mitochondrial fusion provides additional benefit, as
it allows for sustaining functional mitochondria with higher
number of mutated mitochondrial DNA (mtDNA) copies per sen-
escent cell by complementation. Indeed, senescent cells show
increased inherent proton leak that might be caused by damage
to the inner mitochondrial membrane (Hutter et al., 2004). This
leak is compensated by increasing absolute values of basal
respiration (compared to nonsenescent cells) and thus maintain-
ing the fraction of respiration coupled to ATP synthesis (Hutter
etal.,2004).Itwillbeinteresting to determine whether senescent
cells can maintain the same degree of mitochondrial ATP
synthesis capacity when mitochondrial fragmentation is induced
and when mitochondrial elongation is prevented.
The senescent cell represents a situation of decreased bioen-
ergetic capacity and decreased work load, while the starved cell
has both capacity and workload increased. These different
needs may explain the difference in the molecular mechanism
under each condition: senescent cells show reduced Fis1 and
Drp1 expression and slightly increased Mfn protein levels,
whereas starved cells show no changes in total proteins levels,
only in Drp1 recruitment to the mitochondria (Mai et al., 2010;
Lee et al., 2007; Yoon et al., 2006; Gomes et al., 2011).
Other acute stresses, such as apoptosis activation (early
stages) and oxidative stress (hydrogen peroxide treatment),
have been shown to induce mitochondrial elongation. These
changes were shown to facilitate ATP synthesis (Jendrach
et al., 2008; Tondera et al., 2009).
The examples reviewed here illustrate that respiration under
uncoupling as found in nutrient excess (or treatment with
uncouplers) is associated with fragmentation and inhibition of
Cell Metabolism 17, April 2, 2013 ª2013 Elsevier Inc.
fusion, whereas the opposite situation, starvation, is associated
with inhibition of fission and increased ATP synthesis (and
potentially with more coupled mitochondria under starvation).
This comparison strengthens the hypothesis that mitochondrial
dynamics plays an active role in changes in mitochondrial
bioenergetic efficiency and capacity (see the summary in
The Link between Bioenergetic Efficiency and
Mitochondrial Architecture: The Pancreatic Beta Cell
As discussed above, the beta cell exquisitely adapts nutrient
oxidation to nutrient availability, thereby coupling the latter to
insulin secretion. This makes the beta cell an attractive model
and cellular bioenergetic efficiency.
changes to mitochondrial architecture and dynamics. Exposure
of beta cell line INS1 to high fat alone or in combination
with glucose leads to mitochondrial fragmentation, which is de-
tected after 4 and 24 hr of addition of high glucose and high fat
(Molina et al., 2009) (Figure 2). Remarkably, the two nutrients
show an additive effect in terms of inducing fragmentation.
This suggests that the two are likely to activate the same frag-
Thus, mitochondrial fragmentation in the beta cell is an early
event that could be directly associated with increased nutrient
oxidation. Mechanistically, the observed nutrient-induced frag-
mentation is mediated by inhibition of mitochondrial fusion (as
shown by decreased sharing in mitochondrial matrix protein
content; see Figure 4). Similar studies revealed a marked
decrease in mitochondrial fusion in primary mouse islets
exposed to high glucose and high fatty acids for 48 hr (Molina
et al., 2009).
Is nutrient-induced fragmentation unique to the beta cell? A
model in which this question can be addressed is the brown
adipocyte. The brown adipocyte allows for hormonal mediated
is an example of a sharp increase in nutrient availability and in
proton conductance, moving from efficient respiration to the
most inefficient respiration in terms of ATP synthesis.
Activated brown fat preferably oxidizes fatty acids, which
would be a similar situation to high fat exposure in the beta
cell. Brown adipocytes go through complete mitochondrial
network fragmentation upon induction of uncoupled respiration,
supporting the observed correlation between the two (unpub-
lished data). Consequently, determination of the importance of
mitochondrial fragmentation to brown fat activation can be
a strong evidence that fragmentation is required to stimulate
and/or enhance uncoupled respiration.
Increase in uncoupled respiration in the beta cell may serve as
a mechanism to remove excess nutrient and set bioenergetic
efficiency to balance beta cell nutrient supply and demand
(see Figure 1). High glucose and particularly high fatty acids
have multiple toxic effects in the beta cell, not only related to
excessive ROS production (Las et al., 2011; reviewed in Poitout
and Robertson, 2008). The increase in uncoupled respiration
could be a mechanism to decrease bioenergetic efficiency in
the beta cell, and thusto getting rid of the excess nutrients within
the beta cell, by oxidizing them to generate heat. In this regard,
Barbara Corkey and Marc Prentki suggested that increased
nutrient oxidation and metabolic cycling in response to nutrient
excess were mechanisms acting to permit beta cell detoxifica-
tion. Consistent with this, increased uncoupled respiration and
ification from the excess of nutrients through their oxidation
without overproduction of ROS. Interestingly, fatty acid excess
is more toxic for beta cells than is high glucose (reviewed in Poit-
out and Robertson, 2008). This might explain why fragmentation
is higher in the presence of fatty acids excess than in the pres-
ence of high glucose in the beta cell. Fatty acids might require
extra detoxification capacity within the beta cell because of their
higher caloric content and their potential cytotoxic intermediates
as a result of their incomplete oxidation (as in muscle) (Koves
et al., 2008).
The difference between fatty acids and glucose in mediating
fragmentation can be explained by the following hypotheses.
Respiration with fatty acids as substrates is associated with
increased mitochondrial proton leak and concomitantly with
lower values of membrane potential, whereas glucose oxidation
(feeding pyruvate to the mitochondria) occurs with relatively
higher membrane potential and thus lower proton leak. There-
fore, one could hypothesize that fatty acids are more efficient
at inducing fragmentation because their oxidation and other
additional effects mediated by fatty acids per se are associated
with a higher proton leak.
Fatty acid oxidation has been associated with higher ROS
production by the electron transport chain. This is in part due
to an additional site for superoxide formation (ETF-Qo, an exclu-
sive site for electron entry into the ETC through fatty acid beta
oxidation) (Seifert et al., 2010). Fragmentation and uncoupling
might therefore be a protective mechanism that prevents oxida-
tive damage. This would suggest that ROS, and not the fatty
and energetic state
Figure 3. The Balance of Energy Supply and Demand Is Associated
with Corresponding Changes to Mitochondrial Architecture and to
Physiological processes associated with increased energy demand and
decreased energy supply, such as acute stress, starvation, and G1/S phase,
are characterized by mitochondrial elongation and by respiration coupled to
ATP synthesis. On the other hand, physiological processes associated with
decreased energy demand and increased supply (high levels of nutrients,
obesity, and type 2 diabetes) are associated with mitochondrial fragmentation
and decreased coupling (associated with heat generation or decreased
Cell Metabolism 17, April 2, 2013 ª2013 Elsevier Inc.
acids or their beta oxidation, are the main activators of fragmen-
tation and uncoupling. In this regard, mitochondrial superoxide
has been shown to activate uncoupled respiration (Echtay
et al., 2002). Therefore, one would expect antioxidants to
decrease fragmentation and proton leak induced by high fatty
An alternative mechanism would be that the fatty acids per se
could be causing fragmentation by directly interfering with the
fusion and fission machinery. Indeed, fatty acids were shown
to activate uncoupled respiration in brown fat through UCP1
(Nicholls and Locke, 1984; Williamson, 1970). The mechanism
for this activation would be more likely related to their chemical
structure, rather than to fatty acid metabolism or an intrinsic pro-
tonophoric activity (Shabalina et al., 2008). In this context,
ture was UCP1-independent (Shabalina et al., 2008). Therefore,
signaling mitochondria fragmentation and consequently un-
coupled respiration in a UCP1-independent manner, in addition
to being the fuels oxidized by mitochondria. Consistent with this,
phospholipase activity in the mitochondria is required for mito-
chondrial fusion mediated by mitofusins (Choi et al., 2006).
This study shows a direct connection between acidic lipids
generated in the mitochondria by phospholipase activity and
fusion (Choi et al., 2006). Thus, fatty acid excess or acidic lipid
moieties could bemodulating fusion byinterfering in these phos-
pholipase-dependent processes or others currently unknown
(Huang et al., 2011). However, this pathway has not been
described in beta cells or brown adipocytes so far.
Ultimately, reductive stressand increasedROS generation are
fragmentation could relieve from reductive stress and ROS
generation by decreasing mitochondrial membrane potential
through cristae remodeling and OPA1 processing (i.e., nutrient
excess). At the same time, mitochondrial fragmentation could
be recruited by mechanisms or physiological processes depola-
rizing the mitochondria, to amplify or enhance the capacity of
The Primary Role of Mitochondrial Dynamics in
Bioenergetic Efficiency: Lessons from Genetic Models
Thus far, we have described the association of mitochondrial
network fragmentation and elongation with bioenergetic effi-
ciency. Examination of genetic models in which alteration of
mitochondrial dynamics proteins is the primary change may
allow us to better understand the cause and effect relationship
between the two.
Effects of Specific Changes in Mitochondrial Dynamics
on Mitochondrial Bioenergetic Efficiency
tected from cell death and shifting it toward fission increased
susceptibility to apoptosis (Frank et al., 2001; Lee et al., 2004).
Consistent with thisobservation, apoptosis has been associated
with complete mitochondrial fragmentation (Frank et al., 2001).
Furthermore, smaller and fragmented mitochondria were found
in skeletal muscle from type-2-diabetic and obese subjects,
chain activity and decreased Mfn2 expression (Bach et al., 2003;
Kelley et al., 2002). Together, these findings led to the initial
impression that mitochondrial fragmentation impairs mitochon-
drial respiratory function and is deleterious to cell viability.
However, these generalizations were found to be inaccurate.
For example, inhibition of mitochondrial fission through Drp1
modulation impairs mitochondrial function. HeLa cells with
and a decrease in both state 3 respiration (maximal ATP
synthesis rates) and state 4 respiration (proton leak or uncou-
in mtDNA copy number in cell culture (Parone et al., 2008),
and complete Drp1 abrogation in mice and humans caused
t=0 min t=55 min
20mM Glucose/0.4mM Palmitate
t=0 min t=55 min
Figure 4. Mitochondrial Fragmentation
Induced by Nutrient Excess in the Beta Cell
Is Caused by Decreased Mitochondrial
Mitochondrial fusion activity was quantified with
mitochondrial matrix-targeted photoactivatable
GFP (mtPAGFP, green). A portion of the mito-
chondrial population within a cell is labeled by
laser photoconversion, and the sharing of the
photoconverted molecules across the mitochon-
drial population through fusion events is moni-
tored. Over 55 min, the majority of mitochondria
acquire photoconverted PA-GFP molecules. As
a result of the dilution of the signal across the cell,
the intensity is diminished. TMRE (red) labeling
was used to visualize the entire mitochondrial
population. Right panels: INS-1 cells expressing
mtPAGFP exposed to nutrient overload (20 mM
glucose + 0.4 mM palmitate-BSA) for 4 hr show
a dramatic reduction in fusion rates, as shown by
the lack of mtPAGFP sharing with other mito-
chondria. Note that due to the lack of fusion, the
mtPAGFP intensity in the labeled mitochondria
remains unchanged. Left panels: INS-1 control
cells (5 mM glucose) present almost all mito-
chondria labeled with mtPAGFP 55 min after
photoactivation. The images are adapted from
Molina et al. (2009) with permission.
Cell Metabolism 17, April 2, 2013 ª2013 Elsevier Inc.
lethality with brain developmental defects and severe neurode-
generation (Ishihara et al., 2009; Wakabayashi et al., 2009;
Waterham et al., 2007). Therefore, Drp1-mediated fission is
important to maintain proper quality control (Twig et al.,
2008a), electron transport chain function, mtDNA integrity, and
cell viability. Drp1 also mediates peroxisomal fission and some
of the physiological changes induced by Drp1 modulation can
be attributed to effects on peroxisome function (Schrader,
2006; Waterham et al., 2007).
Mitochondrial fusion and fission occur sequentially in a
repeating cycle (see Figure 5). The direct implication of this real-
ization is that inhibition of either fusion or fission arrest the cycle.
Indeed, similar bioenergetic defects are observed in cells in
which fusion is inhibited. As an example, skeletal muscle
harboring simultaneous deletions in Mfn1 and Mfn2 expression
(Mfn double knockout) show decreased number of mtDNA
copies, increased mutation and deletion load, and decreased
mitochondrial respiration (Chen et al., 2010). On the other
hand, an ineffective compensatory increase in mitochondrial
mass and complex II activity has been observed in Mfn double-
knockout muscles (Chen et al., 2010). The bioenergetic defect
and the accompanying expansion of mitochondrial mass re-
semble the histopathology of patients harboring mutations in
mtDNA causing MERRF (myoclonic epilepsy with red ragged
fibers). Thus, absence of fusion alters mtDNA homeostasis and
electron transport chain function in a similar manner to the
Figure 5. The Life Cycle of Mitochondria and Its Regulation by Nutrient Availability
(A) The life cycle of the mitochondria. The cycle is characterized by fusion and fission events. Fusion generates a network in which components of the two
mitochondria are mixed and reorganized (1). Fission that follows within minutes splits fused mitochondria into two daughter mitochondria with disparate
membrane potential (2). The daughter with the higher membrane potential is the first to return to the cycle of fusion and fission, while the daughter with more
depolarized membrane potential will remain solitary until its membrane potential recovers (3). If membrane potential remains depolarized, this mitochondrion
will lose its ability to fuse and become part of the preautophagic pool characterized by solitary, depolarized mitochondria (4). With a delay of 1–3 hr, these
mitochondria are eliminated by autophagy (5).
(B) Changes to nutrient availability and energy demand can divert mitochondria from the life cycle and extend their stay in the post fusion state (elongation) or the
post fission state (fragmentation). Elongation of mitochondria is a result of increased fusion or decreased fission activity (top section). This is typical for states of
increased energy efficiency (starvation, acute stress, and senescence). Shortening of mitochondria is a result of decreased fusion activity or increased fission
activity (bottom section). This is typical for states of reduced bioenergetic efficiency (increased uncoupled respiration). Since bioenergetic adaptation to high
energy supply requires the arrest of the mitochondria life cycle, extended exposure to excess nutrient environment is expected to impact quality control,
a condition that will contribute to reduced longevity.
Cell Metabolism 17, April 2, 2013 ª2013 Elsevier Inc.
inhibition of fission. The mechanisms by which lack of fusion or
fission would decrease mtDNA levels are not clear. While mito-
chondrial fusion is the main mechanism proposed to allow
complementation of functional components in mitochondria
harboring mutated mtDNA copies, lack of complementation per
levels (along with a compensatory increase in mass and tran-
scription of nuclear-encoded mitochondrial components).
Mfn2 Deletion in Skeletal Muscle Exacerbates the
Effects of Nutrient Excess on Bioenergetic Efficiency
Some of the first observations that associated fragmentation
with excess nutrients were the decreased size of mitochondria
in muscle from type-2-diabetic and obese humans or mouse
models (Bach et al., 2003; Kelley et al., 2002). An accompanying
decrease in Mfn2 expression provided a potential mechanism to
the reduction in mitochondrial size (Bach et al., 2003; Bach et al.,
2005). Consistent with this, specific deletion of Mfn2 in the
muscle is associated with decreased mitochondrial ATP
synthesis efficiency in permeabilized muscle fibers and with
lower protein levels of different ETC subunits (Sebastia ´n et al.,
2012). This lower efficiency is explained by a mild decrease in
ADP-stimulated respiration and by a mild increase in the leak,
accompanied by an increase in ROS production (Sebastia ´n
et al., 2012). Therefore, as in the beta cell, fragmentation through
inhibition of mitochondrial fusion is associated with increased
proton leak in muscle. In addition, this specific deletion of
Mfn2 in the muscle and mild repression in other tissues is suffi-
cient to impair insulin signaling and exacerbates the deleterious
effects of nutrient excess (high-fat diet) (Sebastia ´n et al., 2012).
These results suggest that Mfn2 expression in the muscle is
required for a proper bioenergetic adaptation to nutrient excess
in the form of high fat diet. Thus, Mfn2 deletion in skeletal muscle
causes a prodiabetic effect that involves increased ROS gener-
ation and oxidative damage.
On the other hand, Mfn2 and other mitochondrial dynamics
components are regulated by pathways activated during condi-
tions of increased energy demand (i.e., exercise and cold expo-
sure) and downregulated in type-2-diabetic patients (nutrient
excess), involving the transcriptional coactivators PGC-1a and
PGC-1b (Cartoni et al., 2005; Liesa et al., 2008; Soriano et al.,
2006; Mootha et al., 2003; Patti et al., 2003). This regulation
also supports, to a certain extent, the link between increased
energy demand and mitochondrial elongation described before
(see the illustration in Figure 3).
In conclusion, maintaining both fusion and fission events is the
key parameter to the homeostasis of the bioenergetic function of
the mitochondrial population within the cell. Specific defects in
mitochondrial dynamics can generate cellular energetic states
similar to conditions with altered nutrient supply and demand
balance, as shown in muscle. In the specific case of long-term
nutrient excess, it is possible that the extension in time of
a short-term protective response to nutrient overload, such as
fragmentation to reduce reductive stress, ROS, and membrane
in the long term (Mouli et al., 2009; Twig et al., 2008a). These
effects on mitochondrial quality control mediated by altered
morphology can potentially explain the abnormal mitochondrial
bioenergetic function and cumulative damage associated with
metabolic diseases or aging.
Effects of Nutrient Availability on Mitochondrial Quality
Changes in mitochondrial dynamics affect quality control and
can therefore influence bioenergetic capacity indirectly (Twig
et al., 2008a). Moreover, recent evidence suggests that nutrients
influence quality control function (Las et al., 2011; Singh et al.,
2009). Therefore, for appropriate consideration of the relation-
ship between mitochondrial dynamics and bioenergetics one
has to consider how both interact with mitochondrial quality
Mechanisms for Mitochondrial Quality Control and Its
Regulation by Bioenergetics, Mitophagy, and
The use of confocal microscopy allows the visualization of single
mitochondria units and specifically to track them over time (Twig
etal.,2008a, 2010; reviewed in Liesaetal., 2009).Avery relevant
observation to our discussion is the finding that mitochondrial
units with a cell are heterogeneous in terms of their bioenergetic
This is reflected by the difference in mitochondrial membrane
neity was modulated by nutrient excess and other metabolic
changes (Wikstrom et al., 2007), which regulate mitochondrial
dynamics (Molina et al., 2009). These data demonstrate that
mitochondrial fusion and fission does not completely equilibrate
the bioenergetic properties of the entire mitochondrial popula-
tion. This stands in contrast to the mitochondrial complementa-
tion theory, which hypothesizes that mitochondrial fusion
homogenizes the entire population, a conclusion drawn from
the observation of shared matrix soluble components. Remark-
ably, however, decreasing mitochondrial fusion rate resulted in
increased heterogeneity illustrating the contribution of mito-
chondrial dynamics to the maintenance of the mitochondrial
bioenergetic function. The paradox could be settled by the
understanding that fusion, fission and autophagy are all con-
nected by one axis (Figure 5).
The quality control axis is centered on the fission event, which
might generate two bioenergetically different mitochondria, one
with a higher membrane potential and one with lower membrane
potential. The single daughter mitochondrion with lower mem-
tial and regain the capacity to reconnect with the network or (2)
remain in the solitary period, depolarized. If membrane potential
is not restored during the solitary period, OPA1 will be degraded.
Thus, the solitary mitochondria will not be able to re-engage with
the network and will be degraded by mitophagy. One can
conclude that fission is an important process isolating a poten-
tially damaging organelle and that selective fusion governs the
fate of the mitochondria to be autophagocytosed (Twig et al.,
2008a). Within this context, long-term inhibition (days) of fission
by Drp1 dominant negative overexpression can reduce the
increase in respiration induced by uncouplers in intact cells
(Twig et al., 2008a). These results should not be interpreted as
evidence for the requirement of fragmentation to achieve
maximal respiratory capacity (see the ‘‘Effects of Specific
Changes in Mitochondrial Dynamics on Mitochondrial Bioener-
getic Efficiency’’ section). The effects on bioenergetics caused
by long-term inhibition of fission can be explained by accumula-
tion of irreversibly damaged mitochondria that cannot be
Cell Metabolism 17, April 2, 2013 ª2013 Elsevier Inc.
segregated (Twig et al., 2008a). This finding is supported by
changes of membrane fluidity in isolated mitochondria from cells
with downregulation of Drp1 (Benard et al., 2007), showing that
the alteration is maintained when mitochondria are taken out of
the cells and mitochondrial dynamics are absent.
Although it is widely accepted that fission events produce
uneven daughters that are selected by autophagy, it might be
appropriate to indicate that this was only shown in the beta
cell and COS7 cells. Similarly, that mitochondrial autophagy is
a housekeeping process that targets spontaneously depolarized
mitochondria was thus far shown only in the beta cells.
Multiple studies have identified additional mechanisms for the
inability of mitochondria in the solitary period to fuse and the
signals that label them to be recognized and removed by the au-
tophagic machinery. The U3-ubiquitin ligase Parkin (mutated in
Parkinson’s disease), through PINK1 serine-threonine kinase
activity, is recruited to depolarized mitochondria to target them
for mitophagy (Narendra et al., 2008; Vives-Bauza et al., 2010;
Ziviani et al., 2010). In addition, Parkin ubiquitinates Mfn,
promoting its degradation by the proteasome system and thus
contributing to fusion inhibition of the solitary depolarized mito-
chondria (Chan et al., 2011; Tanaka et al., 2010; Ziviani et al.,
2010). Therefore, we can define that these solitary and dysfunc-
tional mitochondria, with no fusion capabilities, comprise the
preautophagic pool of mitochondria.
A key component dictating the efficiency of mitochondrial
quality control by fusion, fission, and autophagy is the ability of
a full cycle to be completed and the number of cycles per day
(Mouli et al., 2009). A mathematical model that runs multiple iter-
ations of the cycle predicts that the rate of fusion and fission
cycles determines the capacity of the pathway to restore quality
upon damage. In this context, the effect of nutrient on the rate of
fusion, fission, and the formation of the mitochondrial preauto-
phagic pool may be considered as important in its effect on
autophagy (Las et al., 2011; Singh et al., 2009).
Evidence of Mitochondrial Quality Control, Mitophagy,
and Autophagy Modulation by Nutrients and Their
Relationship to the Energetic State
Nutrient excess leads to the inhibition of fusion, resulting in frag-
mentation and an incomplete cycle of fusion, fission, and auto-
phagy (Molina et al., 2009; Las et al., 2011). In addition, it does
not allow for mitochondrial complementation and thus increases
subcellular mitochondrial heterogeneity (Wikstrom et al., 2007).
Given this lack of selective removal, one could expect that mito-
chondrial mass would decrease, as the population will be mostly
comprised of small and depolarized mitochondria (Figure 5).
Therefore, maintenance of mitochondrial health would only
require stimulation of mitochondrial biogenesis. However,
nutrient excess can impair autophagic flux by inhibiting lyso-
somes, which are required for autophagic degradation (Las
et al., 2011). As a consequence, dysfunctional mitochondria
will accumulate and will affect even mitochondria generated de
novo (by unselective fusion and/or increased ROS production).
These alterations can explain different reports demonstrating
mitochondrial dysfunction in pathologies associated with an
imbalance in nutrient supply and demand.
Turnover requires both fusion events and the segregation of
damaged components by fission, which will not enter again
into the network because fusion is bioenergetically selective
(Figure 5). We suggest that the interaction between mitochon-
drial life cycle, dynamics, and bioenergetics evolved to adapt
to changes in nutrient availability, which are physiologically
comprised of feeding and fasting states. Any prolongation in
the feeding or fasting state requires a bioenergetic adaptation
that will shift the balance of mitochondrial dynamics. A pro-
longed shift will have deleterious effects on mitochondrial health
and quality control. In the case of the fasting state, the shift in
dynamics required for bioenergetic adaptation will homogenize
the mitochondrial population, preventing the segregation, the
formation of the preautophagic pool and the removal of
damaged components by mitophagy. In the fed state and/or
nutrient excess (particularly high fat), fragmentation and high
respiratory rates can lead to damage, in addition to mechanisms
affecting the autophagic machinery downstream of the preauto-
phagic pool of mitochondria. This would cause the accumulation
of dysfunctional units and the increase in ROS generation. In this
context, it is likely that caloric restriction (or proper fed/fasting
cycles) would promote a bioenergetic adaptation and a change
in mitochondrial dynamics, permitting the most efficient mito-
chondrial quality control mechanisms. Thus, this interaction
between bioenergetics adaptation, mitochondrial dynamics,
and quality control could explain some of the beneficial effects
associated with caloric restriction.
We thank Professors Gyorgy Hajnoczky, David Nicholls, Daniel Dagan, Susan
K. Fried, and Barbara E. Corkey for inspiring discussions and insightful
thoughts. M.L. is an Evans Center Fellow and was the recipient of a postdoc-
toral fellowship from Fundacio ´n Ramo ´n Areces. O.S. is funded by NIH grants
RO1 DK35914, R01 DK56690, and R01 DK074778. M.L. is funded by 5 P30
DK046200 BNORC P&F grant. We thank Erga Rivis for her help with the
figures. We thank Drs. Guy Las, Fernanda Cerqueira, Jakob Wikstrom, and Gi-
lad Twig for helpful suggestions.
Affourtit, C., and Brand, M.D. (2006). Stronger control of ATP/ADP by proton
leak in pancreatic beta-cells than skeletal muscle mitochondria. Biochem. J.
Almind, K., Manieri, M., Sivitz, W.I., Cinti, S., and Kahn, C.R. (2007). Ectopic
brown adipose tissue in muscle provides a mechanism for differences in risk
of metabolic syndrome in mice. Proc. Natl. Acad. Sci. USA 104, 2366–2371.
Aquila, H., Link, T.A., and Klingenberg, M. (1985). The uncoupling protein from
brown fat mitochondria is related to the mitochondrial ADP/ATP carrier. Anal-
ysis of sequence homologies and of folding of the protein in the membrane.
EMBO J. 4, 2369–2376.
Asami, D.K., McDonald, R.B., Hagopian, K., Horwitz, B.A., Warman, D., Hsiao,
uncouplingprotein 3(UCP3) onmitochondrial proton leakinmice.Exp. Geron-
tol. 43, 1069–1076.
Ashcroft, F.M., Harrison, D.E., and Ashcroft, S.J. (1984). Glucose induces
closure of single potassium channels in isolated rat pancreatic beta-cells.
Nature 312, 446–448.
Bach, D., Pich, S., Soriano, F.X., Vega, N., Baumgartner, B., Oriola, J.,
Daugaard, J.R., Lloberas, J., Camps, M., Zierath, J.R., et al. (2003). Mitofu-
sin-2 determines mitochondrial network architecture and mitochondrial
metabolism. A novel regulatory mechanism altered in obesity. J. Biol. Chem.
Bach, D., Naon, D., Pich, S., Soriano, F.X., Vega, N., Rieusset, J., Laville, M.,
Guillet, C., Boirie, Y., Wallberg-Henriksson, H., et al. (2005). Expression of
Mfn2, the Charcot-Marie-Tooth neuropathy type 2A gene, in human skeletal
Cell Metabolism 17, April 2, 2013 ª2013 Elsevier Inc.
muscle: effects of type 2 diabetes, obesity, weight loss, and the regulatory role
of tumor necrosis factor alpha and interleukin-6. Diabetes 54, 2685–2693.
Benard, G., Bellance, N., James, D., Parrone, P., Fernandez, H., Letellier, T.,
and Rossignol, R. (2007). Mitochondrial bioenergetics and structural network
organization. J. Cell Sci. 120, 838–848.
Vidal, H., and Rieusset, J. (2008). Mitochondrial dysfunction results from
oxidative stress in the skeletal muscle of diet-induced insulin-resistant mice.
J. Clin. Invest. 118, 789–800.
Brand, M.D., Affourtit, C., Esteves, T.C., Green, K., Lambert, A.J., Miwa, S.,
Pakay, J.L., and Parker, N. (2004). Mitochondrial superoxide: production,
biological effects, and activation of uncoupling proteins. Free Radic. Biol.
Med. 37, 755–767.
Cannon, B., and Nedergaard, J. (2004). Brown adipose tissue: function and
physiological significance. Physiol. Rev. 84, 277–359.
Cartoni, R., Le ´ger, B., Hock, M.B., Praz, M., Crettenand, A., Pich, S., Ziltener,
J.L., Luthi, F., De ´riaz, O., Zorzano, A., et al. (2005). Mitofusins 1/2 and
ERRalpha expression are increased in human skeletal muscle after physical
exercise. J. Physiol. 567, 349–358.
Cereghetti, G.M., Stangherlin, A., Martins de Brito, O., Chang, C.R.,
Blackstone, C., Bernardi, P., and Scorrano, L. (2008). Dephosphorylation by
calcineurin regulates translocation of Drp1 to mitochondria. Proc. Natl.
Acad. Sci. USA 105, 15803–15808.
Chan, N.C., Salazar, A.M., Pham, A.H., Sweredoski, M.J., Kolawa, N.J.,
Graham, R.L., Hess, S., and Chan, D.C. (2011). Broad activation of the ubiqui-
tin-proteasome system by Parkin is critical for mitophagy. Hum. Mol. Genet.
Chance, B., and Williams, G.R. (1955). Respiratory enzymes in oxidative phos-
phorylation. III. The steady state. J. Biol. Chem. 217, 409–427.
Chappell, J.B., and Perry, S.V. (1954). The Respiratory and Adenosinetriphos-
phatase Activitiesof Skeletal-Muscle Mitochondria. Biochem. J. 55, 586–595.
Chen, H., McCaffery, J.M., and Chan, D.C. (2007). Mitochondrial fusion
protects against neurodegeneration in the cerebellum. Cell 130, 548–562.
Chen, H., Vermulst, M., Wang, Y.E., Chomyn, A., Prolla, T.A., McCaffery, J.M.,
and Chan, D.C. (2010). Mitochondrial fusion is required for mtDNA stability in
skeletal muscle and tolerance of mtDNA mutations. Cell 141, 280–289.
Chen, Y., Liu, Y., and Dorn, G.W., 2nd. (2011). Mitochondrial fusion is essential
for organelle function and cardiac homeostasis. Circ. Res. 109, 1327–1331.
Choi, S.Y., Huang, P., Jenkins, G.M., Chan, D.C., Schiller, J., and Frohman,
M.A. (2006). A common lipid links Mfn-mediated mitochondrial fusion and
SNARE-regulated exocytosis. Nat. Cell Biol. 8, 1255–1262.
Datta, S.R., Pitter, K.L., Bird, G.H., Wikstrom, J.D., et al. (2008). Dual role of
proapoptotic BAD in insulin secretion and beta cell survival. Nat Med. 14,
Deeney, J.T., Prentki, M., and Corkey, B.E. (2000). Metabolic control of beta-
cell function. Semin. Cell Dev. Biol. 11, 267–275.
Duvezin-Caubet, S., Jagasia, R., Wagener, J., Hofmann, S., Trifunovic, A.,
Hansson, A., Chomyn, A., Bauer, M.F., Attardi, G., Larsson, N.G., et al.
(2006). Proteolytic processing of OPA1 links mitochondrial dysfunction to
alterations in mitochondrial morphology. J. Biol. Chem. 281, 37972–37979.
Echtay, K.S., Roussel, D., St-Pierre, J., Jekabsons, M.B., Cadenas, S., Stuart,
J.A., Harper, J.A., Roebuck, S.J., Morrison, A., Pickering, S., et al. (2002).
Superoxide activates mitochondrial uncoupling proteins. Nature 415, 96–99.
Mitochondrial proton leak in obesity-resistant and obesity-prone mice. Am. J.
Physiol. Regul. Integr. Comp. Physiol. 293, R1773–R1780.
Floyd, J.C., Jr., Fajans, S.S., Conn, J.W., Knopf, R.F., and Rull, J. (1966). Stim-
ulation of insulin secretion by amino acids. J. Clin. Invest. 45, 1487–1502.
Frank, S., Gaume, B., Bergmann-Leitner, E.S., Leitner, W.W., Robert, E.G.,
Catez, F., Smith, C.L., and Youle, R.J. (2001). The role of dynamin-related
protein 1, a mediator of mitochondrial fission, in apoptosis. Dev. Cell 1,
Frezza, C., Cipolat, S., Martins de Brito, O., Micaroni, M., Beznoussenko, G.V.,
Rudka, T., Bartoli, D., Polishuck, R.S., Danial, N.N., De Strooper, B., and Scor-
rano, L. (2006). OPA1 controls apoptotic cristae remodeling independently
from mitochondrial fusion. Cell 126, 177–189.
Gandre-Babbe, S., and van der Bliek, A.M. (2008). The novel tail-anchored
membrane protein Mff controls mitochondrial and peroxisomal fission in
mammalian cells. Mol. Biol. Cell 19, 2402–2412.
Germain, M., Mathai, J.P., McBride, H.M., and Shore, G.C. (2005). Endo-
plasmic reticulum BIK initiates DRP1-regulated remodelling of mitochondrial
cristae during apoptosis. EMBO J. 24, 1546–1556.
Goehring, I., Gerencser, A.A., Schmidt, S., Brand, M.D., Mulder, H., and Nich-
olls, D.G. (2012). Plasma membrane potential oscillations in insulin secreting
Ins-1 832/13 cells do not require glycolysis and are not initiated by fluctuations
in mitochondrial bioenergetics. J. Biol. Chem. 287, 15706–15717.
Gomes, L.C., Di Benedetto, G., and Scorrano, L. (2011). During autophagy
mitochondria elongate, are spared from degradation and sustain cell viability.
Nat. Cell Biol. 13, 589–598.
Griparic, L., Kanazawa, T., and van der Bliek, A.M. (2007). Regulation of the
mitochondrial dynamin-like protein Opa1 by proteolytic cleavage. J. Cell
Biol. 178, 757–764.
Hafner, R.P., Brown, G.C., and Brand, M.D. (1990). Analysis of the control of
respiration rate, phosphorylation rate, proton leak rate and protonmotive force
in isolated mitochondria using the ‘top-down’ approach of metabolic control
theory. Eur. J. Biochem. 188, 313–319.
Heart, E., Corkey, R.F., Wikstrom, J.D., Shirihai, O.S., and Corkey, B.E. (2006).
Glucose-dependent increase in mitochondrial membrane potential, but not
cytoplasmic calcium, correlates with insulin secretion in single islet cells. Am
J Physiol Endocrinol Metab. 290, E143–E148.
Heaton, G.M., Wagenvoord, R.J., Kemp, A., Jr., and Nicholls, D.G. (1978).
Brown-adipose-tissue mitochondria: photoaffinity labelling of the regulatory
site of energy dissipation. Eur. J. Biochem. 82, 515–521.
Himms-Hagen, J., Triandafillou, J., and Gwilliam, C. (1981). Brown adipose
tissue of cafeteria-fed rats. Am. J. Physiol. 241, E116–E120.
Huang, H., Gao, Q., Peng, X., Choi, S.Y., Sarma, K., Ren, H., Morris, A.J., and
Frohman, M.A. (2011). piRNA-associated germline nuage formation and sper-
matogenesis require MitoPLD profusogenic mitochondrial-surface lipid
signaling. Dev. Cell 20, 376–387.
Hutter, E., Renner, K., Pfister, G., Sto ¨ckl, P., Jansen-Du ¨rr, P., and Gnaiger, E.
(2004). Senescence-associated changes in respiration and oxidative phos-
phorylation in primary human fibroblasts. Biochem. J. 380, 919–928.
Ishihara, N., Fujita, Y., Oka, T., and Mihara, K. (2006). Regulation of mitochon-
drial morphology through proteolytic cleavage of OPA1. EMBO J. 25, 2966–
Ishihara, N.,Nomura,M.,Jofuku, A.,Kato,H., Suzuki, S.O.,Masuda, K.,Otera,
H., Nakanishi, Y.,Nonaka, I.,Goto,Y.,etal. (2009). Mitochondrial fission factor
Drp1 is essential for embryonic development and synapse formation in mice.
Nat. Cell Biol. 11, 958–966.
Jendrach, M., Mai, S., Pohl, S., Vo ¨th, M., and Bereiter-Hahn, J. (2008). Short-
and long-term alterations of mitochondrial morphology, dynamics and mtDNA
after transient oxidative stress. Mitochondrion 8, 293–304.
Kelley, D.E., He, J., Menshikova, E.V., and Ritov, V.B. (2002). Dysfunction of
mitochondria in human skeletal muscle in type 2 diabetes. Diabetes 51,
Kibbey, R.G., Pongratz, R.L., Romanelli, A.J., Wollheim, C.B., Cline, G.W., and
Shulman, G.I. (2007). Mitochondrial GTP regulates glucose-stimulated insulin
secretion. Cell Metab. 5, 253–264.
Koves, T.R., Ussher,J.R., Noland, R.C., Slentz, D., Mosedale, M., Ilkayeva,O.,
Bain, J., Stevens, R., Dyck, J.R., Newgard, C.B., et al. (2008). Mitochondrial
overload and incomplete fatty acid oxidation contribute to skeletal muscle
insulin resistance. Cell Metab. 7, 45–56.
Kozak, L.P. (2010). Brown fat and the myth of diet-induced thermogenesis.
Cell Metab. 11, 263–267.
Cell Metabolism 17, April 2, 2013 ª2013 Elsevier Inc.
Las, G., Serada, S.B., Wikstrom, J.D., Twig, G., and Shirihai, O.S. (2011). Fatty
acids suppress autophagic turnover in b-cells. J. Biol. Chem. 286, 42534–
Lee, Y.J., Jeong, S.Y., Karbowski, M., Smith, C.L., and Youle, R.J. (2004).
Roles of the mammalian mitochondrial fission and fusion mediators Fis1,
Drp1, and Opa1 in apoptosis. Mol. Biol. Cell 15, 5001–5011.
Lee, S., Jeong, S.Y., Lim, W.C., Kim, S., Park, Y.Y., Sun, X., Youle, R.J., and
Cho, H. (2007). Mitochondrial fission and fusion mediators, hFis1 and OPA1,
modulate cellular senescence. J. Biol. Chem. 282, 22977–22983.
Legros, F., Lombe `s, A., Frachon, P., and Rojo, M. (2002). Mitochondrial fusion
in human cells is efficient, requires the inner membrane potential, and is
mediated by mitofusins. Mol. Biol. Cell 13, 4343–4354.
Levine, J.A., Eberhardt, N.L., and Jensen, M.D. (1999). Role of nonexercise
activity thermogenesis in resistance to fat gain in humans. Science 283,
Liesa, M., Borda-d’Agua, B., Medina-Go ´mez, G., Lelliott, C.J., Paz, J.C., Rojo,
M., Palacı ´n, M., Vidal-Puig, A., and Zorzano, A. (2008). Mitochondrial fusion is
increased by the nuclear coactivator PGC-1beta. PLoS ONE 3, e3613.
Liesa, M., Palacı ´n, M., and Zorzano, A. (2009). Mitochondrial dynamics in
mammalian health and disease. Physiol. Rev. 89, 799–845.
Liu, X., and Hajno ´czky, G. (2011). Altered fusion dynamics underlie unique
morphological changes in mitochondria during hypoxia-reoxygenation stress.
Cell Death Differ. 18, 1561–1572.
Loso ´n, O.C., Song, Z., Chen, H., and Chan, D.C. (2013). Fis1, Mff, MiD49, and
MiD51 mediate Drp1 recruitment in mitochondrial fission. Mol. Biol. Cell. 24,
Mai, S., Klinkenberg, M., Auburger, G., Bereiter-Hahn, J., and Jendrach, M.
(2010). Decreased expression of Drp1 and Fis1 mediates mitochondrial
elongation in senescent cells and enhances resistance to oxidative stress
through PINK1. J. Cell Sci. 123, 917–926.
Marcinek, D.J., Schenkman, K.A., Ciesielski, W.A., and Conley, K.E. (2004).
Mitochondrial coupling in vivo in mouse skeletal muscle. Am. J. Physiol. Cell
Physiol. 286, C457–C463.
Mitchell, P. (1961). Coupling of phosphorylation to electron and hydrogen
transfer by a chemi-osmotic type of mechanism. Nature 191, 144–148.
Mitra, K., Wunder, C., Roysam, B., Lin, G., and Lippincott-Schwartz, J. (2009).
A hyperfused mitochondrial state achieved at G1-S regulates cyclin E buildup
and entry into S phase. Proc. Natl. Acad. Sci. USA 106, 11960–11965.
Molina, A.J., Wikstrom, J.D., Stiles, L., Las, G., Mohamed, H., Elorza, A.,
Walzer, G., Twig, G., Katz, S., Corkey, B.E., and Shirihai, O.S. (2009).
apoptosis. Diabetes 58, 2303–2315.
and Iossa, S. (2006). Heterogeneous bioenergetic behaviour of subsarcolem-
mal and intermyofibrillarmitochondria infed and fasted rats. Cell.Mol. LifeSci.
Montessuit, S., Somasekharan, S.P., Terrones, O., Lucken-Ardjomande, S.,
Herzig, S., Schwarzenbacher, R., Manstein, D.J., Bossy-Wetzel, E., Basan ˜ez,
G., Meda, P., and Martinou, J.C. (2010). Membrane remodeling induced by
the dynamin-related protein Drp1 stimulates Bax oligomerization. Cell 142,
Mootha, V.K., Lindgren, C.M., Eriksson, K.F., Subramanian, A., Sihag, S.,
Lehar, J., Puigserver, P., Carlsson, E., Ridderstra ˚le, M., Laurila, E., et al.
(2003). PGC-1alpha-responsive genes involved in oxidative phosphorylation
are coordinately downregulated in human diabetes. Nat. Genet. 34, 267–273.
Mouli, P.K., Twig, G., and Shirihai, O.S. (2009). Frequency and selectivity of
mitochondrial fusion are key to its quality maintenance function. Biophys. J.
Muoio, D.M., and Newgard, C.B. (2006). Obesity-related derangements in
metabolic regulation. Annu. Rev. Biochem. 75, 367–401.
Nabben, M., Hoeks, J., Moonen-Kornips, E., van Beurden, D., Briede ´, J.J.,
Hesselink, M.K., Glatz, J.F., and Schrauwen, P. (2011a). Significance of un-
coupling protein 3 in mitochondrial function upon mid- and long-term dietary
high-fat exposure. FEBS Lett. 585, 4010–4017.
Nabben, M., Shabalina, I.G., Moonen-Kornips, E., van Beurden, D., Cannon,
B., Schrauwen, P., Nedergaard, J., and Hoeks, J. (2011b). Uncoupled
respiration, ROS production, acute lipotoxicity and oxidative damage in
isolated skeletal muscle mitochondria from UCP3-ablated mice. Biochim.
Biophys. Acta 1807, 1095–1105.
Narendra, D.,Tanaka, A.,Suen,D.F.,and Youle,R.J.(2008).Parkinisrecruited
selectively to impaired mitochondria and promotes their autophagy. J. Cell
Biol. 183, 795–803.
Nicholls, D.G. (1974). Hamster brown-adipose-tissue mitochondria. The
control of respiration and the proton electrochemical potential gradient by
possible physiological effectors of the proton conductance of the inner
membrane. Eur. J. Biochem. 49, 573–583.
Nicholls, D.G., and Ferguson, S.J. (2002). Bioenergetics 3 (London: Academic
Nicholls, D.G., and Locke, R.M. (1984). Thermogenic mechanisms in brown
fat. Physiol. Rev. 64, 1–64.
Nicholls, D.G., Bernson, V.S., and Heaton, G.M. (1978). The identification of
the component in the inner membrane of brown adipose tissue mitochondria
responsible for regulating energy dissipation. Experientia Suppl. 32, 89–93.
Ouellet, V., Labbe ´, S.M., Blondin, D.P., Phoenix, S., Gue ´rin, B., Haman, F.,
Turcotte, E.E., Richard, D., and Carpentier, A.C. (2012). Brown adipose tissue
oxidative metabolism contributes to energy expenditure during acute cold
exposure in humans. J. Clin. Invest. 122, 545–552.
Palmer, C.S., Osellame, L.D., Laine, D., Koutsopoulos, O.S., Frazier, A.E., and
Ryan, M.T. (2011). MiD49 and MiD51, new components of the mitochondrial
fission machinery. EMBO Rep. 12, 565–573.
Parker, N.D., Crichton, P.G., Vidal-Puig, A.J., and Brand, M.D. (2009). Uncou-
pling protein-1 (UCP1) contributes to the basal proton conductance of brown
adipose tissue mitochondria. J. Bioenerg. Biomembr. 41, 335–342.
Parone, P.A., Da Cruz, S., Tondera, D., Mattenberger, Y., James, D.I., Maech-
ler, P., Barja, F., and Martinou, J.C. (2008). Preventing mitochondrial fission
impairs mitochondrial function and leads to loss of mitochondrial DNA. PLoS
ONE 3, e3257.
Patti, M.E., Butte, A.J., Crunkhorn, S., Cusi, K., Berria, R., Kashyap, S.,
Miyazaki, Y., Kohane, I., Costello, M., Saccone, R., et al. (2003). Coordinated
reduction of genes of oxidative metabolism in humans with insulin resistance
and diabetes: Potential role of PGC1 and NRF1. Proc. Natl. Acad. Sci. USA
Petersen, K.F., Dufour, S., Befroy, D., Garcia, R., and Shulman, G.I. (2004).
Impaired mitochondrial activity in the insulin-resistant offspring of patients
with type 2 diabetes. N. Engl. J. Med. 350, 664–671.
Pi, J., Bai, Y., Zhang, Q., Wong, V., Floering, L.M., Daniel, K., Reece, J.M.,
Deeney, J.T., Andersen, M.E., Corkey, B.E., and Collins, S. (2007). Reactive
oxygen species as a signal in glucose-stimulated insulin secretion. Diabetes
Poitout, V., and Robertson, R.P. (2008). Glucolipotoxicity: fuel excess and
beta-cell dysfunction. Endocr. Rev. 29, 351–366.
Pospisilik, J.A., Knauf, C., Joza, N., Benit, P., Orthofer, M., Cani, P.D., Ebers-
berger, I., Nakashima, T., Sarao, R., Neely, G., et al. (2007). Targeted deletion
of AIF decreases mitochondrial oxidative phosphorylation and protects from
obesity and diabetes. Cell 131, 476–491.
Prentki, M., Tornheim, K., and Corkey, B.E. (1997). Signal transduction
mechanisms in nutrient-induced insulin secretion. Diabetologia 40(Suppl 2),
Prentki, M., Joly, E., El-Assaad, W., and Roduit, R. (2002). Malonyl-CoA
signaling, lipid partitioning, and glucolipotoxicity: role in beta-cell adaptation
and failure in the etiology of diabetes. Diabetes 51(Suppl 3), S405–S413.
Quiro ´s, P.M., Ramsay, A.J., Sala, D., Ferna ´ndez-Vizarra, E., Rodrı ´guez, F.,
Peinado, J.R., Ferna ´ndez-Garcı ´a, M.S., Vega, J.A., Enrı ´quez, J.A., Zorzano,
A., and Lo ´pez-Otı ´n, C. (2012). Loss of mitochondrial protease OMA1 alters
processing of the GTPase OPA1 and causes obesity and defective thermo-
genesis in mice. EMBO J. 31, 2117–2133.
Rambold, A.S., Kostelecky, B., Elia, N., and Lippincott-Schwartz, J. (2011).
Tubular network formation protects mitochondria from autophagosomal
Cell Metabolism 17, April 2, 2013 ª2013 Elsevier Inc.
degradation during nutrient starvation. Proc. Natl. Acad. Sci.USA 108, 10190–
Rial, E., Poustie, A., and Nicholls, D.G. (1983). Brown-adipose-tissue mito-
chondria: the regulation of the 32000-Mr uncoupling protein by fatty acids
and purine nucleotides. Eur. J. Biochem. 137, 197–203.
Rothwell, N.J., and Stock, M.J. (1979). A role for brown adipose tissue in diet-
induced thermogenesis. Nature 281, 31–35.
Rutter, G.A. (2001). Nutrient-secretion coupling in the pancreatic islet beta-
cell: recent advances. Mol. Aspects Med. 22, 247–284.
Schieke, S.M., McCoy, J.P., Jr., and Finkel, T. (2008). Coordination of mito-
chondrial bioenergetics with G1 phase cell cycle progression. Cell Cycle 7,
Schrader, M. (2006). Shared components of mitochondrial and peroxisomal
division. Biochim. Biophys. Acta 1763, 531–541.
Schutz, Y., Bessard, T., and Je ´quier, E. (1984). Diet-induced thermogenesis
measured over a whole day in obese and nonobese women. Am. J. Clin.
Nutr. 40, 542–552.
Sebastia ´n, D., Herna ´ndez-Alvarez, M.I., Segale ´s, J., Sorianello, E., Mun ˜oz,
J.P., Sala, D., Waget, A., Liesa, M., Paz, J.C., Gopalacharyulu, P., et al.
(2012). Mitofusin 2 (Mfn2) links mitochondrial and endoplasmic reticulum func-
tion with insulin signaling and is essential for normal glucose homeostasis.
Proc. Natl. Acad. Sci. USA 109, 5523–5528.
Seifert, E.L., Estey, C., Xuan, J.Y., and Harper, M.E. (2010). Electron transport
sion during long-chain fatty acid oxidation. J. Biol. Chem. 285, 5748–5758.
Shabalina, I.G., Backlund, E.C., Bar-Tana, J., Cannon, B., and Nedergaard, J.
(2008). Within brown-fat cells, UCP1-mediated fatty acid-induced uncoupling
is independent of fatty acid metabolism. Biochim. Biophys. Acta 1777,
Singh,R.,Kaushik, S.,Wang, Y.,Xiang, Y.,Novak, I., Komatsu, M.,Tanaka,K.,
Cuervo, A.M., and Czaja, M.J. (2009). Autophagy regulates lipid metabolism.
Nature 458, 1131–1135.
Song, Z., Chen, H., Fiket, M., Alexander, C., and Chan, D.C. (2007). OPA1
processing controls mitochondrial fusion and is regulated by mRNA splicing,
membrane potential, and Yme1L. J. Cell Biol. 178, 749–755.
Soriano, F.X., Liesa, M., Bach, D., Chan, D.C., Palacı ´n, M., and Zorzano, A.
(2006). Evidence for a mitochondrial regulatory pathway defined by peroxi-
some proliferator-activated receptor-gamma coactivator-1 alpha, estrogen-
related receptor-alpha, and mitofusin 2. Diabetes 55, 1783–1791.
Tanaka, A., Cleland, M.M., Xu, S., Narendra, D.P., Suen, D.F., Karbowski, M.,
and Youle, R.J. (2010). Proteasome and p97 mediate mitophagyand degrada-
tion of mitofusins induced by Parkin. J. Cell Biol. 191, 1367–1380.
Tondera,D.,Grandemange, S.,Jourdain, A.,Karbowski, M.,Mattenberger,Y.,
is required for stress-induced mitochondrial hyperfusion. EMBO J. 28, 1589–
Twig, G., Elorza, A., Molina, A.J., Mohamed, H., Wikstrom, J.D., Walzer, G.,
Stiles, L., Haigh, S.E., Katz, S., Las, G., et al. (2008a). Fission and selective
fusion govern mitochondrial segregation and elimination by autophagy.
EMBO J. 27, 433–446.
Twig, G.,Hyde, B.,and Shirihai, O.S.(2008b). Mitochondrialfusion, fissionand
autophagy as a quality control axis: the bioenergetic view. Biochim. Biophys.
Acta 1777, 1092–1097.
Twig, G., Liu, X., Liesa, M., Wikstrom, J.D., Molina, A.J., Las, G., Yaniv, G.,
Hajno ´czky, G., and Shirihai, O.S. (2010). Biophysical properties of mitochon-
drial fusion events in pancreatic beta-cells and cardiac cells unravel potential
control mechanisms of its selectivity. Am. J. Physiol. Cell Physiol. 299, C477–
Vives-Bauza, C., Zhou, C., Huang, Y., Cui, M., de Vries, R.L., Kim, J., May, J.,
Tocilescu, M.A., Liu, W., Ko, H.S., et al. (2010). PINK1-dependent recruitment
of Parkin to mitochondria in mitophagy. Proc. Natl. Acad. Sci. USA 107,
Wakabayashi, J., Zhang, Z., Wakabayashi, N., Tamura, Y., Fukaya, M.,
Kensler, T.W., Iijima, M., and Sesaki, H. (2009). The dynamin-related GTPase
Waterham, H.R., Koster, J., van Roermund, C.W., Mooyer, P.A., Wanders,
R.J., and Leonard, J.V. (2007). A lethal defect of mitochondrial and peroxi-
somal fission. N. Engl. J. Med. 356, 1736–1741.
Wikstrom, J.D., Katzman, S.M., Mohamed, H., Twig, G., Graf, S.A., Heart, E.,
mitochondria exhibit membrane potential heterogeneity that can be altered by
stimulatory or toxic fuel levels. Diabetes 56, 2569–2578.
Wikstrom, J.D., Twig, G., and Shirihai, O.S. (2009). What can mitochondrial
heterogeneity tell us about mitochondrial dynamics and autophagy? Int. J.
Biochem. Cell Biol. 41, 1914–1927.
Williamson, J.R. (1970). Control of energy metabolism in hamster brown
adipose tissue. J. Biol. Chem. 245, 2043–2050.
Wu, J., Bostro ¨m, P., Sparks, L.M., Ye, L., Choi, J.H., Giang, A.H., Khandekar,
M., Virtanen, K.A., Nuutila, P., Schaart, G., et al. (2012). Beige adipocytes are
a distinct type of thermogenic fat cell in mouse and human. Cell 150, 366–376.
Wredenberg, A., Freyer, C., Sandstro ¨m, M.E., Katz, A., Wibom, R., Wester-
blad, H., and Larsson, N.G. (2006). Respiratory chain dysfunction in skeletal
muscle does not cause insulin resistance. Biochem. Biophys. Res. Commun.
Yoon, Y.S., Yoon, D.S., Lim, I.K., Yoon, S.H., Chung, H.Y., Rojo, M., Malka, F.,
Jou, M.J., Martinou, J.C., and Yoon, G. (2006). Formation of elongated giant
mitochondria in DFO-induced cellular senescence: involvement of enhanced
fusion process through modulation of Fis1. J. Cell. Physiol. 209, 468–480.
Zhang, Z., Wakabayashi, N., Wakabayashi, J., Tamura, Y., Song, W.J.,
Sereda, S., Clerc, P., Polster, B.M., Aja, S.M., Pletnikov, M.V., et al. (2011).
The dynamin-related GTPase Opa1 is required for glucose-stimulated ATP
production in pancreatic beta cells. Mol. Biol. Cell 22, 2235–2245.
Ziviani, E., Tao, R.N., and Whitworth, A.J. (2010). Drosophila parkin requires
PINK1 for mitochondrial translocation and ubiquitinates mitofusin. Proc.
Natl. Acad. Sci. USA 107, 5018–5023.
Cell Metabolism 17, April 2, 2013 ª2013 Elsevier Inc.