5.5: Uncoupling Electron Transport from ATP Synthesis - Biology

So, that is oxidative phosphorylation. The system is normally highly self-regulated due to impermeability of the inner mitochondrial membrane to H+. Thus there is a direct association between respiration rate and physiological energy need.

Interestingly, there is an exception to this tight coupling of the electron transport chain and formation of ATP. The purpose of brown fat (aka brown adipose tissue), which is most often found in newborn and hibernating mammals, is to generate non-shivering (non-movement-based) heat to keep the animal warm. This is accomplished by uncoupling the electron transport chain from the ATP synthesis. This uncoupling is a hormonally controlled process based on the presence of a mitochondrial proton channel called thermogenin. The hormone norepinephrine increases production of free fatty acids, which open the thermogenin channel. This allows protons to ow from the intermembrane space back into the matrix without having to go through ATP synthase. Because of this, the electron transport chain can keep chugging away, ATP levels do not build up, there is no reduction in respiration rate, and the excess energy not being used in ATP production is released as heat.

In fact, 2,4-dinitrophenol, which is used in a variety of research and industrial applications today, was at one time used as dieting drug (in the 1930’s) because through a different mechanism, it too uncoupled electron transport from ATP synthesis. Its mechanism of action derived from its ability to carry and release protons as it freely diffused through the mitochondrial membrane (since it is a small hydrophobic molecule). As this continues, cells catabolize more and more stores of carbohydrates and fats, which is the reason for the interest by dieters. Unfortunately for some of those dieters, this pharmacological means of uncoupling the electron transport chain from the ATP synthesis had no regulation other than the amount of DNP taken. In cases of overdose, respiration rates could rise dramatically while producing little ATP and a great deal of heat. In fact, overdose illness and death are generally due to the spike in body temperature rather than lowered ATP availability. Unfortunately, there are still some dieters and bodybuilders who self-medicate with DNP despite the dangers.

5.5: Uncoupling Electron Transport from ATP Synthesis - Biology

The electrochemical gradient couples the rate of the electron transport chain to the rate of ATP synthesis. Because electron flow requires proton pumping, electron flow cannot occur faster than protons are used for ATP synthesis (coupled oxidative phosphorylation) or returned to the matrix by a mechanism that short circuits the ATP synthase pore (uncoupling).

7.7.1. Regulation through Coupling

As ATP chemical bond energy is used by energy-requiring reactions, ADP and Pi concentrations increase. The more ADP present to bind to the ATP synthase, the greater will be proton flow through the ATP synthase pore, from the intermembrane space to the matrix.Thus, as ADP levels rise, proton influx increases, and the electrochemical gradient decreases (Fig. 7.2 and 7.3). The proton pumps of the electron transport chain respond with increased proton pumping and electron flow to maintain the electrochemical gradient. The result is increased O2 consumption. The increased oxidation of NADH in the electron transport chain and the increased concentration of ADP stimulate the pathways of fuel oxidation, such as the TCA cycle, to supply more NADH and FADH2 to the electron transport chain. For example, during exercise, we use more ATP for muscle contraction, consume more oxygen, oxidize more fuel (which means burn more calories), and generate more heat from the electron transport chain. If we rest, and the rate of ATP utilization decreases, proton influx decreases, the electrochemical gradient increases, and proton “back-pressure” decreases the rate of the electron transport chain. NADH and FADH2 cannot be oxidized as rapidly in the electron transport chain, and consequently, their build-up inhibits the enzymes that generate them.The electron transport chain has a very high capacity and can respond very rapidly to any increase in ATP utilization.

7.7.2. Uncoupling ATP synthesis from electron transport

When protons leak back into the matrix without going through the ATP synthase pore, they dissipate the electrochemical gradient across the membrane without generating ATP. This phenomenon is called “uncoupling” oxidative phosphorylation. It occurs with chemical compounds, known as uncouplers , and it occurs physiologically with uncoupling proteins that form proton conductance channels through the membrane. Uncoupling of oxidative phosphorylation results in increased oxygen consumption and heat production as electron flow and proton pumping attempt to maintain the electrochemical gradient.

Mitochondria are often described as the "powerhouse" of a cell as it is here that energy is largely released from the oxidation of food. Reducing equivalents generated from beta-oxidation of fatty acids and from the Krebs cycle enter the electron transport chain (also called the respiratory chain). During a series of redox reactions, electrons travel down the chain releasing their energy in controlled steps. These reactions drive the active transport of protons from the mitochondrial matrix , through the inner membrane to the intermembrane space. The respiratory chain consists of five main types of carrier flavins, iron-sulfur centres, quinones, cytochromes (heme proteins) and copper. The two main reducing equivalents entering the respiratory chain are NADH and FADH2. NADH is linked through the NADH-specific dehydrogenase whereas FADH2 is reoxidised within succinate dehydrogenase and a ubiquinone reductase of the fatty acid oxidation pathway. Oxygen is the final acceptor of electrons and with protons, is converted to form water, the end product of aerobic cellular respiration. A proton electrochemical gradient (often called protonmotive force) is established across the inner membrane, with positive charge in the intermembrane space relative to the matrix. Protons driven by the proton-motive force, can enter ATP synthase thus returning to the mitochondrial matrix. ATP synthases use this exergonic flow to form ATP in the matrix, a process called chemiosmotic coupling. A by-product of this process is heat generation.

An antiport, ATP-ADP translocase, preferentially exports ATP from the matrix thereby maintaining a high ADP:ATP ratio in the matrix. The tight coupling of electron flow to ATP synthesis means oxygen consumption is dependent on ADP availability (termed respiratory control). High ADP (low ATP) increases electron flow thereby increasing oxygen consumption and low ADP (high ATP) decreases electron flow and thereby decreases oxygen consumption. There are many inhibitors of mitochondrial ATP synthesis. Most act by either blocking the flow of electrons (eg cyanide, carbon monoxide, rotenone) or uncoupling electron flow from ATP synthesis (eg dinitrophenol). Thermogenin is a natural protein found in brown fat. Newborn babies have a large amount of brown fat and the heat generated by thermogenin is an alternative to ATP synthesis (and thus electron flow only produces heat) and allows the maintenance of body temperature in newborns.

The electron transport chain is located in the inner mitochondrial membrane and comprises some 80 proteins organized in four enzymatic complexes (I-IV). Complex V generates ATP but has no electron transfer activity. In addition to these 5 complexes, there are also two electron shuttle molecules Coenzyme Q (also known as ubiquinone, CoQ) and Cytochrome c (Cytc). These two molecules shuttle electrons between the large complexes in the chain.

How many ATPs are generated by this process? Theoretically, for each glucose molecule, 32 ATPs can be produced. As electrons drop from NADH to oxygen in the chain, the number of protons pumped out and returning through ATP synthase can produce 2.5 ATPs per electron pair. For each pair donated by FADH2, only 1.5 ATPs can be formed. Twelve pairs of electrons are removed from each glucose molecule

10 by NAD+ = 25 ATPs
2 by FADH2 = 3 ATPs.

Making a total of 28 ATPs. However, 2 ATPs are formed during the Krebs' cycle and 2 ATPs formed during glycolysis for each glucose molecule therefore making a total ATP yield of 32 ATPs. In reality, the energy from the respiratory chain is used for other processes (such as active transport of important ions and molecules) so under conditions of normal respiration, the actual ATP yield probably does not reach 32 ATPs.

The reducing equivalents that fuel the electron transport chain, namely NADH and FADH2, are produced by the Krebs cycle (TCA cycle) and the beta-oxidation of fatty acids. At three steps in the Krebs cycle (isocitrate conversion to oxoglutarate oxoglutarate conversion to succinyl-CoA Malate conversion to oxaloacetate), a pair of electrons (2e-) are removed and transferred to NAD+, forming NADH and H+. At a single step, a pair of electrons are removed from succinate, reducing FAD to FADH2. From the beta-oxidation of fatty acids, one step in the process forms NADH and H+ and another step forms FADH2.

Cytoplasmic NADH, generated from glycolysis, has to be oxidized to reform NAD+, essential for glycolysis, otherwise glycolysis would cease to function. There is no carrier that transports NADH directly into the mitochondrial matrix and the inner mitochondrial membrane is impermeable to NADH so the cell uses two shuttle systems to move reducing equivalents into the mitochondrion and regenerate cytosolic NAD+.
The first is the glycerol phosphate shuttle, which uses electrons from cytosolic NADH to produce FADH2 within the inner membrane. These electrons then flow to Coenzyme Q. Complex I is bypassed so only 1.5 ATPs can be formed per NADH via this route. The overall balanced equation, summing all the reactions in this system, is

NADH (cytosol) + H+ (cytosol) + NAD+ (mito.) = NAD+ (cytosol) + NADH (mito.) + H+ (mito.)

The malate-aspartate shuttle uses the oxidation of malate to generate NADH in the mitochondrial matrix. This NADH can then be fed directly to complex I and thus can form 3 ATPs via the respiratory chain. The overall balanced equation is

NADH (cytosol) + H+ (cytosol) + FAD (inner memb.) = NAD+ (cytosol) + FADH2 (inner memb.)

Both of these shuttle systems regenerate cytosolic NAD+.

The entry point for NADH is complex I (NADH dehydrogenase) and the entry point for FADH2 is Coenzyme Q. The input of electrons from fatty acid oxidation via ubiquinone is complicated and not shown in the diagram.

Sperm mitochondrial uncoupling

Mitochondria generate ATP by coupling the activities of two large transport protein complexes on the inner mitochondrial membrane (IMM) – the Electron Transport Chain (ETC) and ATP Synthase. The ETC pumps Hydrogen ions (H + ) out of the mitochondrial matrix to generate an electrochemical H + gradient (ΔΨ) across the IMM. ATP Synthase then allows H + to diffuse back into the mitochondrial matrix and, like a molecular water wheel, uses the energy released by this diffusion to synthesize ATP.

It was originally assumed that H + ions could only return to the matrix through ATP synthase, but it is now well-established that there are other proteins in the IMM which can allow H + to return to the mitochondrial matrix, but do not generate ATP and instead dissipate the released energy as heat. This phenomenon, known as mitochondrial uncoupling , is crucial for mitochondrial function and integrity. In the specialized thermogenic tissues, brown and beige fat, mitochondrial uncoupling helps to maintain core body temperature and control body weight, and is mediated by uncoupling protein 1 (UCP1).

In most other tissues, mitochondria are more rarely uncoupled, and when they are uncoupled, they carry a smaller H + current across the IMM. However, because this “mild” uncoupling occurs in the majority of tissues, it may have a significant impact on thermogenesis, body weight, healthy metabolism and reproduction potential.

Recently, it was demonstrated that mild mitochondrial uncoupling is mediated by Adenosine Nucleotide Transporter (ANT) proteins . These proteins’ primary function is to transport ATP out of the mitochondrial matrix, but when activated by chemical uncouplers, they also allow H + ions to pass through, thereby uncoupling the mitochondria – reducing the IMM potential and slowing the synthesis of ATP. In humans and mice, there are several ANT isoforms, and human ANT4 is exclusively expressed in sperm.

ANT4 has been previously recognized as an excellent contraceptive target, and we aim to activate its uncoupling function, thereby draining sperm of energy and making them unable to find and fertilize an egg. This approach may lead to the creation of unisex contraceptives with fewer side effects than were found in previous attempts to target ANT4.


A procedure is evolved to assess the maximum uncoupling activity of the classical unsubstituted phenolic uncouplers of mitochondrial oxidative phosphorylation (OX PHOS) 2,4-dinitrophenol and 2,6-dinitrophenol. The uncoupler concentrations, C, required for maximum uncoupling efficacy are found to be a strong function of the pH, and a linear relationship of pC with pH is obtained between pH 5 to pH 9. The slopes of the uncoupler concentrations in the aqueous and lipid phases as a function of pH have been estimated. It is shown that the experimental results can be derived from first principles by an enzyme kinetic model for uncoupling that is based on the same equations as formulated for the coupling of ion transport to ATP synthesis in a companion paper after imposition of the special conditions arising from the uncoupling process. The results reveal the catalysis of a reaction that involves both the anionic and protonated forms of the phenolic uncouplers in the vicinity of their binding sites in a non-aqueous region of the cristae membranes of mitochondria. The rate-limiting step in the overall process of uncoupling has been identified based on the uncoupling data. The data cannot be explained by a simple conduction of protons by uncouplers from one bulk aqueous phase to another as postulated by Mitchell's chemiosmotic theory. It is shown that Nath's two-ion theory of energy coupling/uncoupling in ATP synthase is consistent with the results. A molecular mechanism for uncoupling of ATP synthesis by the dinitrophenols is presented and the chief differences between coupling and uncoupling in ATP catalysis are summarized. The pharmacological consequences of our analysis of uncoupling are discussed, with particular reference to the mode of action of the anti-tuberculosis drug bedaquiline that specifically targets the c-subunit of the F1FO-ATP synthase and uncouples respiration from ATP synthesis in Mycobacterium tuberculosis. Hence the work is shown to be important both from the point of view of fundamental biology and is also pregnant with possibilities for practical pharmaceutical applications.

The energetics of electron transport

In discussing the driving forces of electron transport above, we have referred to both the free energy and the redox potential. Before considering the energetics of the respiratory chain in more detail, we will briefly review how exactly these two physical terms relate to one another.

Redox reactions can be compartmentalized to produce a measurable voltage

This slide illustrates the experimental setup for measuring the redox potential of an electron carrier. Left panel: coenzyme Q withdraws electrons from the standard hydrogen electrode, which by definition gives it a positive redox potential (Δ E ). Right: NADH pushes electrons toward the standard electrode, making its Δ E negative.

In the experimental setup, the molecule of interest and a reference solute are contained in two adjacent buffer-filled chambers. Platinum electrodes are immersed in both solutions and connected through a voltmeter ( V ). As electrons are withdrawn from the solute in one chamber and delivered to the other, the voltmeter indicates the direction and magnitude of the potential difference. Protons and other ions can flow across a salt bridge between the chambers so as to preserve electroneutrality. In order to allow the flow of ions but prevent mixing of the chamber contents by convection, this hole is covered with a porous membrane or plugged with agar.

The reference solute commonly used in chemistry is H2 , equilibrated with hydrogen gas at 1 atm above the solution. The corresponding oxidized form, H + , is adjusted to 1 mol/l or pH 0. The immersed platinum electrode not only conducts electrons but also serves as a catalyst for the interconversion between H2 and H + .

The potential of a redox carrier measured against this electrode is defined as its standard redox potential or Δ E 0. For biochemical purposes, the standard electrode solution is buffered at pH 7 rather than pH 0, and the redox potentials measured against this electrode are referred to as Δ E 0 ′ . A pH of 7 is just as arbitrary a reference point as pH 0, but we will stick with it because the textbooks do so, too.

The redox potential (Δ E ) is proportional to the free energy (Δ G )

Δ G (frac< ext>< ext>)
Δ E (frac< ext>< ext>)
Δ G = (frac< ext>< ext> imes frac< ext>< ext>)
ΔG = (Delta E imes frac< ext>< ext>)
Δ G = (- Delta E imes n imes ext) (6.1)

From the previous slide, it is clear that electrons will flow spontaneously from one redox cofactor to another if the corresponding Δ E is positive. We also know that reactions proceed spontaneously if their Δ G is negative. The two parameters are directly related to one another according to equation 6.1 . Either one is therefore sufficient to describe the energetics of the reaction the reason why redox potentials are more commonly used in this context is that they can be measured more directly than Δ G .

In the equation, Δ E is the difference in the redox potentials between two cofactors. The parameter n is the number of electrons transferred in the reaction for example, NADH feeds two electrons at a time into the chain, which means that n equals two for this reaction. In contrast, heme typically accepts and donates single electrons, which means that n =1. The F in the equation is Faraday’s constant, which tells us how many units of charge are carried by one mole electrons (96,500 coulombs/mol). 36 One can think of a cofactor’s redox potential as its affinity for electrons—the higher it is, the more strongly the cofactor will attract electrons. 37

Redox potentials and free energies in the respiratory chain

This slide shows the redox potentials, and the corresponding free energy levels, of some selected electron carriers in the respiratory chain. The lowest potential is found with NAD + , in keeping with its position at the start of the transport chain. The next carrier in sequence, FMN, is part of complex I. It has a slightly higher potential than NADH and is therefore able to accept its electrons. The redox potential increases continuously along the respiratory chain to reach its highest value at oxygen, which therefore has the highest affinity for the electrons and gets to keep them. Reduced oxygen, which recombines with protons to yield water, thus is the end product of respiration.

The iron-sulfur cluster N2, which occupies the lowermost position within complex I as shown in slide 6.4 , has a significantly higher potential than the FMN. This step in potential corresponds to a significant amount of free energy that is released at some point within complex I as the electrons travel through it from FMN toward N2. Complex I uses this energy to expel protons from the mitochondrion, against their concentration gradient. Major steps in potential that drive proton expulsion also occur within complex III and complex IV.

Only minor steps of potential occur in the delivery of electrons from complex I to complex III via coenzyme Q, and between complexes III and IV via cytochrome C. Likewise, with complex II, the potentials of both entry and exit points must fall into the narrow interval between FADH2 and coenzyme Q, which means that very little energy is released as electrons traverse this complex. Such minor steps in redox potential suffice to jog the electrons along, but they are too small to contribute to proton pumping.

Experimental and Computational Evidence for Self-Assembly of Mitochondrial UCP2 in Lipid Bilayers

Uncoupling proteins (UCPs) are members of the mitochondrial carrier family (MCF) that transport protons across the inner mitochondrial membrane, thereby uncoupling electron transport from ATP synthesis. The stoichiometry of UCPs, and the possibility of co-existence of this protein as mono-meric and associated forms in lipid membranes remain an intriguing open question. In the current study, the tertiary structure of UCP2 was analyzed both experimentally and through molecular dynamics (MD) simulations. After recombinant expression of UCP2 in the inner membrane of E. coli, the protein was directly extracted from the bacterial membranes with a non-denaturing detergent and purified both as a pure monomer and as a mixture of monomers, dimers and tetramers. Both protein preparations were re-constituted in egg yolk lipid vesicles. Gel electrophoresis, circular dichroism spectroscopy and fluorescence methods were used to characterize the structure and the proton transport function of protein. UCP2 showed unique stable tetrameric forms in lipid bilayers. MD simulations using membrane lipids and principal component analysis support the experimental results and provided new molecular insights into the nature of noncovalent interactions in oligomeric UCP2. MD simulations indicate that UCP2 tetramers are asymmetric dimers of dimers, in which the interactions between the monomers forming the dimer are stronger than the interactions between the dimers within the tetramer. It is also shown that UCP2 has a specific tendency to form functional tetramers in lipid bilayers, capable of proton transport. The asymmetric nature of the UCP2 tetramer could act as a scaffold for regulating the activity of the monomeric units through cooperative intercommunication between these subunits. Under similar experimental conditions, the structurally comparable ADP/ATP carrier protein did not form tetramers in vesicles, implying that spontaneous tetramerization cannot be generalized to all MCF members.

STATEMENT OF SIGNIFICANCE Self-assembly of membrane proteins plays a significant role in their biological function. In this article, both experimental and computational evidence are provided for spontaneous tetramerization of one of the mitochondrial uncoupling proteins (UCP2) in model lipid membranes. It is also shown that the tetrameric form of UCP2 is capable of proton transport, which leads to regulation of ATP synthesis in mitochondrion. Molecular dynamics simulations confirm the presence of asymmetric UCP2 tetramers as a potential scaffold for regulating the activity of the monomeric units through mutual intercommunication. The outcome of this study provides a solid ground for potential co-existence of monomeric and multimeric functional forms of UCPs that contributes to a deeper molecular insight into their structure and function.

Leaders in Pharmaceutical Business Intelligence (LPBI) Group

The Colors of Respiration and Electron Transport

Reporter & Curator: Larry H. Bernstein, MD, FCAP

Molecular Biology of the Cell. 4th edition

Having considered in general terms how a mitochondrion uses electron
transport to create an electrochemical proton gradient, we need to
examine the mechanisms that underlie this membrane-based energy-conversion process. In doing so, we also accomplish a larger purpose.
As emphasized at the beginning of this chapter, very similar chemi-
osmotic mechanisms are used by mitochondria, chloroplasts, archea,
and bacteria. In fact, these mechanisms underlie the function of nearly
all living organisms— including anaerobes that derive all their energy
from electron transfers between two inorganic molecules. It is therefore
rather humbling for scientists to remind themselves that the existence
of chemiosmosis has been recognized for only about 40 years.

Overview of The Electron Transport Chain

We begin with a look at some of the principles that underlie the electron-transport process, with the aim of explaining how it can pump protons
across a membrane.

Although protons resemble other positive ions such as Na+ and K+
in their movement across membranes, in some respects they are unique.
Hydrogen atoms are by far the most abundant type of atom in living
organisms they are plentiful not only in all carbon-containing
biological molecules, but also in the water molecules that surround
them. The protons in water are highly mobile, flickering through the
hydrogen-bonded network of water molecules by rapidly
dissociating from one water molecule to associate with its neighbor,
as illustrated in Figure 14-20A. Protons are thought to move across a
protein pump embedded in a lipid bilayer in a similar way: they
transfer from one amino acid side chain to another, following a
special channel through the protein.

Protons are also special with respect to electron transport. Whenever
a molecule is reduced by acquiring an electron, the electron (e -) brings
with it a negative charge. In many cases, this charge is rapidly
neutralized by the addition of a proton (H+) from water, so that
the net effect of the reduction is to transfer an entire hydrogen atom,
H+ + e – (Figure 14-20B). Similarly, when a molecule is oxidized,
a hydrogen atom removed from it can be readily dissociated into
its constituent electron and proton—allowing the electron to
be transferred separately to a molecule that accepts electrons,
while the proton is passed to the water. Therefore, in a membrane
in which electrons are being passed along an electron-transport
chain, pumping protons from one side of the membrane to
another can be relatively simple. The electron carrier merely
needs to be arranged in the membrane in a way that causes it to
pick up a proton from one side of the membrane when it accepts
an electron, and to release the proton on the other side of the
membrane as the electron is passed to the next carrier molecule
in the chain (Figure 14-21).

protons pumped across membranes ch14f21

How protons can be pumped across membranes. As an electron
passes along an electron-transport chain embedded in a lipid-bilayer
membrane, it can bind and release a proton at each step.
In this diagram, electron carrier B picks up a proton (H+)
from one (more…)

The Redox Potential Is a Measure of Electron Affinities

In biochemical reactions, any electrons removed from one
molecule are always passed to another, so that whenever one
molecule is oxidized, another is reduced. Like any other chemical r
eaction, the tendency of such oxidation-reduction reactions, or
redox reactions, to proceed spontaneously depends on the free-
energy change (ΔG) for the electron transfer, which in turn
depends on the relative affinities of the two molecules for electrons.

Because electron transfers provide most of the energy for living
things, it is worth spending the time to understand them. Many
readers are already familiar with acids and bases, which donate
and accept protons (see Panel 2-2, pp. 112–113). Acids and bases
exist in conjugate acid-base pairs, in which the acid is readily
converted into the base by the loss of a proton. For example,
acetic acid (CH3COOH) is converted into its conjugate base
(CH3COO-) in the reaction:

In exactly the same way, pairs of compounds such as NADH and
NAD+ are called redox pairs, since NADH is converted to NAD+
by the loss of electrons in the reaction:

NADH is a strong electron donor: because its electrons are held
in a high-energy linkage, the free-energy change for passing its
electrons to many other molecules is favorable (see Figure 14-9).
It is difficult to form a high-energy linkage. Therefore its redox
partner, NAD+, is of necessity a weak electron acceptor.

The tendency to transfer electrons from any redox pair can be
measured experimentally. All that is required is the formation
of an electrical circuit linking a 1:1 (equimolar) mixture of the
redox pair to a second redox pair that has been arbitrarily selected
as a reference standard, so the voltage difference can be measured
between them (Panel 14-1, p. 784). This voltage difference is
defined as the redox potential as defined, electrons move
spontaneously from a redox pair like NADH/NAD+ with a low
redox potential (a low affinity for electrons) to a redox pair like
O2/H2O with a high redox potential (a high affinity for electrons).
Thus, NADH is a good molecule for donating electrons to the
respiratory chain, while O2 is well suited to act as the “sink” for
electrons at the end of the pathway. As explained in Panel 14-1,
the difference in redox potential, ΔE0′, is a direct measure of
the standard free-energy change (ΔG°) for the transfer of an
electron from one molecule to another.

Electron Transfers Release Large Amounts of Energy

As just discussed, those pairs of compounds that have the most negative
redox potentials have the weakest affinity for electrons and therefore
contain carriers with the strongest tendency to donate electrons.
Conversely, those pairs that have the most positive redox potentials
have the strongest affinity for electrons and therefore contain carriers
with the strongest tendency to accept electrons. A 1:1 mixture of NADH
and NAD+ has a redox potential of -320 mV, indicating that NADH has
a strong tendency to donate electrons a 1:1 mixture of H2O and ½O2
has a redox potential of +820 mV, indicating that O2 has a strong
tendency to accept electrons. The difference in redox potential is
1.14 volts (1140 mV), which means that the transfer of each electron
from NADH to O2 under these standard conditions is enormously
favorable, where ΔG° = -26.2 kcal/mole (-52.4 kcal/mole for the two
electrons transferred per NADH molecule see Panel 14-1). If we
compare this free-energy change with that for the formation of the
phosphoanhydride bonds in ATP (ΔG° = -7.3 kcal/mole, see Figure 2-75), we see that more than enough energy is released by the oxidization
of one NADH molecule to synthesize several molecules of ATP from
ADP and Pi.

Phosphate dependence of pyruvate oxidation

Living systems could certainly have evolved enzymes that would
allow NADH to donate electrons directly to O2 to make water in the reaction:

But because of the huge free-energy drop, this reaction would proceed
with almost explosive force and nearly all of the energy would be released
as heat. Cells do perform this reaction, but they make it proceed much
more gradually by passing the high-energy electrons from NADH to
O2 via the many electron carriers in the electron-transport chain.
Since each successive carrier in the chain holds its electrons more
tightly, the highly energetically favorable reaction 2H+ + 2e – + ½O2
→ H2O is made to occur in many small steps. This enables nearly half
of the released energy to be stored, instead of being lost to the
environment as heat.

Spectroscopic Methods Have Been Used to Identify Many Electron
Carriers in the Respiratory Chain

Many of the electron carriers in the respiratory chain absorb visible
light and change color when they are oxidized or reduced. In general,
each has an absorption spectrum and reactivity that are distinct enough
to allow its behavior to be traced spectroscopically, even in crude mixtures.
It was therefore possible to purify these components long before their
exact functions were known. Thus, the cytochromes were discovered
in 1925 as compounds that undergo rapid oxidation and reduction in
living organisms as disparate as bacteria, yeasts, and insects. By observing
cells and tissues with a spectroscope, three types of cytochromes were
identified by their distinctive absorption spectra and designated
cytochromes a, b, and c. This nomenclature has survived, even though
cells are now known to contain several cytochromes of each type and
the classification into types is not functionally important.

The cytochromes constitute a family of colored proteins that are
related by the presence of a bound heme group, whose iron atom
changes from the ferric oxidation state (Fe3+) to the ferrous oxidation
state (Fe2+) whenever it accepts an electron. The heme group consists
of a porphyrin ring with a tightly bound iron atom held by four nitrogen
atoms at the corners of a square (Figure 14-22). A similar porphyrin ring
is responsible for the red color of blood and for the green color of
leaves, being bound to iron in hemoglobin and to magnesium in
chlorophyll, respectively.

The structure of the heme group attached covalently to cytochrome c ch14f22

Figure 14-22. The structure of the heme group attached covalently
to cytochrome c.

The structure of the heme group attached covalently to cytochrome c.
The porphyrin ring is shown in blue. There are five different
cytochromes in the respiratory chain. Because the hemes in different
cytochromes have slightly different structures and (more…)

Iron-sulfur proteins are a second major family of electron carriers. In these
proteins, either two or four iron atoms are bound to an equal number of
sulfur atoms and to cysteine side chains, forming an iron-sulfur center
on the protein (Figure 14-23). There are more iron-sulfur centers than
cytochromes in the respiratory chain. But their spectroscopic detection
requires electron spin resonance (ESR) spectroscopy, and they are less
completely characterized. Like the cytochromes, these centers carry one
electron at a time.

structure of iron sulfur centers ch14f23

Figure 14-23. The structures of two types of iron-sulfur centers.

The structures of two types of iron-sulfur centers. (A) A center of the
2Fe2S type. (B) A center of the 4Fe4S type. Although they contain
multiple iron atoms, each iron-sulfur center can carry only one
electron at a time. There are more than seven different (more…)

The simplest of the electron carriers in the respiratory chain—and
the only one that is not part of a protein—is a small hydrophobic
molecule that is freely mobile in the lipid bilayer known as ubiquinone,
or coenzyme Q. A quinone (Q) can pick up or donate either one or
two electrons upon reduction, it picks up a proton from the medium
along with each electron it carries (Figure 14-24).

quinone electron carriers ch14f24

Figure 14-24. Quinone electron carriers.

Quinone electron carriers. Ubiquinone in the respiratory chain picks
up one H+ from the aqueous environment for every electron it accepts,
and it can carry either one or two electrons as part of a hydrogen atom
(yellow). When reduced ubiquinone donates (more…)

In addition to six different hemes linked to cytochromes, more than
seven iron-sulfur centers, and ubiquinone, there are also two copper
atoms and a flavin serving as electron carriers tightly bound to respiratory-chain proteins in the pathway from NADH to oxygen. This pathway
involves more than 60 different proteins in all.

As one would expect, the electron carriers have higher and higher
affinities for electrons (greater redox potentials) as one moves along
the respiratory chain. The redox potentials have been fine-tuned
during evolution by the binding of each electron carrier in a particular
protein context, which can alter its normal affinity for electrons. However,
because iron-sulfur centers have a relatively low affinity for electrons,
they predominate in the early part of the respiratory chain in contrast,
the cytochromes predominate further down the chain, where a higher
affinity for electrons is required.

The order of the individual electron carriers in the chain was
determined by sophisticated spectroscopic measurements (Figure 14-25),
and many of the proteins were initially isolated and characterized as
individual polypeptides. A major advance in understanding the
respiratory chain, however, was the later realization that most of
the proteins are organized into three large enzyme complexes.

path of electrons ch14f25

Figure 14-25. The general methods used to determine the path of
electrons along an electron-transport chain.

The general methods used to determine the path of electrons along
an electron-transport chain. The extent of oxidation of electron
carriers a, b, c, and d is continuously monitored by following their
distinct spectra, which differ in their oxidized and (more…)

The Respiratory Chain Includes Three Large Enzyme Complexes
Embedded in the Inner Membrane

Membrane proteins are difficult to purify as intact complexes
because they are insoluble in aqueous solutions, and some of
the detergents required to solubilize them can destroy normal
protein-protein interactions. In the early 1960s, however, it
was found that relatively mild ionic detergents, such as deoxycholate,
can solubilize selected components of the inner mitochondrial
membrane in their native form. This permitted the identification
and purification of the three major membrane-bound respiratory
enzyme complexes in the pathway from NADH to oxygen (Figure 14-26).
As we shall see in this section, each of these complexes acts as an
electron-transport-driven H+ pump however, they were
initially characterized in terms of the electron carriers that
they interact with and contain:

mitochondrial oxidative phosphorylation

Figure 14-26. The path of electrons through the three respiratory
enzyme complexes.

The path of electrons through the three respiratory enzyme complexes.
The relative size and shape of each complex are shown. During the
transfer of electrons from NADH to oxygen (red lines), ubiquinone
and cytochrome c serve as mobile carriers that ferry (more…)

The NADH dehydrogenase complex (generally known as complex I)
is the largest of the respiratory enzyme complexes, containing more
than 40 polypeptide chains. It accepts electrons from NADH and
passes them through a flavin and at least seven iron-sulfur centers
to ubiquinone. Ubiquinone then transfers its electrons to a second
respiratory enzyme complex, the cytochrome b-c1 complex.

The cytochrome b-c1 complex contains at least 11 different
polypeptide chains and functions as a dimer. Each monomer
contains three hemes bound to cytochromes and an iron-sulfur
protein. The complex accepts electrons from ubiquinone
and passes them on to cytochrome c, which carries its electron
to the cytochrome oxidase complex.

The cytochrome oxidase complex also functions as a dimer each
monomer contains 13 different polypeptide chains, including two
cytochromes and two copper atoms. The complex accepts one electron
at a time from cytochrome c and passes them four at a time to oxygen.

The cytochromes, iron-sulfur centers, and copper atoms can carry
only one electron at a time. Yet each NADH donates two electrons,
and each O2 molecule must receive four electrons to produce water.
There are several electron-collecting and electron-dispersing points
along the electron-transport chain where these changes in electron
number are accommodated. The most obvious of these is cytochrome

An Iron-Copper Center in Cytochrome Oxidase Catalyzes Efficient
O2 Reduction

Because oxygen has a high affinity for electrons, it releases a
large amount of free energy when it is reduced to form water.
Thus, the evolution of cellular respiration, in which O2 is
converted to water, enabled organisms to harness much more
energy than can be derived from anaerobic metabolism. This
is presumably why all higher organisms respire. The ability of
biological systems to use O2 in this way, however, requires a
very sophisticated chemistry. We can tolerate O2 in the air we
breathe because it has trouble picking up its first electron this
fact allows its initial reaction in cells to be controlled closely by
enzymatic catalysis. But once a molecule of O2 has picked up one
electron to form a superoxide radical (O2 -), it becomes dangerously
reactive and rapidly takes up an additional three electrons wherever
it can find them. The cell can use O2 for respiration only because
cytochrome oxidase holds onto oxygen at a special bimetallic
center, where it remains clamped between a heme-linked iron
atom and a copper atom until it has picked up a total of four electrons.
Only then can the two oxygen atoms of the oxygen molecule be
safely released as two molecules of water (Figure 14-27).

Figure 14-27. The reaction of O2 with electrons in cytochrome oxidase.

The reaction of O2 with electrons in cytochrome oxidase. As indicated,
the iron atom in heme a serves as an electron queuing point this
heme feeds four electrons into an O2 molecule held at the bimetallic
center active site, which is formed by the other (more…)

The cytochrome oxidase reaction is estimated to account for 90%
of the total oxygen uptake in most cells. This protein complex is
therefore crucial for all aerobic life. Cyanide and azide are extremely
toxic because they bind tightly to the cell’s cytochrome oxidase
complexes to stop electron transport, thereby greatly reducing
ATP production.

Although the cytochrome oxidase in mammals contains 13
different protein subunits, most of these seem to have a subsidiary
role, helping to regulate either the activity or the assembly of the
three subunits that form the core of the enzyme. The complete
structure of this large enzyme complex has recently been determined
by x-ray crystallography, as illustrated in Figure 14-28. The atomic
resolution structures, combined with mechanistic studies of the effect
of precisely tailored mutations introduced into the enzyme by genetic
engineering of the yeast and bacterial proteins, are revealing the
detailed mechanisms of this finely tuned protein machine.

Figure 14-28. The molecular structure of cytochrome oxidase.

The molecular structure of cytochrome oxidase. This protein
is a dimer formed from a monomer with 13 different protein
subunits (monomer mass of 204,000 daltons). The three colored
subunits are encoded by the mitochondrial genome, and they
form the functional (more…)

Electron Transfers Are Mediated by Random Collisions in
the Inner Mitochondrial Membrane

The two components that carry electrons between the three
major enzyme complexes of the respiratory chain—ubiquinone
and cytochrome c—diffuse rapidly in the plane of the inner
mitochondrial membrane. The expected rate of random collisions
between these mobile carriers and the more slowly diffusing
enzyme complexes can account for the observed rates of electron
transfer (each complex donates and receives an electron about
once every 5–20 milliseconds). Thus, there is no need to postulate
a structurally ordered chain of electron-transfer proteins in the
lipid bilayer indeed, the three enzyme complexes seem to exist as
independent entities in the plane of the inner membrane, being
present in different ratios in different mitochondria.

The ordered transfer of electrons along the respiratory chain
is due entirely to the specificity of the functional interactions
between the components of the chain: each electron carrier is
able to interact only with the carrier adjacent to it in the sequence
shown in Figure 14-26, with no short circuits.

Electrons move between the molecules that carry them in
biological systems not only by moving along covalent bonds
within a molecule, but also by jumping across a gap as large
as 2 nm. The jumps occur by electron “tunneling,” a quantum-
mechanical property that is critical for the processes we are
discussing. Insulation is needed to prevent short circuits that
would otherwise occur when an electron carrier with a low redox
potential collides with a carrier with a high redox potential. This
insulation seems to be provided by carrying an electron deep
enough inside a protein to prevent its tunneling interactions
with an inappropriate partner.

How the changes in redox potential from one electron carrier
to the next are harnessed to pump protons out of the mitochondrial
matrix is the topic we discuss next.

A Large Drop in Redox Potential Across Each of the Three Respiratory
Enzyme Complexes Provides the Energy for H+ Pumping

We have previously discussed how the redox potential reflects
electron affinities (see p. 783). An outline of the redox potentials
measured along the respiratory chain is shown in Figure 14-29.
These potentials drop in three large steps, one across each major
respiratory complex. The change in redox potential between any
two electron carriers is directly proportional to the free energy
released when an electron transfers between them. Each enzyme
complex acts as an energy-conversion device by harnessing some
of this free-energy change to pump H+ across the inner membrane,
thereby creating an electrochemical proton gradient as electrons
pass through that complex. This conversion can be demonstrated
by purifying each respiratory enzyme complex and incorporating
it separately into liposomes: when an appropriate electron donor
and acceptor are added so that electrons can pass through the complex,
H+ is translocated across the liposome membrane.

Figure 14-29. Redox potential changes along the mitochondrial
electron-transport chain.

Redox potential changes along the mitochondrial electron-transport
chain. The redox potential (designated E′0) increases as electrons
flow down the respiratory chain to oxygen. The standard free-energy
change, ΔG°, for the transfer (more…)

The Mechanism of H+ Pumping Will Soon Be Understood in
Atomic Detail

Some respiratory enzyme complexes pump one H+ per electron
across the inner mitochondrial membrane, whereas others pump
two. The detailed mechanism by which electron transport is coupled
to H+ pumping is different for the three different enzyme complexes.
In the cytochrome b-c1 complex, the quinones clearly have a role.
As mentioned previously, a quinone picks up a H+ from the aqueous
medium along with each electron it carries and liberates it when it
releases the electron (see Figure 14-24). Since ubiquinone is freely
mobile in the lipid bilayer, it could accept electrons near the inside
surface of the membrane and donate them to the cytochrome b-c1
complex near the outside surface, thereby transferring one H+
across the bilayer for every electron transported. Two protons are
pumped per electron in the cytochrome b-c1 complex, however, and
there is good evidence for a so-called Q-cycle, in which ubiquinone
is recycled through the complex in an ordered way that makes this
two-for-one transfer possible. Exactly how this occurs can now be
worked out at the atomic level, because the complete structure of
the cytochrome b-c1 complex has been determined by x-ray
crystallography (Figure 14-30).

Figure 14-30. The atomic structure of cytochrome b-c 1.

The atomic structure of cytochrome b-c 1. This protein is a dimer.
The 240,000-dalton monomer is composed of 11 different protein
molecules in mammals. The three colored proteins form the
functional core of the enzyme: cytochrome b (green), cytochrome (more…)

Allosteric changes in protein conformations driven by electron
transport can also pump H+, just as H+ is pumped when ATP
is hydrolyzed by the ATP synthase running in reverse. For both the
NADH dehydrogenase complex and the cytochrome oxidase complex,
it seems likely that electron transport drives sequential allosteric
changes in protein conformation that cause a portion of the protein
to pump H+ across the mitochondrial inner membrane. A general
mechanism for this type of H+ pumping is presented in Figure 14-31.

Figure 14-31. A general model for H+ pumping.

A general model for H+ pumping. This model for H+ pumping
by a transmembrane protein is based on mechanisms that are
thought to be used by both cytochrome oxidase and the light-driven
procaryotic proton pump, bacteriorhodopsin. The protein
is driven through (more…)

H+ Ionophores Uncouple Electron Transport from ATP Synthesis

Since the 1940s, several substances—such as 2,4-dinitrophenol—
have been known to act as uncoupling agents, uncoupling electron
transport from ATP synthesis. The addition of these low-molecular-weight organic compounds to cells stops ATP synthesis by mitochondria
without blocking their uptake of oxygen. In the presence of an
uncoupling agent, electron transport and H+ pumping continue at
a rapid rate, but no H+ gradient is generated. The explanation for
this effect is both simple and elegant: uncoupling agents are lipid-
soluble weak acids that act as H+ carriers (H+ ionophores), and
they provide a pathway for the flow of H+ across the inner mitochondrial
membrane that bypasses the ATP synthase. As a result of this short-
circuiting, the proton-motive force is dissipated completely, and
ATP can no longer be made.

Respiratory Control Normally Restrains Electron Flow
Through the Chain

When an uncoupler such as dinitrophenol is added to cells,
mitochondria increase their oxygen uptake substantially because
of an increased rate of electron transport. This increase reflects
the existence of respiratory control. The control is thought to
act via a direct inhibitory influence of the electrochemical proton
gradient on the rate of electron transport. When the gradient is
collapsed by an uncoupler, electron transport is free to run unchecked
at the maximal rate. As the gradient increases, electron transport
becomes more difficult, and the process slows. Moreover, if an
artificially large electrochemical proton gradient is experimentally
created across the inner membrane, normal electron transport
stops completely, and a reverse electron flow can be detected in
some sections of the respiratory chain. This observation suggests
that respiratory control reflects a simple balance between the
free-energy change for electron-transport-linked proton pumping
and the free-energy change for electron transport—that is, the
magnitude of the electrochemical proton gradient affects both
the rate and the direction of electron transport, just as it affects
the directionality of the ATP synthase (see Figure 14-19).

Respiratory control is just one part of an elaborate interlocking
system of feedback controls that coordinate the rates of glycolysis,
fatty acid breakdown, the citric acid cycle, and electron transport.
The rates of all of these processes are adjusted to the ATP:ADP ratio,
increasing whenever an increased utilization of ATP causes the ratio
to fall. The ATP synthase in the inner mitochondrial membrane,
for example, works faster as the concentrations of its substrates
ADP and Pi increase. As it speeds up, the enzyme lets more H+ flow
into the matrix and thereby dissipates the electrochemical proton
gradient more rapidly. The falling gradient, in turn, enhances the
rate of electron transport.

Similar controls, including feedback inhibition of several key enzymes
by ATP, act to adjust the rates of NADH production to the rate of
NADH utilization by the respiratory chain, and so on. As a result of
these many control mechanisms, the body oxidizes fats and sugars
5–10 times more rapidly during a period of strenuous exercise than
during a period of rest.

Natural Uncouplers Convert the Mitochondria in Brown Fat into
Heat-generating Machines

In some specialized fat cells, mitochondrial respiration is normally
uncoupled from ATP synthesis. In these cells, known as brown fat
cells, most of the energy of oxidation is dissipated as heat rather
than being converted into ATP. The inner membranes of the large
mitochondria in these cells contain a special transport protein that
allows protons to move down their electrochemical gradient, by-
passing ATP synthase. As a result, the cells oxidize their fat stores
at a rapid rate and produce more heat than ATP. Tissues containing
brown fat serve as “heating pads,” helping to revive hibernating animals
and to protect sensitive areas of newborn human babies from the cold.

Bacteria Also Exploit Chemiosmotic Mechanisms to Harness Energy

Bacteria use enormously diverse energy sources. Some, like animal
cells, are aerobic they synthesize ATP from sugars they oxidize to
CO2 and H2O by glycolysis, the citric acid cycle, and a respiratory
chain in their plasma membrane that is similar to the one in the
inner mitochondrial membrane. Others are strict anaerobes, deriving
their energy either from glycolysis alone (by fermentation) or from an
electron-transport chain that employs a molecule other than oxygen
as the final electron acceptor. The alternative electron acceptor can
be a nitrogen compound (nitrate or nitrite), a sulfur compound
(sulfate or sulfite), or a carbon compound (fumarate or carbonate),
for example. The electrons are transferred to these acceptors by a
series of electron carriers in the plasma membrane that are comparable
to those in mitochondrial respiratory chains.

Despite this diversity, the plasma membrane of the vast majority of
bacteria contains an ATP synthase that is very similar to the one in
mitochondria. In bacteria that use an electron-transport chain to
harvest energy, the electron-transport pumps H+ out of the cell and
thereby establishes a proton-motive force across the plasma membrane
that drives the ATP synthase to make ATP. In other bacteria, the
ATP synthase works in reverse, using the ATP produced by glycolysis
to pump H+ and establish a proton gradient across the plasma
membrane. The ATP used for this process is generated by
fermentation processes (discussed in Chapter 2).

Thus, most bacteria, including the strict anaerobes, maintain a proton
gradient across their plasma membrane. It can be harnessed to drive
a flagellar motor, and it is used to pump Na+ out of the bacterium via
a Na+-H+ antiporter that takes the place of the Na+-K+ pump of
eucaryotic cells. This gradient is also used for the active inward transport
of nutrients, such as most amino acids and many sugars: each nutrient is
dragged into the cell along with one or more H+ through a specific symporter
(Figure 14-32). In animal cells, by contrast, most inward transport across
the plasma membrane is driven by the Na+ gradient that is established by the
Na+-K+ pump.

Figure 14-32. The importance of H+-driven transport in bacteria.

The importance of H+-driven transport in bacteria. A proton-motive force
generated across the plasma membrane pumps nutrients into the cell and
expels Na+. (A) In an aerobic bacterium, an electrochemical proton gradient
across the plasma membrane is produced (more…)

Some unusual bacteria have adapted to live in a very alkaline
environment and yet must maintain their cytoplasm at a physiological
pH. For these cells, any attempt to generate an electrochemical H+
gradient would be opposed by a large H+ concentration gradient in
the wrong direction (H+ higher inside than outside). Presumably for
this reason, some of these bacteria substitute Na+ for H+ in all of their
chemiosmotic mechanisms. The respiratory chain pumps Na+ out of
the cell, the transport systems and flagellar motor are driven by an
inward flux of Na+, and a Na+-driven ATP synthase synthesizes
ATP. The existence of such bacteria demonstrates that the principle
of chemiosmosis is more fundamental than the proton-motive force
on which it is normally based.

The respiratory chain in the inner mitochondrial membrane contains
three respiratory enzyme complexes through which electrons pass on
their way from NADH to O2.

Each of these can be purified, inserted into synthetic lipid vesicles,
and then shown to pump H+ when electrons are transported through it.
In the intact membrane, the mobile electron carriers ubiquinone and
cytochrome c complete the electron-transport chain by shuttling between
the enzyme complexes. The path of electron flow is NADH → NADH
dehydrogenase complex → ubiquinone → cytochrome b-c1 complex →
cytochrome c → cytochrome oxidase complex → molecular oxygen (O2).

The respiratory enzyme complexes couple the energetically favorable
transport of electrons to the pumping of H+ out of the matrix. The
resulting electrochemical proton gradient is harnessed to make ATP
by another transmembrane protein complex, ATP synthase, through
which H+ flows back into the matrix. The ATP synthase is a reversible
coupling device that normally converts a backflow of H+ into ATP
phosphate bond energy by catalyzing the reaction ADP + Pi → ATP,
but it can also work in the opposite direction and hydrolyze ATP to
pump H+ if the electrochemical proton gradient is sufficiently reduced.
Its universal presence in mitochondria, chloroplasts, and procaryotes
testifies to the central importance of chemiosmotic mechanisms in cells.

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Copyright © 2002, Bruce Alberts, Alexander Johnson, Julian Lewis,
Martin Raff, Keith Roberts, and Peter Walter Copyright © 1983, 1989,
1994, Bruce Alberts, Dennis Bray, Julian Lewis, Martin Raff, Keith
Roberts, and James D. Watson .


Cinti, S.Obesity, Type 2 Diabetes and the Adipose Organ: A Pictorial Atlas from Research to Clinical Applications 1st edn (Springer, 2017).

Wu, J., Cohen, P. & Spiegelman, B. M. Adaptive thermogenesis in adipocytes: is beige the new brown? Genes Dev. 27, 234–250 (2013).

Harms, M. & Seale, P. Brown and beige fat: development, function and therapeutic potential. Nat. Med. 19, 1252–1263 (2013).

Lidell, M. E. et al. Evidence for two types of brown adipose tissue in humans. Nat. Med. 19, 631–634 (2013).

Wu, J. et al. Beige adipocytes are a distinct type of thermogenic fat cell in mouse and human. Cell 150, 366–376 (2012).

Shinoda, K. et al. Genetic and functional characterization of clonally derived adult human brown adipocytes. Nat. Med. 21, 389–394 (2015).

Cypess, A. M. et al. Anatomical localization, gene expression profiling and functional characterization of adult human neck brown fat. Nat. Med. 19, 635–639 (2013).

Ikeda, K., Maretich, P. & Kajimura, S. The common and distinct features of brown and beige adipocytes. Trends Endocrinol. Metab. 29, 191–200 (2018).

Lepper, C. & Fan, C. M. Inducible lineage tracing of Pax7-descendant cells reveals embryonic origin of adult satellite cells. Genesis 48, 424–436 (2010).

Seale, P. et al. PRDM16 controls a brown fat/skeletal muscle switch. Nature 454, 961–967 (2008).

Atit, R. et al. β-Catenin activation is necessary and sufficient to specify the dorsal dermal fate in the mouse. Dev. Biol. 296, 164–176 (2006).

Sanchez-Gurmaches, J. & Guertin, D. A. Adipocytes arise from multiple lineages that are heterogeneously and dynamically distributed. Nat. Commun. 5, 4099 (2014).

Sebo, Z. L., Jeffery, E., Holtrup, B. & Rodeheffer, M. S. A mesodermal fate map for adipose tissue. Development 145, dev166801 (2018).

Wang, W. et al. Ebf2 is a selective marker of brown and beige adipogenic precursor cells. Proc. Natl Acad. Sci. USA 111, 14466–14471 (2014).

Zhang, L. et al. Generation of functional brown adipocytes from human pluripotent stem cells via progression through a paraxial mesoderm state. Cell Stem Cell 27, 784–797.e11 (2020). This study generates human brown adipocytes from pluripotent stem cells by a serum-free directed differentiation strategy.

Xue, B. et al. Genetic variability affects the development of brown adipocytes in white fat but not in interscapular brown fat. J. Lipid Res. 48, 41–51 (2007).

Lee, Y. H., Petkova, A. P., Mottillo, E. P. & Granneman, J. G. In vivo identification of bipotential adipocyte progenitors recruited by β-adrenoceptor activation and high-fat feeding. Cell Metab. 15, 480–491 (2012).

Berry, D. C., Jiang, Y. & Graff, J. M. Mouse strains to study cold-inducible beige progenitors and beige adipocyte formation and function. Nat. Commun. 7, 10184 (2016).

Liu, W. et al. A heterogeneous lineage origin underlies the phenotypic and molecular differences of white and beige adipocytes. J. Cell Sci. 126, 3527–3532 (2013).

Oguri, Y. et al. CD81 controls beige fat progenitor cell growth and energy balance via FAK signaling. Cell 182, 563–577.e20 (2020).

Long, J. Z. et al. A smooth muscle-like origin for beige adipocytes. Cell Metab. 19, 810–820 (2014).

Vishvanath, L. et al. Pdgfrβ + mural preadipocytes contribute to adipocyte hyperplasia induced by high-fat-diet feeding and prolonged cold exposure in adult mice. Cell Metab. 23, 350–359 (2016).

Schulz, T. J. et al. Identification of inducible brown adipocyte progenitors residing in skeletal muscle and white fat. Proc. Natl Acad. Sci. USA 108, 143–148 (2011).

Rodeheffer, M. S., Birsoy, K. & Friedman, J. M. Identification of white adipocyte progenitor cells in vivo. Cell 135, 240–249 (2008).

Berry, R. & Rodeheffer, M. S. Characterization of the adipocyte cellular lineage in vivo. Nat. Cell Biol. 15, 302–308 (2013).

Cattaneo, P. et al. Parallel lineage-tracing studies establish fibroblasts as the prevailing in vivo adipocyte progenitor. Cell Rep. 30, 571–582.e2 (2020).

Finlin, B. S. et al. The β3-adrenergic receptor agonist mirabegron improves glucose homeostasis in obese humans. J. Clin. Invest. 130, 2319–2331 (2020). This study reports that chronic activation of the β3-AR by mirabegron improves insulin sensitivity and activates beige fat in humans with obesity.

Finlin, B. S. et al. Human adipose beiging in response to cold and mirabegron. JCI Insight 3, e121510 (2018).

Min, S. Y. et al. Human ‘brite/beige’ adipocytes develop from capillary networks, and their implantation improves metabolic homeostasis in mice. Nat. Med. 22, 312–318 (2016).

Raajendiran, A. et al. Identification of metabolically distinct adipocyte progenitor cells in human adipose tissues. Cell Rep. 27, 1528–1540.e7 (2019).

Singh, A. M. et al. Human beige adipocytes for drug discovery and cell therapy in metabolic diseases. Nat. Commun. 11, 2758 (2020).

Wang, Q. A., Tao, C., Gupta, R. K. & Scherer, P. E. Tracking adipogenesis during white adipose tissue development, expansion and regeneration. Nat. Med. 19, 1338–1344 (2013).

Himms-Hagen, J. et al. Multilocular fat cells in WAT of CL-316243-treated rats derive directly from white adipocytes. Am. J. Physiol. Cell Physiol. 279, C670–C681 (2000).

Barbatelli, G. et al. The emergence of cold-induced brown adipocytes in mouse white fat depots is determined predominantly by white to brown adipocyte transdifferentiation. Am. J. Physiol. 298, E1244–E1253 (2010).

Shao, M. et al. Cellular origins of beige fat cells revisited. Diabetes 68, 1874–1885 (2019). This study reports the quantitative contribution of beige adipocyte biogenesis via de novo differentiation versus reinstallation of existing adipocytes in vivo.

Lee, Y. H., Petkova, A. P., Konkar, A. A. & Granneman, J. G. Cellular origins of cold-induced brown adipocytes in adult mice. FASEB J. 29, 286–299 (2015).

Tajima, K. et al. Mitochondrial lipoylation integrates age-associated decline in brown fat thermogenesis. Nat. Metab. 1, 886–898 (2019).

Berry, D. C. et al. Cellular aging contributes to failure of cold-induced beige adipocyte formation in old mice and humans. Cell Metab. 25, 481 (2017).

Rosenwald, M., Perdikari, A., Rulicke, T. & Wolfrum, C. Bi-directional interconversion of brite and white adipocytes. Nat. Cell Biol. 15, 659–667 (2013).

Altshuler-Keylin, S. et al. Beige adipocyte maintenance is regulated by autophagy-induced mitochondrial clearance. Cell Metab. 24, 402–419 (2016).

Lu, X. et al. Mitophagy controls beige adipocyte maintenance through a Parkin-dependent and UCP1-independent mechanism. Sci. Signal. 11, eaap8526 (2018).

Roh, H. C. et al. Warming induces significant reprogramming of beige, but not brown, adipocyte cellular identity. Cell Metab. 27, 1121–1137.e5 (2018).

Gnad, T. et al. Adenosine activates brown adipose tissue and recruits beige adipocytes via A2A receptors. Nature 516, 395–399 (2014).

Bordicchia, M. et al. Cardiac natriuretic peptides act via p38 MAPK to induce the brown fat thermogenic program in mouse and human adipocytes. J. Clin. Invest. 122, 1022–1036 (2012).

Fisher, F. M. et al. FGF21 regulates PGC-1α and browning of white adipose tissues in adaptive thermogenesis. Genes Dev. 26, 271–281 (2012).

Ohno, H., Shinoda, K., Spiegelman, B. M. & Kajimura, S. PPARγ agonists induce a white-to-brown fat conversion through stabilization of PRDM16 protein. Cell Metab. 15, 395–404 (2012).

Inagaki, T., Sakai, J. & Kajimura, S. Transcriptional and epigenetic control of brown and beige adipose cell fate and function. Nat. Rev. Mol. Cell Biol. 17, 480–495 (2016).

Sidossis, L. & Kajimura, S. Brown and beige fat in humans: thermogenic adipocytes that control energy and glucose homeostasis. J. Clin. Invest. 125, 478–486 (2015).

Sun, W. et al. Cold-induced epigenetic programming of the sperm enhances brown adipose tissue activity in the offspring. Nat. Med. 24, 1372–1383 (2018).

Jiang, Y., Berry, D. C. & Graff, J. M. Distinct cellular and molecular mechanisms for β3 adrenergic receptor-induced beige adipocyte formation. eLife 6, e30329 (2017).

Bronnikov, G., Houstek, J. & Nedergaard, J. β-Adrenergic, cAMP-mediated stimulation of proliferation of brown fat cells in primary culture. Mediation via β1 but not via β3 adrenoceptors. J. Biol. Chem. 267, 2006–2013 (1992).

McQueen, A. E. et al. The C-terminal fibrinogen-like domain of angiopoietin-like 4 stimulates adipose tissue lipolysis and promotes energy expenditure. J. Biol. Chem. 292, 16122–16134 (2017).

Goh, Y. Y. et al. Angiopoietin-like 4 interacts with integrins β1 and β5 to modulate keratinocyte migration. Am. J. Pathol. 177, 2791–2803 (2010).

Zhu, Y. et al. Connexin 43 mediates white adipose tissue beiging by facilitating the propagation of sympathetic neuronal signals. Cell Metab. 24, 420–433 (2016). This study identifies the role of the gap junction in beige fat biogenesis via propagation of the sympathetically derived cAMP signal to neighbouring adipocytes.

Chen, Y. et al. Thermal stress induces glycolytic beige fat formation via a myogenic state. Nature 565, 180–185 (2019).

Jun, H. et al. Adrenergic-independent signaling via CHRNA2 regulates beige fat activation. Dev. Cell 54, 106–116.e5 (2020).

Wu, Y., Kinnebrew, M. A., Kutyavin, V. I. & Chawla, A. Distinct signaling and transcriptional pathways regulate peri-weaning development and cold-induced recruitment of beige adipocytes. Proc. Natl Acad. Sci. USA 117, 6883–6889 (2020).

Song, A. et al. Low- and high-thermogenic brown adipocyte subpopulations coexist in murine adipose tissue. J. Clin. Invest. 130, 247–257 (2019).

Lee, K. Y. et al. Developmental and functional heterogeneity of white adipocytes within a single fat depot. EMBO J. 38, e99291 (2019).

Min, S. Y. et al. Diverse repertoire of human adipocyte subtypes develops from transcriptionally distinct mesenchymal progenitor cells. Proc. Natl Acad. Sci. USA 116, 17970–17979 (2019). This study reports diverse adipocyte progenitors in human adipose tissue that give rise to beige adipocytes.

Xue, R. et al. Clonal analyses and gene profiling identify genetic biomarkers of the thermogenic potential of human brown and white preadipocytes. Nat. Med. 21, 760–768 (2015).

Sun, W. et al. Single-nucleus RNA-seq reveals a new type of brown adipocyte regulating thermogenesis. Nature 587, 98–102 (2020). This study employs single-nucleus RNA-sequencing to characterize adipocyte heterogeneity in mice and humans, and identifies a subpopulation of adipocytes that uses acetate to regulate the thermogenic capacity of neighbouring adipocytes.

Schwalie, P. C. et al. A stromal cell population that inhibits adipogenesis in mammalian fat depots. Nature 559, 103–108 (2018). This study, by single-cell RNA-sequencing analysis, identifies distinct subpopulations of adipose precursor cells, including adipogenesis-regulatory cells, in mouse adipose tissue.

Hepler, C. et al. Identification of functionally distinct fibro-inflammatory and adipogenic stromal subpopulations in visceral adipose tissue of adult mice. eLife 7, e39636 (2018). This study reveals the functional heterogeneity of visceral WAT perivascular cells and identifies fibro-inflammatory progenitors.

Merrick, D. et al. Identification of a mesenchymal progenitor cell hierarchy in adipose tissue. Science 364, eaav2501 (2019). This study employs single-cell RNA sequencing to identify mesenchymal progenitor cells that give rise to adipocytes in mice and humans.

Seale, P. et al. Transcriptional control of brown fat determination by PRDM16. Cell Metab. 6, 38–54 (2007).

Kajimura, S. et al. Regulation of the brown and white fat gene programs through a PRDM16/CtBP transcriptional complex. Genes Dev. 22, 1397–1409 (2008).

Seale, P. et al. Prdm16 determines the thermogenic program of subcutaneous white adipose tissue in mice. J. Clin. Invest. 121, 96–105 (2011).

Cohen, P. et al. Ablation of PRDM16 and beige adipose causes metabolic dysfunction and a subcutaneous to visceral fat switch. Cell 156, 304–316 (2014).

Ohno, H., Shinoda, K., Ohyama, K., Sharp, L. Z. & Kajimura, S. EHMT1 controls brown adipose cell fate and thermogenesis through the PRDM16 complex. Nature 504, 163–167 (2013).

Berg, F., Gustafson, U. & Andersson, L. The uncoupling protein 1 gene (UCP1) is disrupted in the pig lineage: a genetic explanation for poor thermoregulation in piglets. PLoS Genet. 2, e129 (2006).

Gaudry, M. J. et al. Inactivation of thermogenic UCP1 as a historical contingency in multiple placental mammal clades. Sci. Adv. 3, e1602878 (2017).

Ricquier, D. & Kader, J. C. Mitochondrial protein alteration in active brown fat: a sodium dodecyl sulfate-polyacrylamide gel electrophoretic study. Biochem. Biophys. Res. Commun. 73, 577–583 (1976).

Nicholls, D. G. Hamster brown-adipose-tissue mitochondria. Purine nucleotide control of the ion conductance of the inner membrane, the nature of the nucleotide binding site. Eur. J. Biochem. 62, 223–228 (1976).

Aquila, H., Link, T. A. & Klingenberg, M. The uncoupling protein from brown fat mitochondria is related to the mitochondrial ADP/ATP carrier. Analysis of sequence homologies and of folding of the protein in the membrane. EMBO J. 4, 2369–2376 (1985).

Bouillaud, F., Ricquier, D., Thibault, J. & Weissenbach, J. Molecular approach to thermogenesis in brown adipose tissue: cDNA cloning of the mitochondrial uncoupling protein. Proc. Natl Acad. Sci. USA 82, 445–448 (1985).

Enerback, S. et al. Mice lacking mitochondrial uncoupling protein are cold-sensitive but not obese. Nature 387, 90–94 (1997).

Arsenijevic, D. et al. Disruption of the uncoupling protein-2 gene in mice reveals a role in immunity and reactive oxygen species production. Nat. Genet. 26, 435–439 (2000).

Gong, D. W. et al. Lack of obesity and normal response to fasting and thyroid hormone in mice lacking uncoupling protein-3. J. Biol. Chem. 275, 16251–16257 (2000).

Klingenberg, M. UCP1 — a sophisticated energy valve. Biochimie 134, 19–27 (2017).

Ricquier, D. UCP1, the mitochondrial uncoupling protein of brown adipocyte: a personal contribution and a historical perspective. Biochimie 134, 3–8 (2017).

Winkler, E. & Klingenberg, M. Effect of fatty acids on H + transport activity of the reconstituted uncoupling protein. J. Biol. Chem. 269, 2508–2515 (1994).

Jezek, P., Orosz, D. E., Modriansky, M. & Garlid, K. D. Transport of anions and protons by the mitochondrial uncoupling protein and its regulation by nucleotides and fatty acids. A new look at old hypotheses. J. Biol. Chem. 269, 26184–26190 (1994).

Urbankova, E., Voltchenko, A., Pohl, P., Jezek, P. & Pohl, E. E. Transport kinetics of uncoupling proteins. Analysis of UCP1 reconstituted in planar lipid bilayers. J. Biol. Chem. 278, 32497–32500 (2003).

Schreiber, R. et al. Cold-induced thermogenesis depends on ATGL-mediated lipolysis in cardiac muscle, but not brown adipose tissue. Cell Metab. 26, 753–763.e7 (2017).

Shin, H. et al. Lipolysis in brown adipocytes is not essential for cold-induced thermogenesis in mice. Cell Metab. 26, 764–777.e5 (2017).

Anderson, C. M. et al. Dependence of brown adipose tissue function on CD36-mediated coenzyme Q uptake. Cell Rep. 10, 505–515 (2015).

Putri, M. et al. CD36 is indispensable for thermogenesis under conditions of fasting and cold stress. Biochem. Biophys. Commun. 457, 520–525 (2015).

Simcox, J. et al. Global analysis of plasma lipids identifies liver-derived acylcarnitines as a fuel source for brown fat thermogenesis. Cell Metab. 26, 509–522.e6 (2017). This study identifies a mechanism whereby FFAs from adipose tissue promote acylcarnitine production in the liver, which provides fuel for cold-induced thermogenesis.

Chouchani, E. T. et al. Mitochondrial ROS regulate thermogenic energy expenditure and sulfenylation of UCP1. Nature 532, 112–116 (2016).

Wang, G. et al. Regulation of UCP1 and mitochondrial metabolism in brown adipose tissue by reversible succinylation. Mol. Cell 74, 844–857.e7 (2019).

Ukropec, J., Anunciado, R. P., Ravussin, Y., Hulver, M. W. & Kozak, L. P. UCP1-independent thermogenesis in white adipose tissue of cold-acclimated Ucp1 –/– mice. J. Biol. Chem. 281, 31894–31908 (2006).

Ikeda, K. et al. UCP1-independent signaling involving SERCA2b-mediated calcium cycling regulates beige fat thermogenesis and systemic glucose homeostasis. Nat. Med. 23, 1454–1465 (2017). This paper provides direct evidence of a UCP1-independent mechanism in beige fat that controls thermogenesis and glucose homeostasis.

de Meis, L. Uncoupled ATPase activity and heat production by the sarcoplasmic reticulum Ca 2+ -ATPase. Regulation by ADP. J. Biol. Chem. 276, 25078–25087 (2001).

Bal, N. C. et al. Sarcolipin is a newly identified regulator of muscle-based thermogenesis in mammals. Nat. Med. 18, 1575–1579 (2012).

Tajima, K. et al. Wireless optogenetics protects against obesity via stimulation of non-canonical fat thermogenesis. Nat. Commun. 11, 1730 (2020).

Aquilano, K. et al. Low-protein/high-carbohydrate diet induces AMPK-dependent canonical and non-canonical thermogenesis in subcutaneous adipose tissue. Redox Biol. 36, 101633 (2020).

Kazak, L. et al. A creatine-driven substrate cycle enhances energy expenditure and thermogenesis in beige fat. Cell 163, 643–655 (2015). This study identifies a UCP1-independent thermogenic mechanism that involves creatine futile cycling.

Kazak, L. et al. Genetic depletion of adipocyte creatine metabolism inhibits diet-induced thermogenesis and drives obesity. Cell Metab. 26, 660–671.e3 (2017).

Kazak, L. et al. Ablation of adipocyte creatine transport impairs thermogenesis and causes diet-induced obesity. Nat. Metab. 1, 360–370 (2019).

Guan, H. P. et al. A futile metabolic cycle activated in adipocytes by antidiabetic agents. Nat. Med. 8, 1122–1128 (2002).

Flachs, P. et al. Induction of lipogenesis in white fat during cold exposure in mice: link to lean phenotype. Int. J. Obes. 41, 372–380 (2017).

Reidy, S. P. & Weber, J. M. Accelerated substrate cycling: a new energy-wasting role for leptin in vivo. Am. J. Physiol. 282, E312–E317 (2002).

Silva, J. E. Thermogenic mechanisms and their hormonal regulation. Physiol. Rev. 86, 435–464 (2006).

DosSantos, R. A., Alfadda, A., Eto, K., Kadowaki, T. & Silva, J. E. Evidence for a compensated thermogenic defect in transgenic mice lacking the mitochondrial glycerol-3-phosphate dehydrogenase gene. Endocrinology 144, 5469–5479 (2003).

Anunciado-Koza, R., Ukropec, J., Koza, R. A. & Kozak, L. P. Inactivation of UCP1 and the glycerol phosphate cycle synergistically increases energy expenditure to resist diet-induced obesity. J. Biol. Chem. 283, 27688–27697 (2008).

Long, J. Z. et al. The secreted enzyme pm20d1 regulates lipidated amino acid uncouplers of mitochondria. Cell 166, 424–435 (2016).

Kajimura, S., Spiegelman, B. M. & Seale, P. Brown and beige fat: physiological roles beyond heat generation. Cell Metab. 22, 546–559 (2015).

Cooney, G. J., Caterson, I. D. & Newsholme, E. A. The effect of insulin and noradrenaline on the uptake of 2-[ 1–14 C]deoxyglucose in vivo by brown adipose tissue and other glucose-utilising tissues of the mouse. FEBS Lett. 188, 257–261 (1985).

Guerra, C. et al. Brown adipose tissue-specific insulin receptor knockout shows diabetic phenotype without insulin resistance. J. Clin. Invest. 108, 1205–1213 (2001).

Dallner, O. S., Chernogubova, E., Brolinson, K. A. & Bengtsson, T. β3-Adrenergic receptors stimulate glucose uptake in brown adipocytes by two mechanisms independently of glucose transporter 4 translocation. Endocrinology 147, 5730–5739 (2006).

Olsen, J. M. et al. Glucose uptake in brown fat cells is dependent on mTOR complex 2-promoted GLUT1 translocation. J. Cell Biol. 207, 365–374 (2014).

Lowell, B. B. et al. Development of obesity in transgenic mice after genetic ablation of brown adipose tissue. Nature 366, 740–742 (1993).

Stanford, K. I. et al. Brown adipose tissue regulates glucose homeostasis and insulin sensitivity. J. Clin. Invest. 123, 215–223 (2013).

de Souza, C. J., Hirshman, M. F. & Horton, E. S. CL-316,243, a β3-specific adrenoceptor agonist, enhances insulin-stimulated glucose disposal in nonobese rats. Diabetes 46, 1257–1263 (1997).

Roberts-Toler, C., O’Neill, B. T. & Cypess, A. M. Diet-induced obesity causes insulin resistance in mouse brown adipose tissue. Obesity 23, 1765–1770 (2015).

Bartelt, A. et al. Brown adipose tissue activity controls triglyceride clearance. Nat. Med. 17, 200–205 (2011).

Berbee, J. F. et al. Brown fat activation reduces hypercholesterolaemia and protects from atherosclerosis development. Nat. Commun. 6, 6356 (2015).

Bartelt, A. et al. Thermogenic adipocytes promote HDL turnover and reverse cholesterol transport. Nat. Commun. 8, 15010 (2017). This study reports a possible atheroprotective role of thermogenic fat via increased cholesterol flux through HDL.

Balaz, M. et al. Inhibition of mevalonate pathway prevents adipocyte browning in mice and men by affecting protein prenylation. Cell Metab. 29, 901–916.e8 (2019).

Worthmann, A. et al. Cold-induced conversion of cholesterol to bile acids in mice shapes the gut microbiome and promotes adaptive thermogenesis. Nat. Med. 23, 839–849 (2017).

Sponton, C. H. et al. The regulation of glucose and lipid homeostasis via PLTP as a mediator of BAT–liver communication. EMBO Rep. 21, e49828 (2020).

Neinast, M. D. et al. Quantitative analysis of the whole-body metabolic fate of branched-chain amino acids. Cell Metab. 29, 417–429.e4 (2019).

Yoneshiro, T. et al. BCAA catabolism in brown fat controls energy homeostasis through SLC25A44. Nature 572, 614–619 (2019). This study reports the role of thermogenic fat in BCAA metabolism and identified the first mitochondrial BCAA transporter.

Newgard, C. B. et al. A branched-chain amino acid-related metabolic signature that differentiates obese and lean humans and contributes to insulin resistance. Cell Metab. 9, 311–326 (2009).

Huffman, K. M. et al. Relationships between circulating metabolic intermediates and insulin action in overweight to obese, inactive men and women. Diabetes Care 32, 1678–1683 (2009).

Pietilainen, K. H. et al. Global transcript profiles of fat in monozygotic twins discordant for BMI: pathways behind acquired obesity. PLoS Med. 5, e51 (2008).

Wang, T. J. et al. Metabolite profiles and the risk of developing diabetes. Nat. Med. 17, 448–453 (2011).

Newgard, C. B. Interplay between lipids and branched-chain amino acids in development of insulin resistance. Cell Metab. 15, 606–614 (2012).

Liu, J. et al. Metabolomics based markers predict type 2 diabetes in a 14-year follow-up study. Metabolomics 13, 104 (2017).

Guasch-Ferre, M. et al. Metabolomics in prediabetes and diabetes: a systematic review and meta-analysis. Diabetes Care 39, 833–846 (2016).

Felig, P., Marliss, E. & Cahill, G. F. Jr. Plasma amino acid levels and insulin secretion in obesity. N. Engl. J. Med. 281, 811–816 (1969).

Crown, S. B., Marze, N. & Antoniewicz, M. R. Catabolism of branched chain amino acids contributes significantly to synthesis of odd-chain and even-chain fatty acids in 3T3-L1 adipocytes. PloS ONE 10, e0145850 (2015).

Green, C. R. et al. Branched-chain amino acid catabolism fuels adipocyte differentiation and lipogenesis. Nat. Chem. Biol. 12, 15–21 (2016).

Wallace, M. et al. Enzyme promiscuity drives branched-chain fatty acid synthesis in adipose tissues. Nat. Chem. Biol. 14, 1021–1031 (2018).

Su, X. et al. Adipose tissue monomethyl branched-chain fatty acids and insulin sensitivity: effects of obesity and weight loss. Obesity 23, 329–334 (2015).

Gunawardana, S. C. & Piston, D. W. Reversal of type 1 diabetes in mice by brown adipose tissue transplant. Diabetes 61, 674–682 (2012).

Ali Khan, A. et al. Comparative secretome analyses of primary murine white and brown adipocytes reveal novel adipokines. Mol. Cell Proteom. 17, 2358–2370 (2018).

Villarroya, J., Cereijo, R., Giralt, M. & Villarroya, F. Secretory proteome of brown adipocytes in response to camp-mediated thermogenic activation. Front. Physiol. 10, 67 (2019).

Deshmukh, A. S. et al. Proteomics-based comparative mapping of the secretomes of human brown and white adipocytes reveals EPDR1 as a novel batokine. Cell Metab. 30, 963–975.e7 (2019).

Villarroya, J. et al. New insights into the secretory functions of brown adipose tissue. J. Endocrinol. 243, R19–R27 (2019).

Whittle, A. J. et al. BMP8B increases brown adipose tissue thermogenesis through both central and peripheral actions. Cell 149, 871–885 (2012).

Svensson, K. J. et al. A secreted slit2 fragment regulates adipose tissue thermogenesis and metabolic function. Cell Metab. 23, 454–466 (2016).

Kristof, E. et al. Interleukin-6 released from differentiating human beige adipocytes improves browning. Exp. Cell Res. 377, 47–55 (2019).

Sun, K. et al. Dichotomous effects of VEGF-A on adipose tissue dysfunction. Proc. Natl Acad. Sci. USA 109, 5874–5879 (2012).

Mahdaviani, K., Chess, D., Wu, Y., Shirihai, O. & Aprahamian, T. R. Autocrine effect of vascular endothelial growth factor-A is essential for mitochondrial function in brown adipocytes. Metabolism 65, 26–35 (2016).

Cereijo, R. et al. CXCL14, a brown adipokine that mediates brown-fat-to-macrophage communication in thermogenic adaptation. Cell Metab. 28, 750–763.e6 (2018).

Campderros, L. et al. Brown adipocytes secrete GDF15 in response to thermogenic activation. Obesity 27, 1606–1616 (2019).

Nisoli, E., Tonello, C., Benarese, M., Liberini, P. & Carruba, M. O. Expression of nerve growth factor in brown adipose tissue: implications for thermogenesis and obesity. Endocrinology 137, 495–503 (1996).

Zeng, X. et al. Innervation of thermogenic adipose tissue via a calsyntenin 3β-S100b axis. Nature 569, 229–235 (2019).

Wang, G. X. et al. The brown fat-enriched secreted factor Nrg4 preserves metabolic homeostasis through attenuation of hepatic lipogenesis. Nat. Med. 20, 1436–1443 (2014).

Kong, X. et al. Brown adipose tissue controls skeletal muscle function via the secretion of myostatin. Cell Metab. 28, 631–643.e3 (2018).

Ruan, C. C. et al. A2A receptor activation attenuates hypertensive cardiac remodeling via promoting brown adipose tissue-derived FGF21. Cell Metab. 28, 476–489.e5 (2018).

Lynes, M. D. et al. The cold-induced lipokine 12,13-diHOME promotes fatty acid transport into brown adipose tissue. Nat. Med. 23, 631–637 (2017). This study reports a cold-inducible batokine, 12,13-diHOME, that stimulates fatty acid uptake in brown fat.

Stanford, K. I. et al. 12,13-DiHOME: an exercise-induced lipokine that increases skeletal muscle fatty acid uptake. Cell Metab. 27, 1111–1120.e3 (2018).

Chen, Y. et al. Exosomal microRNA miR-92a concentration in serum reflects human brown fat activity. Nat. Commun. 7, 11420 (2016).

Thomou, T. et al. Adipose-derived circulating miRNAs regulate gene expression in other tissues. Nature 542, 450–455 (2017).

Sun, K., Tordjman, J., Clement, K. & Scherer, P. E. Fibrosis and adipose tissue dysfunction. Cell Metab. 18, 470–477 (2013).

Lackey, D. E. et al. Contributions of adipose tissue architectural and tensile properties toward defining healthy and unhealthy obesity. Am. J. Physiol. 306, E233–E246 (2014).

Muir, L. A. et al. Adipose tissue fibrosis, hypertrophy, and hyperplasia: correlations with diabetes in human obesity. Obesity 24, 597–605 (2016).

Divoux, A. et al. Fibrosis in human adipose tissue: composition, distribution, and link with lipid metabolism and fat mass loss. Diabetes 59, 2817–2825 (2010).

Reggio, S. et al. Increased basement membrane components in adipose tissue during obesity: links with TGFβ and metabolic phenotypes. J. Clin. Endocrinol. Metab. 101, 2578–2587 (2016).

Henegar, C. et al. Adipose tissue transcriptomic signature highlights the pathological relevance of extracellular matrix in human obesity. Genome Biol. 9, R14 (2008).

Khan, T. et al. Metabolic dysregulation and adipose tissue fibrosis: role of collagen VI. Mol. Cell. Biol. 29, 1575–1591 (2009).

Hasegawa, Y. et al. Repression of adipose tissue fibrosis through a PRDM16–GTF2IRD1 complex improves systemic glucose homeostasis. Cell Metab. 27, 180–194.e6 (2018).

Wang, W. et al. A PRDM16-driven metabolic signal from adipocytes regulates precursor cell fate. Cell Metab. 30, 174–189.e5 (2019).

Heaton, J. M. The distribution of brown adipose tissue in the human. J. Anat. 112, 35–39 (1972).

Hany, T. F. et al. Brown adipose tissue: a factor to consider in symmetrical tracer uptake in the neck and upper chest region. Eur. J. Nucl. Med. Mol. Imaging 29, 1393–1398 (2002).

Cohade, C., Osman, M., Pannu, H. K. & Wahl, R. L. Uptake in supraclavicular area fat (“USA-Fat”): description on 18 F-FDG PET/CT. J. Nucl. Med. 44, 170–176 (2003).

van Marken Lichtenbelt, W. D. et al. Cold-activated brown adipose tissue in healthy men. N. Engl. J. Med. 360, 1500–1508 (2009).

Virtanen, K. A. et al. Functional brown adipose tissue in healthy adults. N. Engl. J. Med. 360, 1518–1525 (2009).

Saito, M. et al. High incidence of metabolically active brown adipose tissue in healthy adult humans: effects of cold exposure and adiposity. Diabetes 58, 1526–1531 (2009).

Cypess, A. M. et al. Identification and importance of brown adipose tissue in adult humans. N. Engl. J. Med. 360, 1509–1517 (2009).

Leitner, B. P. et al. Mapping of human brown adipose tissue in lean and obese young men. Proc. Natl Acad. Sci.USA 114, 8649–8654 (2017). This study maps brown fat in six distinct anatomical depots in young men, comparing lean individuals and individuals with obesity.

Chen, K. Y. et al. Brown adipose Reporting Criteria in Imaging STudies (BARCIST 1.0): recommendations for standardized FDG-PET/CT experiments in humans. Cell Metab. 24, 210–222 (2016).

Sharp, L. Z. et al. Human BAT possesses molecular signatures that resemble beige/brite cells. PloS ONE 7, e49452 (2012).

Yoneshiro, T. et al. Recruited brown adipose tissue as an antiobesity agent in humans. J. Clin. Invest. 123, 3404–3408 (2013).

Hanssen, M. J. et al. Short-term cold acclimation improves insulin sensitivity in patients with type 2 diabetes mellitus. Nat. Med. 21, 863–865 (2015).

Chondronikola, M. et al. Brown adipose tissue improves whole-body glucose homeostasis and insulin sensitivity in humans. Diabetes 63, 4089–4099 (2014).

Lee, P. et al. Temperature-acclimated brown adipose tissue modulates insulin sensitivity in humans. Diabetes 63, 3686–3698 (2014).

Hanssen, M. J. et al. Short-term cold acclimation recruits brown adipose tissue in obese humans. Diabetes 65, 1179–1189 (2016). This study shows that short-term cold exposure can lead to the recruitment of brown fat in humans with obesity.

Vijgen, G. H. et al. Increase in brown adipose tissue activity after weight loss in morbidly obese subjects. J. Clin. Endocrinol. Metab. 97, E1229–E1233 (2012).

Raiko, J., Orava, J., Savisto, N. & Virtanen, K. A. High brown fat activity correlates with cardiovascular risk factor levels cross-sectionally and subclinical atherosclerosis at 5-year follow-up. Arterioscler. Thromb. Vasc. Biol. 40, 1289–1295 (2020). This study finds that the presence of cold-induced brown fat activity correlates with lower cardiovascular risk factors and decreased carotid intima-media thickness and higher carotid elasticity on 5-year follow-up.

Becher, T. et al. Brown adipose tissue is associated with cardiometabolic health. Nat. Med. 27, 58–65 (2021). This study finds that brown fat in humans is associated with protection from cardio-metabolic diseases, particularly in individuals that are overweight and obese.

Ma, S. et al. Caloric restriction reprograms the single-cell transcriptional landscape of rattus norvegicus aging. Cell 180, 984–1001.e22 (2020).

Yoneshiro, T. et al. Impact of UCP1 and β3AR gene polymorphisms on age-related changes in brown adipose tissue and adiposity in humans. Int. J. Obes. 37, 993–998 (2013).

Bakker, L. E. et al. Brown adipose tissue volume in healthy lean South Asian adults compared with white Caucasians: a prospective, case-controlled observational study. Lancet Diabetes Endocrinol. 2, 210–217 (2014).

Vosselman, M. J., Vijgen, G. H., Kingma, B. R., Brans, B. & van Marken Lichtenbelt, W. D. Frequent extreme cold exposure and brown fat and cold-induced thermogenesis: a study in a monozygotic twin. PloS ONE 9, e101653 (2014).

Riveros-McKay, F. et al. Genetic architecture of human thinness compared to severe obesity. PLoS Genet. 15, e1007603 (2019).

Zhang, F. et al. An adipose tissue atlas: an image-guided identification of human-like BAT and beige depots in rodents. Cell Metab. 27, 252–262.e3 (2018).

Fitzgibbons, T. P. et al. Similarity of mouse perivascular and brown adipose tissues and their resistance to diet-induced inflammation. Am. J. Physiol. Heart Circ. Physiol. 301, H1425–H1437 (2011).

Sacks, H. S. et al. Uncoupling protein-1 and related messenger ribonucleic acids in human epicardial and other adipose tissues: epicardial fat functioning as brown fat. J. Clin. Endocrinol. Metab. 94, 3611–3615 (2009).

Lynch, C. J. & Adams, S. H. Branched-chain amino acids in metabolic signalling and insulin resistance. Nat. Rev. Endocrinol. 10, 723–736 (2014).

Villarroya, F., Cereijo, R., Villarroya, J., Gavalda-Navarro, A. & Giralt, M. Toward an understanding of how immune cells control brown and beige adipobiology. Cell Metab. 27, 954–961 (2018).

Sakamoto, T. et al. Macrophage infiltration into obese adipose tissues suppresses the induction of UCP1 level in mice. Am. J. Physiol. 310, E676–E687 (2016).

Goto, T. et al. Proinflammatory cytokine interleukin-1β suppresses cold-induced thermogenesis in adipocytes. Cytokine 77, 107–114 (2016).

Valladares, A., Roncero, C., Benito, M. & Porras, A. TNF-α inhibits UCP-1 expression in brown adipocytes via ERKs. Opposite effect of p38MAPK. FEBS Lett. 493, 6–11 (2001).

Chiang, S. H. et al. The protein kinase IKKε regulates energy balance in obese mice. Cell 138, 961–975 (2009).

Mowers, J. et al. Inflammation produces catecholamine resistance in obesity via activation of PDE3B by the protein kinases IKKε and TBK1. eLife 2, e01119 (2013).

Kumari, M. et al. IRF3 promotes adipose inflammation and insulin resistance and represses browning. J. Clin. Invest. 126, 2839–2854 (2016).

Yadav, H. et al. Protection from obesity and diabetes by blockade of TGF-β/Smad3 signaling. Cell Metab. 14, 67–79 (2011).

Koncarevic, A. et al. A novel therapeutic approach to treating obesity through modulation of TGFβ signaling. Endocrinology 153, 3133–3146 (2012).

Guo, T. et al. Adipocyte ALK7 links nutrient overload to catecholamine resistance in obesity. eLife 3, e03245 (2014).

Rajbhandari, P. et al. Single cell analysis reveals immune cell-adipocyte crosstalk regulating the transcription of thermogenic adipocytes. eLife 8, e49501 (2019).

Rajbhandari, P. et al. IL-10 signaling remodels adipose chromatin architecture to limit thermogenesis and energy expenditure. Cell 172, 218–233 e217 (2018). This study characterizes adipocytes and stromal cells identifying crosstalk between immune cells and thermogenic adipocytes.

Wolf, Y. et al. Brown-adipose-tissue macrophages control tissue innervation and homeostatic energy expenditure. Nat. Immunol. 18, 665–674 (2017).

Pirzgalska, R. M. et al. Sympathetic neuron-associated macrophages contribute to obesity by importing and metabolizing norepinephrine. Nat. Med. 23, 1309–1318 (2017).

Camell, C. D. et al. Inflammasome-driven catecholamine catabolism in macrophages blunts lipolysis during ageing. Nature 550, 119–123 (2017).

Chung, K. J. et al. A self-sustained loop of inflammation-driven inhibition of beige adipogenesis in obesity. Nat. Immunol. 18, 654–664 (2017).

Hu, B. et al. γδ T cells and adipocyte IL-17RC control fat innervation and thermogenesis. Nature 578, 610–614 (2020).

Brestoff, J. R. et al. Group 2 innate lymphoid cells promote beiging of white adipose tissue and limit obesity. Nature 519, 242–246 (2015).

Lee, M. W. et al. Activated type 2 innate lymphoid cells regulate beige fat biogenesis. Cell 160, 74–87 (2015).

Zhang, X. et al. Functional inactivation of mast cells enhances subcutaneous adipose tissue browning in mice. Cell Rep. 28, 792–803.e4 (2019).

Finlin, B. S. et al. Mast cells promote seasonal white adipose beiging in humans. Diabetes 66, 1237–1246 (2017).

Lynch, L. et al. iNKT cells induce FGF21 for thermogenesis and are required for maximal weight loss in GLP1 therapy. Cell Metab. 24, 510–519 (2016).

Cypess, A. M. et al. Cold but not sympathomimetics activates human brown adipose tissue in vivo. Proc. Natl Acad. Sci. USA 109, 10001–10005 (2012).

Cypess, A. M. et al. Activation of human brown adipose tissue by a β3-adrenergic receptor agonist. Cell Metab. 21, 33–38 (2015).

O’Mara, A. E. et al. Chronic mirabegron treatment increases human brown fat, HDL cholesterol, and insulin sensitivity. J. Clin. Invest. 130, 2209–2219 (2020). This study shows that chronic treatment with mirabegron increases human brown fat activity, which is associated with increased HDL and improved insulin sensitivity.

Blondin, D. P. et al. Human brown adipocyte thermogenesis is driven by β2-AR stimulation. Cell Metab. 32, 287–300.e7 (2020).

Broeders, E. P. et al. The bile acid chenodeoxycholic acid increases human brown adipose tissue activity. Cell Metab. 22, 418–426 (2015).

Ramage, L. E. et al. Glucocorticoids acutely increase brown adipose tissue activity in humans, revealing species-specific differences in UCP-1 regulation. Cell Metab. 24, 130–141 (2016).

Yoneshiro, T., Aita, S., Kawai, Y., Iwanaga, T. & Saito, M. Nonpungent capsaicin analogs (capsinoids) increase energy expenditure through the activation of brown adipose tissue in humans. Am. J. Clin. Nutr. 95, 845–850 (2012).

Ohyama, K. et al. A synergistic antiobesity effect by a combination of capsinoids and cold temperature through promoting beige adipocyte biogenesis. Diabetes 65, 1410–1423 (2016).

Wang, S. et al. Curcumin promotes browning of white adipose tissue in a norepinephrine-dependent way. Biochem. Biophys. Res. Commun. 466, 247–253 (2015).

Jiang, J. et al. Cinnamaldehyde induces fat cell-autonomous thermogenesis and metabolic reprogramming. Metabolism 77, 58–64 (2017).

5.5: Uncoupling Electron Transport from ATP Synthesis - Biology

Cellular respiration is the set of metabolic reactions used by cells to harvest energy from food. The catabolism of glucose under aerobic conditions occurs in three sequential metabolic pathways: glycolysis, pyruvate oxidation, and the citric acid cycle. The reduced coenzymes produced from these metabolic pathways are then oxidized by the respiratory chain, and ATP is made. By these pathways, glucose has been completely oxidized and the cell has gained many molecules of ATP—a versatile energy carrier that fuels most kinds of cellular work.

In this tutorial we will examine the operation of the electron transport chain and the production of ATP. In cellular respiration, it is the action of the electron transport chain that produces the bulk of the ATP for the cell.


During the early phases of cellular respiration, glucose is completely broken down. CO2 is liberated into the atmosphere, and the hydrogen atoms from glucose are donated to the energy carriers NAD + and FAD to form NADH + H + and FADH2. In order for cellular respiration to continue to operate on additional glucose molecules, these energy carriers must be recycled.

The work of the respiratory chain is, in part, to recycle these carriers. The carriers donate their extra hydrogen atoms to the respiratory chain and thereby convert back into NAD + and FAD. In the accompanying animation, we focused on NADH, which donates a hydrogen atom to the first complex in the chain. FADH2 (not shown in the animation) donates to a different complex.

The other work of the respiratory chain is to transform the chemical energy of the hydrogen atoms (specifically, their electrons) into potential energy. In a series of redox reactions, electrons jump from one complex to another and, in the process, release energy. The chain uses the released energy to pump protons across the membrane, from a region of low concentration inside the mitochondrion to a region of high concentration within the intermembrane space. This concentration gradient represents potential energy.

The cell taps the potential energy of the gradient when protons flow back across the membrane through a pore in the ATP synthase complex. As the protons flow, they release energy, which the complex uses to convert ADP and inorganic phosphate to ATP. The production of ATP from energy derived from the flow of electrons through the respiratory chain is referred to as oxidative phosphorylation. Chemiosmosis is another term for ATP synthesis, referring to the use of a proton gradient to fuel the production of ATP.

Textbook Reference: Concept 6.2 Carbohydrate Catabolism in the Presence of Oxygen Releases a Large Amount of Energy

Watch the video: Uncoupling of Electron Transport Chain (January 2022).