Download Cardiac mitochondrial matrix and respiratory complex - AJP

Am J Physiol Heart Circ Physiol 303: H940–H966, 2012.
First published August 10, 2012; doi:10.1152/ajpheart.00077.2012.
Review
Cardiac mitochondrial matrix and respiratory complex protein phosphorylation
Raul Covian and Robert S. Balaban
Laboratory of Cardiac Energetics, National Heart Lung and Blood Institute, Bethesda, Maryland
Submitted 26 January 2012; accepted in final form 1 August 2012
kinase; phosphatase; citric acid cycle; oxidative phosphorylation; protein transport;
mitochondria intermembrane space; phosphohistidine; adenine nucleotide translocase; pyruvate dehydrogenase; branched chain ␣-ketoacid dehydrogenase
is part of a collection on Post-translational Protein Modification in Metabolic Stress. Other articles appearing in this collection, as well as a full archive of
all Review collections, can be found online at http://ajpheart.
physiology.org/.
THIS REVIEW ARTICLE
Introduction
The heart is a tissue with one of the highest rates of energy
conversion in the body, being critically dependent on mitochondrial oxidative phosphorylation as a major source of
adenosine 5=-triphosphate (ATP). The remarkably fast turnover
of ATP in the heart is in the order of 10 s at high workloads
(12, 13), and a mismatch of just a few percent between the
production and utilization rates can result in major disruptions
Address for reprint requests and other correspondence: R. S. Balaban, Laboratory of Cardiac Energetics, National Heart Lung and Blood Institute, 9000
Rockville Pike, Bethesda, MD, 20817 (e-mail: [email protected]).
H940
in the potential energy available for function (12, 13). The
dysregulation of the cardiac energy conversion process has
been suggested to be one of the major elements in heart failure
(144). Thus the precise orchestration of mitochondrial energy
conversion is critical for normal heart function, though the
mechanism for this regulatory process is still under investigation. Since enzyme protein phosphorylation [i.e., pyruvate
dehydrogenase (PDH) and glycogen phosphorylase (GP)] was
one of the first mechanism of modulating metabolic processes
to match physiological needs, it was logical to further explore
enzyme protein phosphorylation at other levels of energy
metabolism of the heart. Therefore, the focus of the present
article is to review the evidence, or lack thereof, supporting a
role of protein phosphorylation in the regulation of mitochondrial oxidative phosphorylation. It should be pointed out that
other post-translational modifications (PTMs) reviewed elsewhere, such as acetylation (125), S-nitrosylation (142), nitration (35), and glutathionylation (91), among others, have also
been implicated in the regulation of mitochondrial metabolism.
http://www.ajpheart.org
Downloaded from http://ajpheart.physiology.org/ by 10.220.32.246 on May 5, 2017
Covian R, Balaban RS. Cardiac mitochondrial matrix and respiratory complex
protein phosphorylation. Am J Physiol Heart Circ Physiol 303: H940 –H966, 2012.
First published August 10, 2012; doi:10.1152/ajpheart.00077.2012.—It has become
appreciated over the last several years that protein phosphorylation within the
cardiac mitochondrial matrix and respiratory complexes is extensive. Given the
importance of oxidative phosphorylation and the balance of energy metabolism in
the heart, the potential regulatory effect of these classical signaling events on
mitochondrial function is of interest. However, the functional impact of protein
phosphorylation and the kinase/phosphatase system responsible for it are relatively
unknown. Exceptions include the well-characterized pyruvate dehydrogenase and
branched chain ␣-ketoacid dehydrogenase regulatory system. The first task of this
review is to update the current status of protein phosphorylation detection primarily
in the matrix and evaluate evidence linking these events with enzymatic function or
protein processing. To manage the scope of this effort, we have focused on the
pathways involved in energy metabolism. The high sensitivity of modern methods
of detecting protein phosphorylation and the low specificity of many kinases
suggests that detection of protein phosphorylation sites without information on the
mole fraction of phosphorylation is difficult to interpret, especially in metabolic
enzymes, and is likely irrelevant to function. However, several systems including
protein translocation, adenine nucleotide translocase, cytochrome c, and complex
IV protein phosphorylation have been well correlated with enzymatic function
along with the classical dehydrogenase systems. The second task is to review the
current understanding of the kinase/phosphatase system within the matrix. Though
it is clear that protein phosphorylation occurs within the matrix, based on 32P
incorporation and quantitative mass spectrometry measures, the kinase/phosphatase
system responsible for this process is ill-defined. An argument is presented that
remnants of the much more labile bacterial protein phosphoryl transfer system may
be present in the matrix and that the evaluation of this possibility will require the
application of approaches developed for bacterial cell signaling to the mitochondria.
Review
MITOCHONDRIA PROTEIN PHOSPHORYLATION
catalyzes the phosphorylation of many proteins in cell extracts,
consistent with a low absolute specificity. Indeed, as stated by
the authors of KESTREL, a significant challenge in this type of
study is deciphering the kinase phosphorylations that impact
function versus neutral sites. Taking this concept further,
several web-based tools (NETPHOS, NETPHOSK, and GPS)
are available to predict the vulnerability of a given protein
amino acid to kinase activity. These probabilities are based on
a variety of criteria (22, 224, 226). Using the human mitochondrial complex V ␥-subunit as an example (sp_P36542–2_
ATPG_HUMAN), the GPS and NETPHOS predicted ⬎46
phosphorylation sites (see Fig. 1) from numerous kinases based
on the consensus recognition of these enzymes even using the
highest threshold of detection. Similar results were obtained
with NETPHOSK. These results support the notion that protein
kinases are rather “nonspecific,” depending on what threshold
is applied. In general, some threshold of “phosphorylation
potential” is selected to determine the likelihood of phosphorylation by a given kinase. This is illustrated by the gray line in
Fig. 1A. The issue with many of today’s methods of detecting
phosphorylated peptides is that no threshold, in terms of mole
fraction or “occupancy,” is given for the protein phosphorylations reported. Indeed, in most cases the mole fraction of a
phosphorylated peptide is simply not known. Thus one of the
reasons for the explosion in detection of protein phosphorylation sites is that we are simply dropping the threshold for
detection with our sensitive methods revealing potentially less
specific and possibly neutral phosphorylation sites. This is
analogous to bringing the gray line in Fig. 1A closer to zero as
the methods get more sensitive detecting more of the “low
specificity” sites of protein kinase activity.
The lack of absolute specificity of kinases together with the
increased sensitivity of current phosphopeptide detection methods might explain the finding of a number of phosphorylation
sites in the presequence of mitochondrial proteins, including
NADH dehydrogenase ubiqinone flavoprotein (NDUF)S4 (208)
and NDUFS8 (48, 94) of complex I, electron transfer flavoprotein
(ETF) ␣-subunit (ETFA) (227), succinate dehydrogenase subunit
A (flavoprotein) (SDHA) in complex II (151), ubiquinolcytochrome c reductase binding protein (UQCRB) in complex III
(24), and subunits-␤ (151) and c (217) of complex V. Some of
these sites were found in isolated mitochondria, where the
amount of preassembled precursors is very low compared with
the mature and assembled proteins. These sites are likely
irrelevant in terms of the function of the enzymatic complexes.
However, it could be argued that these phosphorylation sites
are important for protein import and, consequently, affect
steady-state levels of mitochondrial complexes. Except for a
few isolated cases, such as NDUFS4 at a site within the mature
protein (54, 55) and several subunits of the translocase of the
outer membrane (TOM) complex (170, 191), phosphorylation
has not been correlated with protein import to mitochondria.
An interesting example is the ␤-subunit of complex V, where
a significant amount of phosphorylated precursor was detected
when cAMP levels were increased by treatment of mouse
lymphoma cells with isoproterenol (202). However, import
was not altered by this PTM, and the phosphorylated precursor
was either rapidly degraded or dephosphorylated before assembly into the mature complex. The conclusion of this study was
that phosphorylation of the nascent subunit-␤ polypeptide by
PKA in the cytosol was merely accidental and physiologically
AJP-Heart Circ Physiol • doi:10.1152/ajpheart.00077.2012 • www.ajpheart.org
Downloaded from http://ajpheart.physiology.org/ by 10.220.32.246 on May 5, 2017
Since the late 1960s, protein phosphorylation was implicated
as one of the regulatory mechanisms balancing energy conversion with utilization in the heart. Protein phosphorylation of
mitochondrial PDH, described almost simultaneously by Linn
et al. (123) and Weiland and colleagues (219, 220), was one of
the first examples of metabolic control by enzyme phosphorylation. This discovery and the previously described regulation
of GP by protein phosphorylation (114) were two of the early
examples of acute modulation of enzyme activity by protein
phosphorylation events [see the early review (194)], and both
of these reactions are important for cardiac energy conversion.
PDH constitutes the major entry point of reducing equivalents
from glycolysis to oxidative phosphorylation. Indeed, PDH
molar activity via dephosphorylation increases with cardiac
workload in the perfused heart (113, 132), consistent with its
playing a role in the adaptation of metabolic conversion rates
to cardiac workload. Similarly, glycogen breakdown, via GP,
has been correlated with the early metabolic response to
cardiac work transitions (72). Thus protein phosphorylation
has already been demonstrated to play a key role in the
regulation of cardiac energy metabolism by regulating the
delivery of reducing equivalents to the citric acid cycle.
Many models of the regulation of cardiac mitochondrial
oxidative phosphorylation suggest a distributed control of
enzymatic activity (11, 237). Therefore, it is a reasonable
extrapolation, based on PDH and GP regulation, to look first at
mitochondrial protein phosphorylation events that might contribute to metabolic regulation. Today, due to the explosion of
proteomic screening methods with remarkable sensitivity, the
number of potentially important mitochondrial protein phosphorylation sites is staggering. It is also important to note that
the original results on PDH and GP were derived by initially
observing acute alterations in enzymatic activity in vitro and
then determining the PTMs responsible. Because of the current
explosion in phosphoprotein analysis, this proven and timehonored strategy has been generally reversed. That is, many
PTMs are being constantly discovered in the mitochondrial
proteome that are yet to be established as functionally significant, rather than mining for PTMs as sources of demonstrated
alterations in extracted enzyme molar activities. This reversed
strategy may be an inefficient approach for many reasons. First,
many protein amino acids, and likely associated PTMs, are not
conserved and apparently not critical for enzyme function.
Thus PTMs on these amino acids, and even on some that are
conserved, will likely not influence function if they are not in
a critical region of the enzyme. In addition, the actual energetic
cost of most cellular protein PTMs, assuming a slow turnover,
is very low compared with the demands of differentiated
function, such as muscle contraction, especially in active tissues like the heart. This energetic analysis suggests that the
effect of neutral PTM events on the natural selection of
PTM-generating systems (i.e., kinases) might be very low.
Indeed, it is likely more “expensive” to “tune” a kinase for a
specific set of active sites than to allow some neutral phosphorylation sites to be populated at negligible cost in energy or
function. These neutral sites may also contribute to signaling
variation when one of these sites becomes “active” during
mutation alterations in protein sequence.
What is the absolute selectivity of known protein kinases?
Several protein 32P phosphorylation screening methods [see
KESTREL (111), for example] show that a given kinase
H941
Review
H942
MITOCHONDRIA PROTEIN PHOSPHORYLATION
Phosphorylation potential
A
Serine
Threonine
Tyrosine
Threshold
1
0
50
100
150
200
250
300
Sequence position
B
Fig. 1. Predicted phosphorylation sites in the human mitochondrial complex V ␥-subunit. The P-Ser (green), P-Thr (blue), and P-Tyr (red) sites predicted by the
NETPHOS (A) and GPS (B) program (224) on the human ␥-subunit sequence (sp_P36542–2_ATPG_HUMAN). A: graphical presentation of phosphorylation
potential as a function of position. Gray vertical line indicates a 0.5 potential. B: graphical presentation of GPS model with the “high” threshold. Agreement
between methods was good; similar results were obtained with NETPHOSK.
irrelevant, although tolerated. Results like this highlight that
unspecific phosphorylation events do occur in experimental
settings and not just in algorithm predictions, although not all
sites found in silico will necessarily be found experimentally.
To complicate matters further, very low levels of nonenzymatic phosphorylation of Tyr, Ser, and even Thr residues can
be achieved when proteins are warmed in the presence of
divalent cations (189), so the greater instrument sensitivity
achieved requires additional precautions in sample preparation
to avoid detection of deeply substoichiometric events that are
completely unrelated to biology. Because of the nonquantitative nature of most phosphoprotein detection schemes, we
rarely know the mole fraction of a phosphopeptide, so the
extent of this problem is still unknown and a major concern of
these authors.
In this review, we will attempt to initially provide an
updated list of protein phosphorylation sites within the mitochondrial matrix enzymes involved in energy metabolism.
These proteins are divided into membrane transporters, intermediary metabolism enzymes, and the complexes of oxidative
phosphorylation. We also review the current understanding of
the matrix kinase/phosphatase system. In some cases, because
of the lack of information, data outside of the heart will be
included. A critical evaluation will be provided of which of
these modifications have clearly been linked to alterations in
activity using the criteria outlined below. In addition, we will
attempt to characterize the sites with regard to the probability
that they may influence function based on our understanding of
the molecular mechanisms within the oxidative phosphorylation complexes. Since this exciting field is moving very
quickly, we apologize in advance for any literature omissions
in this survey that we are sure will occur at our error.
The criteria for establishing the functional impact of a
protein phosphorylation event are highly variable depending on
the genetic control of the system, as well as the cellular tools
available to modify the protein and the cellular physiology. In
general, the simple correlation of protein phosphorylation with
enzymatic activity in vivo and/or in vitro is the first line of
evidence and is an absolute requirement as an initial step.
Ideally, these initial correlations should be accomplished with
physiological manipulations that reflect the operation of the
heart in vivo. Another important element is to demonstrate that
a significant mole fraction of the enzyme is affected by protein
phosphorylation. This is particularly important when dealing
with cardiac energy conversion enzymes since at “resting”
metabolic rates most of the enzymes are present in excess and
only approach Vmax conditions at maximum workloads (140,
163). Thus, to significantly impact energy metabolism along
AJP-Heart Circ Physiol • doi:10.1152/ajpheart.00077.2012 • www.ajpheart.org
Downloaded from http://ajpheart.physiology.org/ by 10.220.32.246 on May 5, 2017
0
Review
MITOCHONDRIA PROTEIN PHOSPHORYLATION
it would depend on the particular enzyme, but it can be
assumed that when movement is required at subunit interfaces
during the catalytic cycle, one or more phosphorylation events
could potentially block activity. Phosphorylation sites close to
the entry or exit points for protons in respiratory enzymes are
also potentially relevant if they can block access to protons
channels or affect the pKa of residues that are sequentially
protonated and deprotonated to facilitate proton movement
(135). We will attempt to point out phosphorylation sites that
have been found in heart or at least in muscle, taking into
consideration what is known about the particular mechanism of
each enzymatic complex and its structure even when there is an
absence of functional data. We omit from the discussion sites
that have been reported in other tissues, unless they are located
in a structurally relevant region of the protein or have been
associated with kinases or modifications of enzymatic activity.
A significant number of sites have been reported in cancer cell
lines or tumors (33, 37, 39, 61, 73, 95, 107, 131, 141, 150, 151,
176, 223, 231) but are not discussed further in this review
under the assumption that metabolic regulation in these cells is
in disarray and is therefore largely unrelated to the physiology
of differentiated tissues like the heart.
Phosphorylation of Inner Membrane Transporters
The critical communication between the mitochondrion and
the cytosol across the outer mitochondrial membrane and the
remarkably high-resistance inner membrane is dependent on
numerous transporters. Thus modulation of these transporters
via protein phosphorylation could dramatically affect the ability of the cytosol to influence mitochondrial reaction pathways
via the exchange of metabolites and signaling molecules, as
well as proteins.
The ADP/ATP translocase (ANT) performs the exchange of
cytosolic ADP with matrix ATP generated by oxidative phosphorylation (109). Phosphorylation sites have been found in all
of the isoforms of ANT by numerous investigators. These sites
include, in ANT-1, Ser7 (24, 236), Ser22 (24, 236), Tyr81 (24),
Thr84 (24), Tyr191 (236), and Tyr195 (24, 67); in ANT-2,
Thr84 (68, 236), Tyr191 (68), and Tyr195 (68); and in ANT-3
(236), Tyr112, Ser119, and Tyr195. All these phosphorylation
sites are shown in Fig. 2. ANT of cardiac mitochondria from
pig and rat was phosphorylated with ␥-[32P]ATP in blue native
gels, suggesting the possibility of autophosphorylation activity
(161), although the specific sites are unknown. A series of
studies by Feng et al. (67, 68) have confirmed the phosphorylation of Tyr191 and Tyr195 on the H4 helix of ANT-1 (158),
showing that phosphorylation is required for maximal translocase function. In addition, Src kinases are capable of phosphorylating ANT-1 in vitro, whereas phosphorylation in intact cells
was found to be blocked with 4-amino-5-(4-chloro-phenyl)-7(t-butyl)pyrazolol[3,4,d]pyrimidine (PP2), a specific inhibitor
of Src kinases. Several phenotypic changes in cells, including
the inhibition of oxidative growth in yeast (68), are induced by
genetically modifying these phosphorylation sites, and Tyr
phosphorylation of ANT-1 has also been correlated with preconditioning in the heart (68). The Tyr phosphorylation sites
on the H4 helix are exposed at the outer surface of the inner
membrane (158), where intermembrane space Src kinases
could phosphorylate these sites without requiring translocation
of these kinases into the matrix. The one missing piece of
AJP-Heart Circ Physiol • doi:10.1152/ajpheart.00077.2012 • www.ajpheart.org
Downloaded from http://ajpheart.physiology.org/ by 10.220.32.246 on May 5, 2017
the energy conversion pathway, a significant fraction of the
enzyme must be modified to have an impact on the metabolic
rate. That is, if only 2% of a given complex subunit is
phosphorylated, this will have little or no impact on the net flux
through the system where flux alterations approach 500% from
rest to maximum workloads. Naturally, any site with a 2%
PTM mole fraction could grow to 100% under different experimental conditions. Thus low mole fraction PTMs do not rule
out a site as being important under all conditions, just under the
condition evaluated. Indeed, detection of the phosphorylation
site confirms the presence of a potent regulating kinase. The
importance of PTM mole fraction is not as critical for signaling
molecules, such as kinases, acetylases, deacetylases, phosphatases, or ion channels, where the modification of a few copies
can influence numerous downstream events or the membrane
potential. This requirement is often overlooked when using
high-sensitivity methods for detecting phosphorylated proteins,
including phosphopeptide enhancement mass spectrometry
(MS) (24, 58, 121, 165, 236), radiolabeling (6, 29, 128), and
high-sensitivity MS (156, 175), all of which permit the detection of a few copies of phosphorylated peptides in the enzyme
protein pool. Another line of evidence for the regulatory role of
a protein phosphorylation site may include changes in enzyme
activity upon specific amino acid substitution that simulate or
eliminate the PTM (1, 101, 159). Though this might be a
powerful approach in some circumstances, the substitution
frequently leads to false-positive results because of changes in
protein structure or assembly induced by the amino acid
substitution that are seductive but could alter activity in a
manner that does not actually occur upon phosphorylation of
the wild-type residue. Finally, direct structural simulation studies demonstrating alteration of enzyme active sites or other
critical structural parameters by phosphorylation provide the
most reliable and definite confirmation of the functional relevance. Regrettably, as this review will illustrate, the vast
majority of protein phosphorylation sites detected in the mitochondrial matrix proteome, which increase in number by the
month, have not even been correlated with enzymatic activity,
limiting the evaluation of their relevance to the examination of
available protein structures to obtain a glimpse of which sites
could in theory affect function.
A particular phosphorylation site can be safely assumed to
have a potential impact in enzymatic function if it is located in
the active site and if phosphorylation would sterically hinder
the binding of substrates or interact with residues that participate in catalysis. The same can be said of sites that are in
contact with a redox group such as a heme, an iron sulfur
center, a copper atom, or a flavin molecule and that should
significantly decrease the midpoint potential of the cofactor by
the presence of a negative charge upon phosphorylation. However, it is more difficult to predict the impact of phosphorylation at sites that are farther away from a redox group, given that
conformational changes can have a large role in determining
the electrochemical properties of the cofactor (97). In this
regard, experiments in which charges have been introduced by
mutagenesis at different distances from a redox group suggest
that electrostatic interactions become greatly attenuated at
distances larger than ⬃10 Å because of the high dielectric
constant in the interior of most proteins (124). The potential
functional role of phosphorylation sites found on the interface
of subunits within a protein complex cannot be generalized, for
H943
Review
H944
MITOCHONDRIA PROTEIN PHOSPHORYLATION
Y112
Y195
S119
Y191
CAT
T84
Y81
S7
Fig. 2. Reported sites of phosphorylation in the ADP/ATP translocase (ANT).
The structure of the bovine ANT-1 (PDB ID 1OKC) crystallized in the
presence of carboxyatractyloside (CAT) is shown looking down from the
intermembrane space to the central cavity. Only the side chains of those
residues reported to be sites of phosphorylation are shown space-filled on
the protein C␣ backbone with colors representing each type of amino acid (Ser,
red; Thr, green; Tyr, purple).
evidence in this extensive series of studies is a measure of the
mole fraction of phosphorylated ANT-1. To our knowledge,
these sites have only been detected by MS after phosphopeptide enrichment or with very sensitive antibodies and radio
labeling. This might imply that only a small fraction of ANT-1
is phosphorylated and thus active, which could contribute to
the rate limitation of oxidative phosphorylation by this transporter, although this latter point is still the subject of controversy. A determination of the mole fraction of ANT-1 phosphorylated at Tyr191 and -195 would be extremely useful in
characterizing the impact of regulation by PTMs under physiological conditions. However, Boja et al. (24) found essentially no change in Tyr196 (Tyr195 in human) phosphorylation
in pig isolated cardiac mitochondria after treatment with dichloroacetate (DCA) or upon Ca2⫹ activation, with a statistically insignificant increase of only 20% when mitochondria
were deenergized. Since these studies were conducted on
isolated mitochondria, the recruitment of cytosolic kinases,
such as the Src kinases, could not occur and may influence
these negative results.
Another inner membrane transport protein that is believed to
play a role in the maintenance of the total adenosine pool of the
matrix is the ATP-Mg/Pi carrier protein (7). This transporter is
phosphorylated at Tyr324 (68), but no functional data have
been reported for this site. Metabolites for intermediary metabolism also need to be transported across the inner membrane. In a very recent high-sensitivity screen by Zhao et al.
(236), the aspartate-glutamate transporter Aralar 1 was found
phosphorylated at Ser662 and -101, whereas the 2-oxogluta-
Phosphorylation of the Enzymes of Intermediary Metabolism
The phosphorylation of PDH via the PDH kinase (PDHK)
and PDH phosphatase (PDHP) system is the first mitochondrial
enzyme shown to be regulated by protein phosphorylation as
reviewed in the introduction. PDH phosphorylation and regulation are thus well documented and will not be discussed at
length here. We will further discuss the PDHK and PDHP
system in the kinase/phosphatases discussion below. Readers
interested in this specific system are referred to the original
papers (60, 102, 123, 219, 220) as well as the several reviews
available on this topic (59, 133). Since quantitative measures
are one of the focuses of this review, it is interesting to note
that Boja et al. (24) performed one of the first quantitative MS
studies on PDH phosphorylation in isolated heart mitochondria. The perturbations previously shown to activate PDH that
were used in this study included additions of DCA or Ca2⫹, as
well as substrate starved deenergization. These perturbations
significantly decreased PDH serine phosphorylation, with
Ser292 and Ser231 phosphorylation decreasing in the order of
three- to fourfold and correlating the best with the modulation
of enzyme activity. These studies demonstrate that quantitative
AJP-Heart Circ Physiol • doi:10.1152/ajpheart.00077.2012 • www.ajpheart.org
Downloaded from http://ajpheart.physiology.org/ by 10.220.32.246 on May 5, 2017
S22
rate/malate carrier was phosphorylated at several sites (Ser6,
Tyr102, Tyr202, and Ser203). Boja et al. (24) also found the
Tyr102 site in addition to Thr36 on this transporter. Cui et al.
(47) found several serine phosphorylation sites on two cationic
amino acid transporters: Slc7a4 (Ser418, -421, and -422) and
Slc7a2 (Ser463 and -645, as well as Thr466). No information
on the functional significance or mole fraction of these phosphorylation sites is currently available.
The voltage-dependent anion channel (VDAC), or mitochondrial porin, is believed to play a major role in metabolite
exchange across the outer membrane of the mitochondrion and
is thus an important link in the energy conversion process (42).
Though VDAC is not directly linked to the mitochondrial
matrix, it is important to note that the phosphorylation of this
transporter has been detected in most mitochondrial phosphoproteome screens and that correlations with VDAC permeability (19, 38) and enzyme complex formation (157, 197) have
been reported. Thus this important solute transport process in
the outer membrane may be under significant regulation by
cytosolic kinases (such as GSK3b, PKA, and NEK1) and
phosphatases.
It is clear that protein phosphorylation occurs on many of the
transporters in the mitochondrial membranes. Of the numerous
phosphorylations detected, the phosphorylation of ANT-1 and
VDAC has been correlated with the function of these transporters. It should also be pointed out that the major protein
transport pathway in the mitochondria, the TOM and translocase of the inner membrane (TIM) complex is also phosphorylated. Recently, Schmidt et al. (191) demonstrated that protein
phosphorylation can both activate and inhibit the important
protein translocation function of TOM. These data suggest a
major role for protein phosphorylation in the regulation of the
exchange of proteins (TOM) and metabolites (VDAC) between
the cytosol and the mitochondria intermembrane space. To
date, most of these regulatory sites have been found in the
mitochondrial intermembrane space or in the cytosol; thus a
role for a matrix kinase/phosphatase system in this control
network is unclear.
Review
MITOCHONDRIA PROTEIN PHOSPHORYLATION
other kinases such as p38 MAPK, no specific sites have been
found (196). Therefore, unlike the case for PDH, which regulates the entry of reducing equivalents from glycolysis, the role
of phosphorylation as a regulatory mechanism for fatty acid
oxidation seems to be weak. Many phosphorylation sites have
been reported in the rest of the enzyme network of fatty acid
oxidation. Most of the acyl dehydrogenases were detected
using phosphoryl-protein sensitive dyes in heart (6, 83). In
mouse heart (94), the following sites have been located in the
acyl-CoA dehydrogenase family members: in ACAD9, Ser478,
Thr482, and Thr488; in ACADL, Ser54, Ser55, Ser191 (not
conserved in human), and Ser200; and in ACADM, Ser70 and
Thr351. In enoyl-CoA hydratase (ECHS1), Thr194 was found
to be phosphorylated in rat heart (68), whereas in the ␣-subunit
of trifunctional enzyme [hydroxyacyl-CoA dehydrogenase/3ketoacyl-CoA thiolase/enoyl-CoA hydratase (trifunctional protein) ␣-subunit], the phosphorylation sites found in mouse
heart (58, 94) are Tyr283, Ser316, Thr575, Thr586, and Ser647.
Phosphorylation of acetoacetyl-CoA thiolase (ACAT1), which
catalyzes the last step of ketone body catabolism, has also been
reported in mouse heart at Tyr90 and Ser238 (58, 94). However, no functional correlation of any of the phosphorylation
sites in these enzymes with regard to fatty acid or ketone body
oxidation rates has been presented.
Thus, despite the recent screening studies revealing many
new phosphorylation sites in the matrix citric acid and fatty
acid oxidation enzyme system, the functional significance, mole
fraction, and kinase/phosphatase system remain unknown.
These sites might suggest that protein phosphorylation may
play a broader role in the regulation of the citric acid cycle
beyond PDH as well as fatty acid oxidation; however, further
investigations will be required to establish whether this regulatory system is in play.
Phosphorylation of Complex I
Complex I, a proton pumping NADH dehydrogenase, is the
largest of the oxidative phosphorylation complexes, with a
total of 46 subunits in mammals, 14 of which are the core of
electron and proton transfer activities, and conserved in prokaryotes and eukaryotes (26). A total of 69 different phosphorylation sites have been reported in 26 of the complex I subunits.
These are summarized in Fig. 3, A and B. To emphasize the
extensive nature of protein phosphorylation detection, we are
presenting two schematics for each complex: the first (A)
places all of the phosphorylations roughly across the whole
complex, and the second and subsequent B–D (when applicable in the corresponding figure) present a space filling model
with the phosphorylation sites shown relative to active sites as
discussed in the introduction. In complex I, 18 of the sites are
located in the seven central proteins that comprise the membrane extrinsic peripheral arm of complex I that contains all of
the redox cofactors necessary for function, and one more site is
found in the hydrophobic core proteins that are essential for
proton pumping. Since the only high-resolution structures
available for the peripheral arm of complex I are bacterial (20,
64, 188), the relevance of those phosphorylation sites that have
not been linked yet to some change in activity by empirical
evidence are restricted to these eight subunits. The remaining
50 sites are distributed among 18 of the 33 accessory subunits
for which structural information is not yet available. Since the
AJP-Heart Circ Physiol • doi:10.1152/ajpheart.00077.2012 • www.ajpheart.org
Downloaded from http://ajpheart.physiology.org/ by 10.220.32.246 on May 5, 2017
comparisons of phosphorylation and enzyme activity are feasible using rather unambiguous MS techniques and should be
useful in evaluating the functional consequences of phosphorylation sites in other mitochondrial enzymes. Another enzyme
system very similar to PDH is the branched-chain ␣-ketoacid
dehydrogenase (BCKDH) present in the heart and other tissues
for the metabolism of the three essential amino acids: valine,
isoleucine, and leucine (76). This enzyme has been demonstrated to have serine phosphorylation sites on its E1B␣ subunit
(89) that inhibit the enzyme in a similar fashion as they do in
the PDH complex. A separate BCKDH kinase and phosphatase
have also been described with properties that are very similar
to the PDH system (49, 53, 166, 173). The phosphorylation of
BCKDH has been shown to occur along with that of PDH in
the intact heart using 32P (28), which correlated with the
activity of the extracted enzymes. However, Boja et al. (24)
actually found that phosphorylation of BCKDH E1B␣ at
Ser337 increased with DCA and deenergization in isolated
heart mitochondria, whereas that of the E1 subunit of PDH
decreased in the same study. The reason for this discrepancy is
unknown and may reflect differences in the control of the
BCKDH and PDHK/PDHP system.
In addition to the well characterized PDH and BCKDH
systems, most of the enzymes of the citric acid cycle have been
found to possess phosphorylation sites. These include in succinyl-CoA ligase [ADP forming] subunit-␤, Ser257, Ser259
(236), and Ser241 (24); in citrate synthase, Ser190 (236) and
Tyr80 (68); in isocitrate dehydrogenase [NADP], Ser423
(236); in malate dehydrogenase, mitochondrial Ser246, Thr235
(24, 236), Ser51, Ser31 (236), Tyr56, and Tyr80 (68); in
fumarate hydratase, Tyr491 (68); and in aconitate hydratase,
Ser669 (236), Tyr71, Tyr544, and Tyr655 (68). However, none
of these phosphorylations have been shown to modify the
activity of these enzymes, and inspection of crystal structures
shows that these sites are away from the substrate binding
pockets.
An interesting observation was made on succinyl-CoA ligase, where a tightly associated phosphate was found on the
enzyme using 32P binding assays that correlated with enzymatic activity (162). However, it was found that this tightly
associated phosphate that survived detergent, SDS, and twodimensional electrophoresis is not a protein phosphorylation,
but a noncovalent interaction that increased the maximum
velocity of the succinyl-CoA ligase reaction kinetics. This
tightly associated phosphate in succinyl-CoA ligase suggests
that similar metabolite associations may occur in other systems
and may require careful interpretation of radiolabeling experiments (5).
A major source of reducing equivalents in the heart under
normal conditions is fatty acid oxidation. Phosphorylation of
carnitine palmitoyltransferase-1 (CPT-1) in the outer mitochondrial membrane has been reported at Ser741 and Ser747 of
the liver isoform both in vivo and in vitro using protein kinase
casein kinase II, with the consequence of a slightly higher
activity and a different inhibitory effect of malonyl-CoA (103).
In isolated heart mitochondria, where the muscle isoform of
CPT-1 is predominant, incubation with PKA and CAMK-II
also generated slight differences in malonyl-CoA sensitivity of
CPT-1 without altering the catalytic activity, and although
changes in the phosphorylation levels of Ser and Thr residues
in the muscle isoform of CPT-1 were reported using these and
H945
Review
H946
MITOCHONDRIA PROTEIN PHOSPHORYLATION
A
INTERMEMBRANE SPACE
B
Complex I
S117
NDUFA10:
NDUFB11:
NDUFB4:
NDUFB7:
S59 S231
S20
S26 S38
S73 Y89
Y133
NDUFA8:
T29
NDUFA1:
NDUFB6:
S144
Y142
NDUFB5:
T2 T113
S55
NDUFA3:
S182
NDUFB10:
NDUFA4:
ND3:
(Y41/S42)
NDUFB9: S21 Y55
S67
S52
S85
Y56 Y143
MATRIX
Y141
Y86
FeS
T127
T125
FeS
FeS
eS
T611
FeS
S250
FeS
FeS FeS
FeS
S478
S479
T164
NADH
FMN
S250
Fig. 3. Reported sites of phosphorylation in complex I. Residues where phosphorylation has been detected in each indicated subunit are shown schematically
(A) with respect to their presumed relevance based on structural and functional evidences in a structural scheme. Sites that are unlikely to have an effect on activity
based on their location in available high-resolution structures are shown in white. Sites with no known function, but that might influence activity based on their
location in the structure, are shown in yellow. Sites that have been found to be phosphorylated by an identified kinase in vitro or in vivo are shown in purple.
Sites that are located in the presequence and are thus absent in the assembled complex are shown in brackets. The position of each site either in the matrix space
or closer to the inner mitochondrial membrane is also shown based on the structural information available. The probable location of some of these sites is shown
(B) using the structure of the hydrophilic domain of complex I from Thermus thermophilus (PDB ID 3IAM) as a scaffold in which the residues that are
homologous to the mammalian amino acids are colored based on the identity of the reported phosphorylation site (Ser, red; Thr, green; Tyr, purple). The C␣
backbone of NDUFV1 is shown in cyan, NDUFV2 is colored dark green, NDUFS1 is shown in orange, NDUFS2 in pink, NDUFS3 in bright green, NDUFS7
in red, and NDUFS8 in blue. Redox groups (FMN and FeS centers) are space-filled and colored magenta, and NADH is represented as a blue wire model. See
main text for definitions of abbreviations.
role and interaction of all of the subunits of complex I is yet to
be resolved, much of the extrapolation of the impact of a given
protein phosphorylation on a specific site must be considered,
to some extent, to be speculative.
NADH binds to the NDUFV1 subunit, which contains FMN
and one iron sulfur center as electron acceptors (20, 26). This
protein was labeled with a phosphospecific dye (ProQ) in pig
heart (83). Only one phosphorylation site has been found in
this subunit in cancer cells at the COOH-terminal residue
Ser464 (151). The location of this residue is unknown given
the lack of conservation in this region between mammalian
NDUFV1 and the Thermus thermophilus structure but is probably located far away from the redox centers and the NADH
binding site. The only phosphorylation site reported in the
essential subunit NDUFV2, also in HeLa cells (151), is
Thr164. The corresponding residue in the bacterial structure is
Thr112, which sits in a relatively well-conserved loop highly
exposed to the solvent but far away from its iron sulfur center
or subunit contact regions (20). Nevertheless, this subunit is
labeled by ProQ and PhosTag dyes in pig heart, but not in liver
(6), suggesting that other more physiologically relevant sites
may exist that have escaped detection. The chain of iron sulfur
centers continues in the NDUFS1 subunit, the largest of complex I, which contains three of these redox cofactors. Interestingly, this subunit is not only labeled with ProQ and PhosTag
in a tissue specific manner but also with 32P in energized
mitochondria, suggesting a higher turnover rate of phosphorylation (6). Of the five phosphorylation sites detected in
NDUFS1, only Ser250 has been recently found in heart tissue
(58). However, the corresponding residue in the bacterial
structure (20) is not in contact with any other subunit interface
(see Fig. 3B) and is close to an iron sulfur cluster that does not
even exist in eukaryotes (26), casting doubt on its functional
relevance.
The metazoan specific subunit NDUFV3 is located somewhere close to the NDUFV1 and NDUFV2 subunits, given that
the three subunits can be dissociated as a subcomplex from the
rest of complex I (69). NDUFV3 is the most heavily phosphorylated protein of NADH dehydrogenase, with 17 different sites
in non-muscle tissues (40, 58, 61, 74, 80, 84), mostly in a string
of Ser residues from position 152 to 161 and the nearby Ser163
and Ser165 of isoform 2, which exists in human and rodents.
Some of these Ser phosphorylations seem to have been detected in human skeletal muscle (236). Another isoform 2 site
is the Ser located four residues from the COOH-terminus (62),
which exists in both isoforms and has also been detected in
mouse brown fat, heart, and kidney (94). Further studies on the
effect of NDUFV3 isoform expression on complex I activity
are needed to understand the role of these phosphorylation
sites, especially in the heart.
Subunit NDUFS8, also known as TYKY, contains two more
iron sulfur clusters and has also been found to bind phosphospecific dyes in pig heart mitochondria (6, 83). Of the four
phosphorylation sites reported for this protein, only Tyr86,
found in rat inner medulla cells (81), exists in the mature
protein. Tyr86 is close to the interface with NDUFS7 and to the
AJP-Heart Circ Physiol • doi:10.1152/ajpheart.00077.2012 • www.ajpheart.org
Downloaded from http://ajpheart.physiology.org/ by 10.220.32.246 on May 5, 2017
NDUFS7:
NDUFA2:
S39 S41 S117
S7 S78
T125 T127 NDUFS2:
NDUFA7:
NDUFS8: Y141
S63 S96
[Y2]
[S5] [S6]
NDUFS6:
Y51
Y86
NDUFS3:
S59
NDUFS1:
NDUFS4:
S250 S461,
[T32] [S34]
S478 S479 T611
S144 S165
S173
NDUFV3:
S77 S98 S152-161,163,165,237,466
T247
NDUFV2:
NDUFV1: T164
S464
S59
Review
MITOCHONDRIA PROTEIN PHOSPHORYLATION
phosphomimetic substitutions in nearby Ser residues in hamsters show a slight decrease in complex I activity (225),
although in this case the evidence is more indirect because of
the lack of residue conservation. Phosphorylation of a 18-kDa
complex I protein, later shown to correspond to NDUFB11
(36), was reported to increase activity of complex I (210),
although the fraction of total protein that underwent this
modification was not quantified. Tyr133 and Ser144 have also
been reported as phosphorylation sites in human muscle, but
only using highly sensitive methods (236).
Seventeen more sites have been found in other 10 accessory
subunits of the membrane domain. Sites found in heart (94) or
muscle (236) are Tyr41 (or Ser42) of NDUFA3, Ser67 of
NDUFA4, Ser26 and Ser38 of NDUFB4, Ser21 of NDUFB10
[which is labeled by ProQ in pig heart (83)], and Ser73 and
Tyr89 in NDUFB7. NDUFA13, also known as GRIM19, an
apoptosis-inducing factor that is essential for complex I assembly, was found to have a Ca2⫹-insensitive phosphorylation site
at Thr96, detected in pig heart (24). Another accessory subunit,
NDUFS5, has no reported phosphorylation sites, although it
was found to be labeled both by ProQ and PhosTag dyes in
pig heart (but not liver) mitochondria (6, 83). However, any
functional effect of phosphorylation in these accessory subunits is probably restricted to protein assembly, given that most
are either cysteine-rich chaperones likely involved in iron
sulfur cluster synthesis and insertion or short single transmembrane domain polypeptides proposed to aid in the assembly and
stability of the membrane domain (26), but not likely to be
involved directly in proton pumping, given their absence from
prokaryotic complex I.
Phosphorylation of Complex II and the ETF System
Succinate dehydrogenase (complex II) and the enzymes of
the ETF system share ubiquinone as final electron acceptor
in the mitochondrial inner membrane. Complex II is anchored
to the membrane and oxidizes succinate to fumarate, whereas
the ETF system is composed of the heterodimeric soluble ETF
that shuttles electrons from a variety of dehydrogenases, most
notably the acyl dehydrogenases of fatty acid oxidation, to the
membrane-associated ETF: ubiquinone oxidoreductase (ETFDH).
A summary of the identified phosphorylation sites in these
enzymes are schematically presented in Fig. 4, A–C. Phosphorylation of the two hydrophilic subunits of complex II, SDHA,
and succinate dehydrogenase subunit B (iron-sulfer protein)
(SDHB) has been detected in heart and liver mitochondria
using phosphosensitive dyes (6, 83). Phosphorylated subunit-␤
of ETF (ETFB) has also been identified using dyes in mitochondria from heart and more prominently in liver, where
ETFB is also labeled by 32P (6, 83).
The flavoprotein subunit of complex II (SDHA) is where
succinate binding and oxidation takes place. Nine different
phosphorylation sites have been reported in this flavin adenine
dinucleotide (FAD)-containing protein, five of which are tyrosines. The crystal structure of complex II (205) shows that all
the reported sites are located on the surface of SDHA away
from the succinate binding site, the FAD binding pocket, and
the interface with SDHB (see Fig. 4B), in line with the lack of
evidence for a functional role at any of these sites. Two
tyrosine phosphorylation sites (Tyr535 and Tyr596 in rat brain
mitochondria) were detected after in vitro incubation of solu-
AJP-Heart Circ Physiol • doi:10.1152/ajpheart.00077.2012 • www.ajpheart.org
Downloaded from http://ajpheart.physiology.org/ by 10.220.32.246 on May 5, 2017
membrane (14, 20), but away from redox centers (see Fig. 3B),
so its functional effects, if any, remain obscure. Another
central subunit of the peripheral arm labeled with PhosTag in
heart but not in liver (6) is NDUFS2, which lacks redox
cofactors but is phosphorylated in lung cancer cells at Tyr141
(176), located at the interface with subunit NDUFS7, also
known as the PSST protein, which contains the last iron sulfur
center in the chain (Fig. 3B). This interface has been proposed
to form the quinone reduction site (26) and is therefore an ideal
target of regulation if phosphorylation were to occur under
physiological conditions and at a significant mole fraction,
which is not documented yet. Phosphorylation of NDUFS3,
another core protein that lacks redox centers and is labeled with
PhosTag in both heart and liver mitochondria (6), has been
reported at Ser59 in rat uterus homogenates upon addition of
exogenous PKA after heating to inactivate endogenous kinase
and phosphatase activities (88). Even though this residue lies in
a potentially relevant region at the interface with NDUFS2
(Fig. 3B), the experimental manipulations applied to detect it
are not applicable to physiological conditions. Also, the presence of PKA within the matrix space is still in question (see
Mitochondrial Matrix Protein Kinases).
NDUFS7 has five reported phosphorylation sites, two of
them found in mouse heart and kidney at Ser39 and Ser41 (94)
in a poorly conserved and undefined region of the crystal
structure that seems to interact with subunits of the membrane
domain of complex I (20, 64). Thr125, discovered in HeLa
cells upon treatment with rapamycin (37), is in contact with the
iron sulfur center of NDUFS7 if the orientation of the residue
in metazoans is similar to the corresponding residue (Arg) in
the bacterial crystal structure (see Fig. 3B). Unfortunately, no
functional information exists on the effect of phosphorylation
at any of these sites on enzyme activity.
No structural information is available for the accessory
subunits of the peripheral arm, so the role of the phosphorylation sites described in five of these proteins is even less clear.
In the case of Ser173 of NDUFS4, at least a correlation has
been observed between the amount of imported protein and the
phosphorylation of this residue by PKA (54, 55). Ser96 of
NDUFA7 was found to be phosphorylated in bovine heart
(165), although phosphopeptide enrichment was needed to
detect it, as was the case with Ser78 of NDUFA2, found in
mouse brown fat, heart, kidney, and brain (94, 222), suggesting
a very low mole fraction of phosphorylation.
Accessory subunits in the membrane domain that are not
found in prokaryotes have been the focus of more intensive
studies with respect to phosphorylation and its effects, in
particular NDUFA10, NDUFA1 (also called MWFE), and
NDUFB11 (known as ESSS). Ser59 (not well conserved) and
Ser231 of NDUFA10 have been reported to be phosphorylated
in bovine and mouse heart, respectively (58, 190), although no
correlation to function or quantification of mole fraction has
been established. This subunit has also been found to bind the
PhosTag dye in pig heart but not in liver (6). Ser55 of
NDUFA1 was phosphorylated in bovine heart complex I when
incubated in vitro with PKA and cAMP (36, 225). Phosphomimetic mutations of this residue in hamster cells abolish
complex I assembly, suggesting a function for phosphorylation
at this site in regulating the steady-state levels of the enzyme
(225). In the case of NDUFB11, Ser20 was also found to be
phosphorylated in the presence of exogenous PKA (36), and
H947
Review
H948
MITOCHONDRIA PROTEIN PHOSPHORYLATION
A
ETF system
Complex II
INTERMEMBRANE SPACE
ETFDH:
S264
T352
Y361
S550
ETFA:
[S15]
T58
T93
S140
T143
S207 T213
S227
S281
ETFB:
Y2 T8
T77
S98
T194
T242
S234
MATRIX
B
T119
Y208
SDHB:
Y218
S224
SDHA: Y535
[T24]
Y596
Y629
Y365
T638
S509
S530 Y641
C
TTF
T242
b
TTF
Y208
S227
FeS
T234
S281
T119
Y218
FeS
S224
FeS
S207
T213
S509
Y365
FAD
Y535
T143
S190 T93
S530
Y629
eS
FeS
T58
S189
FAD
S140
S264
S550
T77
T
Y361
T194
FAD
UQ
T638
T352
Y641
Y596
bilized mitochondrial fractions with the tyrosine kinase Fgr
followed by enrichment of phosphotyrosine peptides (180), but
it is still unknown what kinases might be responsible for
complex II phosphorylation in vivo. Interestingly, Phillips et
al. (161) found SDHA as one of the 32P labeled proteins
obtained after incubation of blue native gels with ␥-[32P]ATP,
suggesting an intrinsic kinase activity in complex II or in one
of the proteins present in the corresponding region of the gel,
where no known kinases were detected.
The iron sulfur subunit of complex II (SDHB) contains three of
these redox centers and links SDHA to the two smaller hydrophobic subunits embedded in the membrane where ubiquinone
binds and undergoes reduction (205). Four phosphorylation sites
have been reported in SDHB, one of which, Tyr208, identified in
mouse brown fat and testis (94), stands out as potentially relevant
in view of its position in contact with the iron sulfur center of
SDHB that is closer to the membrane, in the vicinity of one of the
ubiquinone binding sites (see Fig. 4B).
Nine phosphorylation sites comprising only Ser and Thr
residues have been found in ETFA, six of which have been
found differentially expressed in mouse tissues, including
Thr93 in heart (94, 222). Despite the lack of studies establishing a correlation with function, three phosphorylation sites
could have some effect on the activity of ETF based on the
crystal structures available (177, 211). Thr143, found in mouse
liver (94), is close to the interface with the ␤-subunit (ETFB),
so it could be important for the stability of the dimer, whereas
Ser227 (found in brown fat) is in the FAD domain that moves
during interaction with dehydrogenases (see Fig. 4C). Ser281,
also detected in mouse brown fat (94), forms a hydrogen bond
to one of the phosphates of FAD, so its phosphorylation could
alter or even impede assembly of the FAD cofactor, although
in the absence of functional data, it could simply represent an
assembly intermediate (94).
The seven phosphorylation sites identified in ETFB include
Thr77, detected in a human-mouse myeloid chimeric cell line
(39), which is located at the surface of ETFB where acyl
dehydrogenases bind (211), so it could theoretically affect
electron transfer rates. Ser98, Ser234, and Thr242 were all
found in mouse brown fat (94), and the last two of these
residues are located, respectively, in the loop that connects to
the COOH-terminal domain of ETFB that interacts with the
movable FAD domain of ETFA and in the interface with ETFA
close to the FAD binding pocket (Fig. 4C). Again, the lack of
functional studies and the likelihood of a low fraction of
phosphorylated protein at these sites preclude any conclusions
on a physiological role of these sites.
ETFDH has an iron sulfur center as well as one FAD as
redox cofactors, and the crystal structure has shown ubiquinone in its binding pocket (235). Only one of the four reported
AJP-Heart Circ Physiol • doi:10.1152/ajpheart.00077.2012 • www.ajpheart.org
Downloaded from http://ajpheart.physiology.org/ by 10.220.32.246 on May 5, 2017
Fig. 4. Reported phosphorylation sites in complex II and the ETF system. Subunits where
phosphorylated residues have been found are
represented schematically along with the respective residues (A) using the same color
coding as in Fig. 2A. All the sites shown are
exposed to the mitochondrial matrix. The location of the reported phosphorylation sites in
complex II (B) and the ETF system (C) is
shown according to their amino acid identities
(Ser, red; Thr, green; Tyr, purple). The structure shown for complex II is that of the pig
heart mitochondrial complex II (PDB ID
1ZP0), with the C␣ backbone SDHA colored
in cyan, SDHB in blue, SDHC in green, and
SDHD in dark yellow. The two molecules of
thenoyltrifluoroacetone (TTFA) shown in blue
close to the b heme serve to identify the
ubiquinone binding sites. The structure used
for ETF and ETFDH are those from human
(PDB ID 1EFV) and pig (PDB ID 2GMH),
respectively. ETFA is colored dark pink,
ETFB is shown in blue, and ETFDH is orange. Redox cofactors (FAD, FeS centers and
the b heme) are colored magenta. Ubiquinone
(UQ) in ETFDH is shown as a blue wire
model. See main text for definitions of
abbreviations.
Review
MITOCHONDRIA PROTEIN PHOSPHORYLATION
phosphorylation sites in EFTDH, Tyr361, found in lung cancer
cells (176), is close to one of the redox cofactors, in this case,
the iron sulfur center (see Fig. 4C). Ser264 and Ser550,
identified respectively in mouse brown fat and in other tissues
including heart and liver (94, 213), are exposed at the edge of
a flat surface opposite to Thr352, which was found to be
phosphorylated in pig heart, but which did not change in its
phosphorylation level upon energization of mitochondria (24),
suggesting a low turnover and low mole fraction.
Complex III Phosphorylation
contrast to UQCRC1 and UQCRC2, has a central role in
electron transfer (230) and would thus constitute a plausible
site for regulation. UQCRFS1 has a tilted transmembrane helix
that traverses and possibly stabilizes the complex III dimer,
and its COOH-terminal half includes a globular domain in the
intermembrane space that contains the 2Fe-2S center that
accepts one of the two electrons from ubiquinol (96, 230). This
extramembrane domain moves from the vicinity of CYTB to
interact with CYC1, functioning as a diffusible electron shuttle
(46). This movement has been proposed to be involved in the
asymmetric activity of complex III that results in the oxidation
of ubiquinol in only one monomer at a time (45). The
UQCRFS1 subunit has been reported to exhibit a phosphatasesensitive shift in its isoelectric point when rat liver mitochondria are exposed to calcium or agents that induce the permeability transition of the mitochondrial membrane, suggesting a
change in phosphorylation state of UQCRFS1 (78). However,
no specific phosphorylation site has been linked to this isoelectric shift, which has not been observed in heart mitochondria
(83). Moreover, UQCRFS1 phosphorylation has been detected
with dyes such as ProQ and PhosTag, but not with 32P labeling
(6, 83), suggesting a low turnover rate of phosphorylation,
which is inconsistent with a role in acute regulation of mitochondrial function, unless a very labile phosphorylation is
assumed. Nevertheless, UQCRFS1 phosphorylation, as reported by binding of antibodies against P-Ser, was observed to
decrease by up to 50% upon ischemic preconditioning in rat
brain, which was accompanied by an almost twofold increase
in complex III activity (52). Thus it is quite feasible that some
P-Ser constitutes a regulatory site in UQCRFS1.
Two of the reported sites in UQCRFS1 correspond to Ser
residues detected in resting human skeletal muscle mitochondria (236). Ser99 is located on the matrix side in a bend right
before the beginning of the transmembrane helix close to the
NH2-terminus of UQCRC1, and thus away from the extrinsic
domain which moves to transfer electrons (see Fig. 5B). Ser157
is located at the top of the movable domain, but away from the
2Fe-2S cluster and the series of residues that control its redox
potential (199), and also at a considerable distance from the
contact surface that interacts with CYTB or CYC1. Although
phosphorylation of Ser157 would not seem to interfere with the
electron transfer function of UQCRFS1 directly, it could serve
as a docking point for other proteins, which upon binding could
hinder the diffusion of this extrinsic domain. However, no
evidence exists yet for such an interaction.
CYC1 is key to the electron transfer function of complex III
because of its binding and electron transfer to cytochrome c.
Interestingly, cytochrome c binding has been found to occur
only to one of the CYC1 subunits in the complex III dimer,
according to crystallized structures from yeast mitochondria
(116, 200). Only one phosphorylation site, Ser182, has recently
been reported in mitochondria from resting human muscle
(236). The equivalent residue in crystallographic structures is
close to the point of contact between the two CYC1 subunits in
the dimer, but far from the actual binding site of cytochrome c
or the c1 heme group. Furthermore, Ser182 is only found in
primates, whereas Pro is found at this position in most other
vertebrates, casting doubt on a functional role for phosphorylation at this site.
Another subunit that is relevant for cytochrome c binding to
complex III is the acidic “hinge” protein, or subunit 6
AJP-Heart Circ Physiol • doi:10.1152/ajpheart.00077.2012 • www.ajpheart.org
Downloaded from http://ajpheart.physiology.org/ by 10.220.32.246 on May 5, 2017
The cytochrome bc1 complex (complex III) is a dimer
comprised of 11 different subunits per monomer in mammalian
mitochondria (228). Phosphorylation has been detected in eight
of these polypeptides, including the three subunits that contain
redox prosthetic groups and are thus directly involved in
electron transfer from ubiquinol to cytochrome c coupled to
proton translocation, namely, cytochrome b (CYTB), the
Rieske iron sulfur protein (UQCRFS1), and cytochrome c1
(CYC1). Of the 24 phosphorylation sites identified so far in
complex III, 16 occur in Ser residues, 6 in Tyr, and only 1 in
Thr. A summary of these sites is presented in Fig. 5, A and B.
More than half of the phosphorylation sites reported in
complex III are located in the two largest subunits, the
ubiquinol cytochrome c reductase core 1 (UQCRC1) and core
2 (UQCRC2) proteins, which protrude toward the matrix side
of the inner mitochondrial membrane (228), which is consistent with a matrix kinase activity. These proteins, however, do
not participate in electron transfer or proton movement within
complex III, although they are required for the assembly of
CYC1, UQCRFS1, and other smaller subunits, as shown by
deletion strains in yeast (232). UQCRC1 and UQCRC2 are
homologous to the dimeric matrix processing metalloproteases
that cleave the presequence of a considerable number of
proteins translated in the cytosol and imported into the mitochondria. Some crystal structures even show the cleaved presequence of UQCRFS1, termed subunit 9, bound to UQCRC2,
which suggests that these subunits are responsible for processing of UQCRFS1 upon its assembly into complex III (96).
Mild detergent treatment activates the UQCRC1-UQCRC2
protease activity in isolated mammalian complex III, although
the catalytic rate is considerably lower than that of the soluble
matrix protease (57). However, no phosphorylation sites have
been found in the active protease site. More importantly,
neither UQCRC1 nor UQCRC2 has been found to change its
phosphorylation state upon manipulation of cellular mitochondrial metabolism (24), and the turnover is slow as judged by
the lack of labeling when mitochondria are incubated with 32P
(6), suggesting that the significance of the phosphorylation
status is likely limited to their role in the assembly of complex
III. An interesting phosphorylation site is Ser473, reported so
far only in mouse kidney (94), but which can be seen in close
proximity to a phosphate molecule (Fig. 5B) in one crystal
structure from bovine heart (87), suggesting a high mole
fraction of phosphorylation. The phosphate molecule is also in
contact with His-221 of CYTB in a region where mutagenesis
in nearby residues has revealed assembly defects or altered
binding properties at the ubiquinone reduction site (27).
The only other complex III subunit where more than one
phosphorylation site has been reported is UQCRFS1, which, in
H949
Review
H950
MITOCHONDRIA PROTEIN PHOSPHORYLATION
A
Complex III
T29 S48
Complex IV
COX2:
COX6B:
S126 Y218 S52 S71
COX4 : COX1:
COX6C:
S158
S115
Y57
S116
Y304
Y49 Y98
S182
S89
c1
Cytochrome c
UQCRFS1:
S157
CYC1:
UQCRH:
S182
S89
CYTB:
Y75
S157
B
INTERMEMBRANE SPACE
FeS
bL
Stg
pY75
Y115
bH Ant
pS473
COX4:
Y33
S58
S72
T74
S89
COX5A:
S47
T78
T79
Y468
COX7C:
S17
S99 S102 Y448
COX5B:
S71
S124
Y459
S88
T86 S87 S226
Y207
S212
S107 S74
MATRIX
S247
S437
S368
C
D
Y218
S126
Y49
S52
S71
Y57
S158
C A
CuA
S116S115
a
S48
S89
Y98
T29
Y304
30
a3 CuB
Y304
30
S115
a3
3
a
T79 T78
Y33
S71
T74
S58
S72
S124
Fig. 5. Reported phosphorylation sites in complex III, cytochrome c, and complex IV. Residues in the indicated subunits or proteins where phosphorylation has
been detected are shown schematically in different colors to indicate their presumed or demonstrated relevance based on structural and functional evidences (A).
Sites that are unlikely to have an effect on activity based on their location in available high-resolution structures are shown in white. Sites with no known function,
but that might influence activity based on their location in the structure, are shown in yellow. Sites that have been found to affect enzymatic activity when
phosphorylated are shown in red. Sites that are located in the presequence and are thus absent in the assembled complex are shown in brackets. The position
of each site relative to the intermembrane space and the matrix is also shown. The location of most of the phosphorylation sites are shown in the crystal structure
of complex III (B), cytochrome c (C), and complex IV (D), colored according to the type of amino acid (Ser, red; Thr, green; Tyr, purple). Phosphate molecules
are shown in contact with two of the reported sites in complex III (pY75 in CYTB and pS473 in UQCRC1), according to one of the bovine structures (PDB
ID 1PPJ). Only one monomer is shown for clarity, and the C␣ backbone of UQCRC1 is colored in red, UQCRC2 is shown in blue, CYTB in green, UQCRFS1
is orange, CYC1 is brown, and UQCRH is cyan. Stigmatellin (Stg) and antimycin (Ant) are shown as blue wire models bound at the quinol oxidation and
reduction sites, respectively. The structures used for cytochrome c (PDB ID 2B4Z) and for complex IV (PDB ID 1V54) were both from bovine heart. COX1
is colored green in both monomers, COX2 is blue, COX4 is red, COX5A is orange, COX5B is magenta, COX5C is pink, and COX6A, which crosses from one
COX1 to the other contacting Tyr304, is colored cyan in one monomer and brown in the other. Redox groups such as hemes, Cu atoms and a FeS center are
colored magenta. See main text for definitions of abbreviations.
(UQCRH). Genetic deletion (192) or removal (108) of this
subunit leads to a loss of binding of cytochrome c to CYC1
depending on the ionic strength, and at least in yeast, to a
decrease in cytochrome c reductase activity of 50%. Crystal
structures show that this subunit does not contribute directly to
the cytochrome c binding pocket (116, 200), but other studies
show a perturbation of the CYC1 heme environment when
UQCRH is removed (106), and its abundance of acidic residues has been proposed to direct cytochrome c to its actual
binding site (200). Phosphorylation of UQCRH has been detected with PhosTag dye (6), and Ser89, located close to the
COOH-terminus, has been identified as a phosphorylation site
AJP-Heart Circ Physiol • doi:10.1152/ajpheart.00077.2012 • www.ajpheart.org
Downloaded from http://ajpheart.physiology.org/ by 10.220.32.246 on May 5, 2017
UQCRFS1: UQCRC2:
UQCRC1: S99
S74 T86
S107
S102
S87
S8 S88
S212
Y115
Y448
Y207 S226
Y459
S247
Y468
S368 S437
S473
UQCR10:
UQCRB:
S12
[S3]
Review
MITOCHONDRIA PROTEIN PHOSPHORYLATION
Cytochrome c and Complex IV Phosphorylation
In contrast to the highly speculative significance of phosphorylation in complexes I, II, and III, a much more solid body
of evidence has emerged concerning the role of this type of
modification in residues of cytochrome c and its oxidase,
complex IV. As summarized in Fig. 5, A and C, there are four
identified sites of phosphorylation in cytochrome c, which
include two Tyr, one Ser, and one Thr residues. Of the 20 sites
in complex IV subunits, four correspond to Tyr, three to Thr,
and 13 to Ser residues (Fig. 5D). Two tissue-specific Tyr
phosphorylation sites have been reported after purification of
bovine cytochrome c in the presence of unspecific phosphatase
inhibitors: Tyr49 in liver (229) and Tyr98 in heart (119). In
both cases, changes in the kinetics of complex IV were observed when using the (at least partially) phosphorylated cytochrome c, which were reverted by alkaline phosphatase
treatment. With the Tyr49 modified cytochrome c purified
from liver, a twofold decrease in the maximal rate of isolated
complex IV was observed (229), similar to what was observed
when mutating this residue to Glu to mimic phosphorylation
(159). This Glu49 cytochrome c also showed a lower redox
potential by 45 mV, had a 30% lower binding affinity to
cardiolipin, and was unable to catalyze cardiolipin oxidation or
induce caspase-3 activity, which are events linked to the
proapoptotic role of cytochrome c when released from the
inner mitochondrial membrane (159). The Tyr98-phosphorylated cytochrome c from heart generated sigmoidal kinetics in
isolated complex IV and exhibited a marked blue shift of the
695-nm spectral peak that reflects the interaction of the c heme
with surrounding Met residues (119), also suggesting that a
high fraction of cytochrome c molecules in the preparation
contained this modification. Both Tyr49 and Tyr98 are well
conserved residues, with the first forming a hydrogen bonding
to the heme propionate (see Fig. 5C), and the second one being
in contact with Lys-7, which is important for apoptosome
formation (119, 229). Two other phosphorylation sites, Thr29
and Ser48, have recently been reported in resting human
muscle (236), but no functional effect has been linked to these
sites, which probably exist phosphorylated at very low mole
fraction since they were detected only after high enrichment of
phosphopeptides. No studies have been reported in which
phosphorylated cytochrome c at any site is assayed as a
substrate of complex III, which would be relevant to determine
whether electron transfer at this segment of the respiratory
chain is also impaired just as has been postulated for complex
IV. In addition, because of the mobility of cytochrome c
between different cellular compartments, the location of the
kinase/phosphatase system responsible for its phosphorylation
remains unclear.
Cytochrome c oxidase (complex IV) crystallizes as a dimer
of 13 subunits in mammalians (212). Subunit I [cytochrome c
oxidase (COX) 1] is the functional and structural core of this
enzyme, containing two a-type hemes, as well as the CuB atom
that constitutes, together with heme a3, the binuclear metal
center where reduction of oxygen to water occurs. Three
phosphorylation sites have been identified in COX1 through
MS of the isolated subunit or from purified complex IV.
Ser115 and Ser116 were identified in rabbit mitochondria after
ischemia and reperfusion of isolated hearts (65), with phosphorylation at these two residues nearly undetectable when an
AJP-Heart Circ Physiol • doi:10.1152/ajpheart.00077.2012 • www.ajpheart.org
Downloaded from http://ajpheart.physiology.org/ by 10.220.32.246 on May 5, 2017
in pig heart mitochondria (24). However, this residue is not
well conserved among species, so the functional relevance of
phosphorylation in this subunit remains to be established.
CYTB constitutes the functional core of complex III with its
two ubiquinol/ubiquinone binding sites and its two b-type
hemes, where electron recycling coupled to proton uptake and
release takes place. The high hydrophobicity of this subunit
makes it difficult to identify by MS (90), and even its staining
in gels is poor. Nevertheless, a phosphorylation site (Tyr75)
has recently been reported in CYTB from human muscle (236).
Crystal structures show this residue to be in one of the few
solvent accessible regions of CYTB, which is mostly embedded in the membrane and surrounded by other subunits (96,
116, 228). Interestingly, as shown in Fig. 5B, a phosphate
molecule can be seen in contact with Tyr75 in at least one
high-resolution structure from the bovine heart enzyme (87),
suggesting a high mole fraction of phosphorylation at this site
under some conditions. This residue is highly conserved
among species and is only four residues away from the essential Arg79 residue, which constitutes the exit pathway for
protons derived from ubiquinol oxidation (90). It would therefore be interesting to determine in the future if phosphorylation
at this site has an effect on the activity of complex III.
Two other small subunits are phosphorylated in complex III.
Subunit 7, also called UQCRB, has been detected in pig heart
with ProQ and PhosTag (6, 83). However, the phosphorylation
site at Ser3 found in this subunit (24) is actually located in the
short import presequence that is cleaved from this subunit
before assembly into the complex, so other as yet unidentified
residues in the mature protein must be responsible for the
binding of phosphorylation-sensitive dyes. The function of this
subunit is more related to the early assembly steps of complex
III (232) and not to the actual electron transfer activity, consistent with its low phosphorylation turnover (6). Subunit 10 is
the smallest subunit of complex III and has been found to be
phosphorylated at Ser12 in various mouse tissues (94). This
subunit 10 is required for insertion of UQCRFS1 as the last
step in complex III assembly (232), so it can be considered
simply as a chaperone, and its phosphorylation as related only
to that function.
It is noteworthy that no specific kinases or phosphatases
have been associated to any of the reported phosphorylation
events at complex III, as is the case for most mitochondrial
proteins. Low-level phosphorylation at complex III was detected when blue native gels were incubated with ␥-[32P]ATP,
suggesting at least a limited autophosphorylation activity in
this complex (161). However, it remains to be determined
which of the sites reported, if any, are the targets of this
intrinsic kinase activity. The fact that the few phosphorylation
sites found in the catalytically relevant subunits of complex III
have been detected only with increasingly sensitive phosphopeptide enrichment techniques (176, 236) suggests a very low
proportion of phosphorylated complexes. Only the isoelectric
shift in UQCRFS1 reported in liver mitochondria upon induction of the membrane permeability transition (78) seems to
reflect a PTM of a large fraction of complex III, although no
Ca2⫹-sensitive site has been identified yet, which could reflect
a very labile phosphorylation or a completely different type of
PTM.
H951
Review
H952
MITOCHONDRIA PROTEIN PHOSPHORYLATION
such as P-His and P-Asp residues (110). This would agree with
the bacterial origin of COX1, which is still encoded in the
mitochondrial genome.
Subunit II (COX2) is the only other redox center containing
subunit of complex IV. Its binuclear CuA center receives
electrons from cytochrome c, which binds on the surface of the
COOH-terminal globular domain that extends into the mitochondrial intermembrane space (212). Of the two sites reported, only one (Ser126) has been recently identified as a
phosphorylation site in heart after titanium dioxide enrichment
(92). However, Ser126 is located away from the cytochrome c
binding interface, although it might contact COX4 when phosphorylated (see Fig. 5D). Nevertheless, the protein band where
COX2 comigrates with subunit III (COX3) appears to be
labeled by ␥-[32P]-ATP when complex IV is incubated with
protein kinase A (18), and is recognized by antibodies against
P-Tyr, P-Ser and P-Thr (79). The tyrosine kinase c-Src has
been shown to phosphorylate COX2 in vitro, and its activity in
cells results in immunodetection of P-Tyr residues in COX2
(and even in COX1 in mouse cells), together with a mild
stimulation of complex IV activity (139). However, no specific
phosphorylation sites were reported in these studies.
Subunit IV (COX4) is the protein with the most phosphorylation sites identified so far in complex IV. This subunit
traverses the membrane by a single transmembrane helix in
close contact with COX1 and also interacts with COX2
through its matrix and intermembrane space domains (212).
COX4 appears to mediate the allosteric effect of ATP on
complex IV activity by directly binding adenine nucleotides at
both its matrix and intermembrane space domain (100), and
expression of a second isoform in some tissues abolishes this
regulation (93). COX4 has also been identified as interacting
with protein kinase Cε upon activation and translocation of
this kinase to mitochondria in neonatal cardiac myocytes
from rat, resulting in 32P labeling of COX4 that was correlated with an increase in complex IV activity (148, 149),
although it was not identified whether phosphorylation occurred at the matrix or intermembrane domain of this
subunit. Given that PKCε has been implicated in cardiac
ischemic or hypoxic preconditioning (137), the proposed
regulation of COX activity by phosphorylation of COX4
stands out as an example of a functionally relevant modification linked to an experimental manipulation, even if,
unfortunately, the mole fraction of phosphorylation has not
been determined. COX4 (and COX1) were observed to
become less phosphorylated when rat liver mitochondria
were treated with inhibitors of a soluble adenylyl cyclase
that is claimed to be present in the mitochondrial matrix and
that is activated by bicarbonate (2). However, the results of
these studies have not addressed the specific sites that
undergo phosphorylation or dephosphorylation. We have
also observed COX4 labeling by ␥-[32P]ATP in situ in blue
native gels where no kinases were detected by MS (161),
suggesting an intrinsic complex IV phosphorylation activity
acting on this and other subunits.
Phosphorylation of Tyr33 in COX4 was recently identified
after purification of complex IV from bovine liver in the
presence of phosphatase inhibitors and calcium chelators
(120), but its identification required enrichment of phosphopeptides, implying a low mole fraction of phosphorylated
Tyr33, or, alternatively, the high lability reported for Tyr
AJP-Heart Circ Physiol • doi:10.1152/ajpheart.00077.2012 • www.ajpheart.org
Downloaded from http://ajpheart.physiology.org/ by 10.220.32.246 on May 5, 2017
inhibitor of protein kinase A was administered during ischemia. No activity assay of complex IV was reported in this
study, although the complex IV levels were found to decrease
significantly because of the half-hour ischemic treatment,
which suggests that phosphorylation at Ser115 and Ser116
could be more related to protein degradation or turnover than
to regulation of electron flow. Nevertheless, Ser115 has been
proposed to be part of the exit channel for protons pumped
through COX1 during normal enzymatic activity (212), and
both Ser115 and Ser116 are well conserved and exposed to the
solvent in the intermembrane space of mitochondria (Fig. 5D),
making them susceptible to the kinase/phosphatase system well
characterized in this compartment (see Mitochondrial Matrix
Protein Kinases and Matrix Protein Phosphatases).
In bovine liver, isolation of mitochondria in the presence of
a phosphodiesterase inhibitor to generate high cAMP levels
resulted in the identification of Tyr304 as a phosphorylation
site in COX1 (120). The same residue was found to be
phosphorylated when bovine or mouse liver homogenates were
treated with TNF␣ before isolation of complex IV (183).
Tyr304 phosphorylation correlated with pronounced sigmoidal
kinetics and inhibition of the maximal rate of oxygen consumption by complex IV when varying the concentration of cytochrome c. This well-conserved residue could be accessible
from the intermembrane space if rotated out and has been
postulated to be part of an adenine nucleotide binding site
based on the crystal structure that shows cholate bound at this
location, which is assumed to resemble ADP (212). Tyr304 is
also in contact with subunit VIa of the opposite complex IV
monomer, which establishes interactions with its own COX1
subunit (Fig. 5D), thereby creating a possible communication
pathway for intermonomeric conformational changes that
could be responsible for the allosteric regulation by ATP (118).
Interestingly, the structure and sequence of the region surrounding Tyr304 is similar to that of the Tyr98 residue of
cytochrome c that has been shown to be phosphorylated in
bovine heart, suggesting that the same kinase/phosphatase
could target both COX1 and cytochrome c in the intermembrane space of mitochondria (119). However, it is not clear
what other residues in COX1 might be phosphorylated to
regulate the activity of complex IV. Because of its high
hydrophobicity, protein coverage of COX1 by MS has been
reported to be only of 6 –33% (65, 118, 183), implying that
most of the protein was not surveyed. Furthermore, antibodies
against P-Ser and P-Thr residues bind to COX1 after incubating isolated complex IV with protein kinase A, cAMP, and
ATP, conditions that also increase the sigmoidicity of complex
IV kinetics and lower the maximal rate in a phosphatase- and
Ca2⫹-sensitive manner (79). Radioactive labeling of COX1
under these same conditions was also observed in the presence
of ␥-[32P]ATP (18). However, in vivo phosphorylation of
complex IV was only detected when rat heart mitochondria
were rapidly isolated with phosphatase inhibitors and complex
IV separated by blue native electrophoresis instead of the usual
long chromatographic procedure (79). We have also recently
reported that bacterial lambda phosphatase activates ATPinhibited complex IV in blue native gels (161). These characteristics of COX1 phosphorylation suggests that several other
sites might have been missed in previous studies and that the
properties of its kinase/phosphatase regulatory system might
resemble those found in bacteria, which involve labile sites
Review
MITOCHONDRIA PROTEIN PHOSPHORYLATION
has been detected by labeling with ␥-[32P]ATP in blue native
gels from pig and rat heart (161). COX6B interacts with COX1
and COX2 on the intermembrane space (212) and is reported to
slow the rate of complex IV and to modify the affinity for
cytochrome c (218). Phosphorylation has also been reported at
Ser17 in subunit VIIc (COX7C) in human resting muscle (236)
after phosphopeptide enrichment, which is the very NH2terminal residue of COX7C, and is probably located very close
to f-Met 1 of COX1 in the matrix, although its position is not
defined in the structure (212). However, since the role of
COX6C and COX7C in complex IV activity and/or assembly
is obscure, very little can be inferred about possible effects of
phosphorylation at these sites or whether they are physiologically relevant at all.
In summary, the functional correlation in cytochrome c and
complex IV with specific phosphorylation sites constitutes
some of the strongest data in support of a role of protein
phosphorylation in the regulation of the cytochrome chain. The
potential involvement of cAMP-sensitive kinases, PKA, and
PKCε in complex IV is also compelling; however, the precise
localization of the kinase activity that interacts with complex
IV remains unclear and could simply occur under certain
conditions that favor translocation of kinases into the intermembrane space. Furthermore, unknown or low confidence
kinase recognition motifs characterize the amino acid sequence
where phosphorylation sites are located in cytochrome c and
complex IV (92), which is also the case for the majority of
other sites reported in the rest of the mitochondrial proteome
(see Mitochondrial Matrix Protein Kinases). Thus the actual
kinase and phosphatase network regulating these sites is still an
open question.
Complex V Phosphorylation
ATP synthesis is accomplished by the rotational catalysis of
the complex V enzyme, comprised by the extrinsic F1 region
and the membrane-embedded Fo sector. A total of 67 different
phosphorylation sites have been reported in 12 of the 16
subunits of mammalian complex V, in addition to five more
sites in the inhibitor protein of F1 (IF1) and factor B, which are
proteins that associate with complex V to modify its activity.
Almost two-thirds of these sites are found in the F1 domain,
mainly in the hexamer formed by the ␣- and ␤-subunits. Of the
remaining sites, 16 are found in four subunits (OSCP, b, d, and
F6) that, although traditionally classified as part of the Fo
region, actually constitute a membrane-anchored stator that
extends to the tip of F1 to hold it in place against the rotation
of a central stalk composed of a ring of c-subunits in Fo and of
subunits-␥, -␦, and -ε in F1. Only six phosphorylation sites
have been reported in four of the eight subunits that constitute
the hydrophobic core of the Fo sector. These sites are schematically presented in Fig. 6, A–D.
In the F1 region, subunit-␣ is labeled by both PhosTag and
ProQ dyes in pig heart and by PhosTag in pig liver (6, 83). It
is also radioactively labeled by incubating pig heart (but not
liver) mitochondria with 32P (6), although part of this can be
due to metabolite association as indicated by the observation
that some labeling occurs when incubating the purified complex V or blue native gel strips with ␣-[32P]ATP (161). Our
own evaluation shows that ADP might be one of the metabolites tightly associated with complex V (R. Covian, R. L.
AJP-Heart Circ Physiol • doi:10.1152/ajpheart.00077.2012 • www.ajpheart.org
Downloaded from http://ajpheart.physiology.org/ by 10.220.32.246 on May 5, 2017
phosphorylation in general in complex IV. Tyr33 is located in
the matrix side of the membrane, close to the adenine nucleotide binding site in the matrix proposed by comparative
modeling of the isoforms 1 and 2, the last of which does not
show ATP allosteric inhibition of complex IV when expressed
(93). Inspection of the crystal structure (see Fig. 5D) also
shows that Tyr33 is in contact with helix XII of COX1, not far
from the entrance of the channels through which protons enter
COX1 to be pumped or consumed by the reduction of oxygen
to water (212). However, no studies have shown whether this
site has an effect on complex IV activity.
The matrix located Ser58 was also identified recently as
phosphorylation site for COX4 in bovine heart after fluoride
treatment of mitochondria to inhibit proteases and phosphatases, followed by phosphopeptide enrichment (79). However,
phosphorylation at this site was related to a slight inhibitory
effect of added ATP in the isolated complex IV, without
induction of sigmoidal kinetics observed with other phosphorylation sites. Thr74 phosphorylation was detected in rabbit
heart after ischemia and reperfusion (65); however, this residue
(see Fig. 5D) is located at the tip of the matrix domain of
COX4, far from the interface with COX1 and somewhat distant
from the proposed adenine nucleotide binding site, so its
functional significance, if any, is unknown.
Subunit Va of complex IV (COX5A) interacts with COX4
and to a lesser extent with COX2 on the matrix side of the
membrane (212). It has been reported to abolish the allosteric
effect of ATP on complex IV kinetics in the presence of
thyroid hormones (8) and is readily detected by phosphospecific dyes and by ␥-[32P]ATP labeling (6, 83, 161). COX5A is
located in contact with the matrix domain of COX4, close to
the proposed regulatory ATP binding site (93), and also interacts with the single matrix loop of COX2 (see Fig. 5D). Ser47
and Thr78 were identified as phosphorylation sites in complex
IV from fresh bovine heart isolated in the presence of phosphatase inhibitors and after phosphopeptide enrichment (79),
suggesting a low mole fraction (as was the case with Ser58 in
COX4), which would explain the lack of significant effects on
enzymatic activity.
Subunit Vb (COX5B) is also located in the matrix in contact
with COX1, COX3, COX4, and COX5A. Its phosphorylation
has been detected with PhosTag (6), by protein kinase A in
vitro (18), in blue native gels by an intrinsic phosphorylation
activity in complex IV (161), and also by P-Ser antibodies
(167). Phosphorylation of Ser71 in COX5B was found after
ischemia and reperfusion of rabbit hearts followed by blue
native electrophoresis of mitochondria without enrichment of
phosphopeptides and was sensitive to an inhibitor of protein
kinase A (65). However, no attempt to quantify complex IV
activity was made to evaluate the functional relevance of this
phosphorylation. Ser124 phosphorylation was reported in pig
heart COX5B after phosphopeptide enrichment (24), although
most other mammals have a Pro residue at this position. Both
Ser71 and Ser124 are located away from the contacting surfaces of COX5B with other complex IV subunits and exposed
to the solvent (see Fig. 5D), so their functional effect in
enzyme catalysis is probably very weak.
Three more subunits in complex IV have phosphorylation
sites for which functional significance and mole fraction are
unknown. Sites in subunit VIb (COX6B) have recently been
reported in mouse, but not in heart or muscle (94), although it
H953
Review
H954
MITOCHONDRIA PROTEIN PHOSPHORYLATION
A
INTERMEMBRANE SPACE
Complex V
c: [S63]
α:
IF 1
S63
T30
T32
T432
d:
Y152
Y150
δ:
Y9
97
T475
S419 ANP
β:
S529
S528
C
Y395
S521
MATRIX
S106
S30
Stator
F6:
1T475 S316
S108
Y395
S57
T312
Y361
S64
(S263/
T213
S65
T262)
T318
S231
b:
Y165
Y269
Y230
T240
S128
S226
T140
T110
T229
T116
T107
OSCP:
[S25] S106
S145
[S23]
Y46
[T21]
S155 S166
Y35
F
T45
T225
S315
T368
T213
S184
T236
Y361 S231
S316 T312
T240 Y269S263
T262
S76
S100
S106
T110
T140
Y230
T116
T107
S99
S106 S128
D
S63
Y77
Y97
Y69
S146
Y165
S108
S39
T45
ANP
T46T48
Y35
S52 S53
S65
S64
S57
S226
S145
S63
S65
S63
Y46
T229
S166
S63
S63
S155
Fig. 6. Reported sites of phosphorylation in complex V. In the schematic representation of phosphorylation sites (A), residues in the indicated regions and subunits
are shown in white if no functional effect has been reported and their location in the structure suggests no impact on activity. Sites colored in yellow have no
known function but are located in a region that might influence activity according to available high-resolution structures. Site that could influence function based
on structure and that have been found to change its phosphorylation level in different physiological states are shown in orange. Sites that are located in the
presequence and absent in the assembled complex are shown in brackets. The location of several of the reported sites in the F1 region are shown in the crystal
structure (PDB ID 2JDI) where only the ␣- and ␤-subunits in the “closed” conformation with adenosine 5=-(␤,␥-imino) triphosphate (ANP) in the active site are
shown (B). Subunit-␣ is colored blue, subunit-␤ is red, and subunit-␥ is cyan. Other phosphorylation sites (C) are shown in another crystal structure (PDB ID
2WSS) that shows part of the stator attached to one of the ␣-subunits (blue). The ␤-subunit closer to the stator is also shown (magenta), with subunit-␥ in the
center of F1 (cyan) and subunit-␦ (pink) at its foot. In the stator region, OSCP is colored brown, subunit-b is orange, F6 is blue, and subunit d is yellow. The
IF1 protein with its two reported phosphorylation sites is also shown attached to F1 (D, top), based on a different crystal structure (PDB ID 1OHH). Only
the ␣ (green)- and ␤ (red)-subunits with which IF1 (blue) establishes contacts are shown, together with subunit-␥ (cyan). The tetrameric (inactive) IF1 is shown
(D, bottom) with Ser63 highlighted in one of the oligomerization interfaces (PDB ID 1GMJ). See main text for definitions of abbreviations.
Levine, and R. S. Balaban, unpublished observations). Subunit-␣ contains 17 reported phosphorylation sites that correspond only to Ser or Thr residues. A cluster of seven sites is
located near the NH2-terminal region of the mature protein,
with Thr46 establishing contact with the OSCP subunit that
forms the tip of the stator (174). This residue was detected in
isolated mouse heart mitochondria (58). Ser53 establishes
contacts with residues from the adjacent ␤-subunit and has
been found to be phosphorylated in human resting muscle
(236). In pig heart, the phosphorylation level of this residue
AJP-Heart Circ Physiol • doi:10.1152/ajpheart.00077.2012 • www.ajpheart.org
Downloaded from http://ajpheart.physiology.org/ by 10.220.32.246 on May 5, 2017
S451
S39 S521
S315
T432
T236
S419
S106
S184
S100
T225
S99
S76
S65
S53
S52
T46
T48
S451
f:
FO
γ: (Y69 /77)
S146
S108
T45
S108
e:
Y30
Y32
A6L:
S63
B:
T154
S162
T167
B
Review
MITOCHONDRIA PROTEIN PHOSPHORYLATION
last of these residues, their location in the structure does not
seem to support a functional role, as shown in a phosphomimetic mutagenic study in yeast that showed primarily assembly
defects with some weak direct enzymatic activity changes
(101). Thr368 is in contact with the ␥-subunit inside the F1
hexamer, but its poor accessibility suggests that its phosphorylation does not occur in the assembled complex V.
Only four phosphorylation sites in subunit-␤ stand out as
potentially regulatory for enzymatic activity based on available
structures because of being part of the substrate binding pocket
(25). As shown in Fig. 6B, Tyr395 and Thr475 are in contact
with the adenine ring, whereas Tyr361 is close to the ␥-phosphate of ATP and Thr213 coordinates the Mg ion of the
substrate complex. Interestingly, all four residues were found
as phosphorylation sites in human skeletal muscle, and both
Thr213 and Tyr361 were shown to have a 30% higher phosphorylation in biopsies from obese individuals (82). Tyr361
also showed a 50% increase in its phosphorylation level upon
insulin administration in healthy individuals but not in diabetics. However, it is not stated in this study what fraction of sites
was phosphorylated at basal levels; an increase of 50% from a
fraction of a percent would be meaningless in terms of modifying the overall rate of oxidative phosphorylation. The fact
that most sites were detected only after a targeted MS/MS
approach indicates that the abundance of phosphopeptides was
very low. Since the effects of obesity and diabetes were also
related to a decrease in protein levels of oxidative phosphorylation enzymes, including complex V (82), it can be speculated
that the phosphorylation of residues in the active site of the
␤-subunit could be a mechanism to prevent ATP hydrolysis by
free F1 that has dissociated from Fo and is in the process of
being disassembled for proteolysis. It is also troubling that
none of the ␤-subunit sites label with 32P in intact heart
mitochondria (5, 6) and the amount of phosphorylation does
not change with a wide range of perturbations (24), suggesting
that these sites are not dynamically regulated.
Subunit-␥ constitutes the axle that transmits the rotation
energy of the ring of c-subunits in Fo to the ␣-␤-hexamer to
induce the conformational changes that modify the catalytic
sites where adenine nucleotides and phosphate bind (203). It is
noteworthy that this subunit is labeled by PhosTag and ProQ
dyes in pig heart, but not in liver (6, 83), and that it is labeled
with 32P in a calcium-sensitive manner that depends on mitochondrial energization which cannot be explained by metabolite association or intrinsic kinase activity in blue native gels
(6, 161), suggesting that an as yet unidentified soluble kinase is
responsible for phosphorylation of ␥ in the mitochondrial
matrix. Four phosphorylation sites have been identified in this
subunit, two of which could in theory impede the rotation of ␥
relative to the ␣-␤-hexamer based on the structural information
available (25, 203). Thr45 is inside the F1 hexamer in close
contact with one of the ␤-subunits (see Fig. 6, B and C);
however, the inaccessibility of this residue from the solvent
and its being found only in a human cancer cell line (151),
suggest that it is not a physiologically relevant site, perhaps
being phosphorylated abnormally when ␥ is disassembled from
the rest of the enzyme. The other residue, Ser108, is less than
7 Å away from the interface with the hexamer (Fig. 6, B and
C), so an attached phosphate in an extended conformation of
the side chain could make contact with the ␣- or ␤-subunits.
This site, however, has only been reported in mouse kidney
AJP-Heart Circ Physiol • doi:10.1152/ajpheart.00077.2012 • www.ajpheart.org
Downloaded from http://ajpheart.physiology.org/ by 10.220.32.246 on May 5, 2017
increased threefold when mitochondria were energized with
substrates and ATP (24). Ser65 phosphorylation was found in
mouse heart (94) and in human muscle (236), but its position
in a solvent exposed loop far away from any other subunit
makes it unlikely to affect catalysis (34). Ser76 is in close
contact with subunit-␤, and its phosphorylation has been detected in pig heart (24) and mouse heart (94, 213, 214, 222).
However, the functional impact of phosphorylation even at
these sites that interact with the OSCP or ␤-subunits is unclear,
given that little conformational changes during the catalytic
cycle of complex V is observed in this upper region of the F1
hexamer (84). The same can be said of phosphorylation at
Ser106, found in mouse heart (58, 94), given that this residue
is away from any subunit interface. Thr236 and Ser521, found
to be phosphorylated in mouse heart (94, 222, 231), also fall in
the category of functionally irrelevant residues because of their
position and poor conservation across species.
Only four residues where phosphorylation has been reported
in subunit-␣ are potentially relevant in a functional sense from
a structural point of view (see Fig. 6B). Ser184, Ser419, and
Thr432 move significantly during the closure of the catalytic
site as the ␣-␤ interface cycles between the empty, loose, and
tight conformations (203) and are also in the path of substrate
entry and exit. Phosphorylation has been found in mouse brown
fat and heart for Ser184 and Ser419 (94) and during mouse
osteoblast differentiation in the case of Thr432 (105). Another
phosphorylation site identified in mouse brain, Ser451, establishes a transient hydrogen bond via a water molecule with an
Arg residue at the foot of the ␥-subunit that is visible in crystal
structures when the ␣-␤ interface is in the empty conformation
(25). Given that this is a region that changes during rotation of
␥, a phosphate group attached to this Ser could prevent this
movement either by strengthening the bond between ␣ and ␥ or
by introducing a steric hindrance to the relative displacement
of the two subunits. However, the lack of data on the mole
fraction of phosphorylation at any of these structurally relevant
residues makes any discussion on their functional significance
speculative.
Subunit-␤ is labeled by the PhosTag dye and by incubation
of mitochondria with 32P in pig heart and liver (6) and also by
ProQ in heart (83). However, most radioactive labeling can be
attributed to tight association of adenine nucleotides and phosphate to the binding site, which are revealed as shifts in
two-dimensional gels (6). Of the 23 phosphorylation sites
reported in subunit-␤, those reported in mouse heart or human
skeletal muscle are Ser128 (58, 151), Tyr230 (82), and the
COOH-terminal residues Ser528 and Ser529 (94, 236). However, these sites seem to have little significance based on their
location on the surface of the ␤-subunit and away from the
catalytic site (Fig. 6B). Tyr269, which was found in skeletal
muscle (82), is in close contact with the adjacent ␣-subunit, but
in the upper half of F1, where little conformational changes
occur during catalysis (25). Thr312 and Ser316, both detected
as phosphorylation sites in mouse heart (94) and the first also
in human muscle (82), are away from any subunit interface and
buried inside the hexamer, which raises questions on how they
can be phosphorylated in the assembled complex. Thr312 has
also been reported in rabbit ventricular myocytes subjected to
pharmacological preconditioning by being exposed to adenosine (9), along with four other sites not reported elsewhere:
Ser106, Thr107, Thr262 or Ser263, and Thr368. Except for the
H955
Review
H956
MITOCHONDRIA PROTEIN PHOSPHORYLATION
known. None of these sites has been found in heart or muscle
proteomic screens.
Other accessory polypeptides that affect activity of complex
V and where phosphorylation sites have been found are factor
B and the inhibitor protein (IF1). The first has been implicated
in enhancing the coupling of proton movement to ATP synthesis by an as yet unidentified mechanism (16). Three phosphorylation sites (Thr154, Ser162, and Thr167) have been
found in this protein, but only in human lymphoma cells (131),
so their physiological role is unlikely. IF1 has been proposed to
be important to prevent hydrolysis of ATP under ischemic
conditions in which mitochondrial membrane potential collapses, binding to the lower region of F1 and preventing the
conformational changes required for catalysis in the ␣- and
␤-subunits (71). IF1 inhibition takes place under slightly acidic
conditions in which its inactive tetrameric form dissociates into
a dimer that binds to two complex V molecules simultaneously
(30, 31, 70). Two phosphorylation sites, Ser39 and Ser63, have
been reported in this protein. The first site was found in human
skeletal muscle (236) and is located inside F1 in the dimeric
form (see Fig. 6D), in contact with subunit-␤ (15), so in theory
it could be envisioned that phosphorylation at this site might
interfere with binding. However, it is not universally conserved, being substituted by Ala in ungulates. The second site
was found in cultured cells transfected with an angiotensin
receptor (40) and lies on the interface of tetramer formation
(32). However, this residue is even less conserved, changing to
Thr in rodents and to Ala in ungulates, which precludes the
assignment of a physiological role to this site.
In summary, many phosphorylation sites have been identified in complex V; however, the functional significance of
these events remains obscure. Furthermore, their kinase/phosphatase system is poorly defined in either the intermembrane
space or the matrix.
Mitochondrial Matrix Protein Kinases
From the discussion above it is clear that phosphorylated
proteins, at some level, are widespread within the mitochondrial matrix. A few years ago, protein phosphorylation was
believed to occur only on PDH or BCKDH; however, the
presence of this potential regulatory PTM is much more widespread. The first issue to address is where did the protein
phosphorylation occur? Since most of the proteins of the
mitochondria come from nuclear DNA, the majority of proteins must traverse the cytosol as well as be processed through
a complex protein trafficking pathway (the TOM-TIM complex) across the outer membrane, the intermembrane space,
and the remarkably “tight” inner membrane (145). Thus protein phosphorylations could occur along this transport pathway, and if appropriate phosphatases are not present in the
matrix, these phosphorylations could persist indefinitely. A
screen of the protein phosphorylations and the theoretical
kinases interactions was performed by Cui et al. (47), who
found that over 52 kinases could be responsible for this profile.
However, the vast majority of these kinases lack a mitochondrial targeting sequence (MTS), implying they are not located
in the matrix, unless novel protein transport mechanisms are in
play. This is also the case in yeast as reviewed by Lubomir
(127) where it was suggested that “mitochondria, like obligatory intracellular bacterial parasites, are no longer dependent
AJP-Heart Circ Physiol • doi:10.1152/ajpheart.00077.2012 • www.ajpheart.org
Downloaded from http://ajpheart.physiology.org/ by 10.220.32.246 on May 5, 2017
(94), suggesting it is unrelated to the phosphospecific labeling
observed in heart (6, 83). Another reported phosphopeptide
from bovine heart mitochondria is phosphorylated on either
Tyr69 or Tyr77 (63), both of which are over 30 Å away from
the interface with the F1 hexamer (Fig. 6C), which in addition
to the low abundance of the corresponding phosphopeptide
(63) argues against its functional significance in terms of
affecting oxidative phosphorylation rates. The most commonly
found phosphorylation site in subunit-␥ is Ser146, which has
been reported in pig heart (24), where its relative abundance
did not change significantly upon calcium stimulation of mitochondrial respiration. This residue is ⬃ 11 Å from the
interface with ␣ and ␤, much closer than the Tyr residues in the
phosphopeptide found in bovine heart (63), but still too far
away to propose a direct interference with rotation by the
simple addition of a phosphate group (see Fig. 6C).
Phosphorylation of the ␦-subunit, which is associated (and
rotates with) ␥ at the interface with the c ring of Fo (203), has
been pinpointed to its only Tyr residue (112, 233). However, in
a structural sense (Fig. 6C), it is not evident how modification
of this residue could affect the rotational catalysis, given that
there is no interaction of the ␦-subunit with other regions of
complex V that do not rotate. Nevertheless, phosphorylation at
this site has been linked to PDGF signaling in mouse cortical
neurons (233), as well as in fibroblasts and kidney cells (112),
although its role, if any, remains obscure.
The stator region of complex V is highly exposed to the
matrix. Its function seems to be much less dynamic that the F1
sector: simply holding the latter in place relative to the rotation
of ␥, although some discussion has taken place regarding the
possible effects of flexibility in the stator region during the
catalytic cycle (174, 215). Therefore, the phosphorylation of
stator subunits does not seem to have an intuitively relevant
role in modifying complex V activity, unless it is assumed that
phosphorylation takes place on the stator sufficiently close to
the middle or lower regions of the F1 hexamer so as to obstruct
the outward displacements of ␣ and ␤ as rotation of ␥ induces the active sites to open and close sequentially. Two of
the identified phosphorylation sites that are somewhat close to
this region are Ser30 and Ser106 of the d-subunit, detected,
respectively, in dog heart (187) and in human skeletal muscle.
Some of these sites might also be close to the interface between
complex V dimers that has been proposed to be important for
oligomerization and subsequent formation of tubular mitochondrial cristae (43), but a role of phosphorylation in this kind
of phenomenon is highly speculative at this point.
All other phosphorylation sites identified in the stator region
are close to the tip of F1 (see Fig. 6C) where contact is made
with the hexamer (174), including Ser155, found in human
muscle (236), and Thr145, identified in pig heart, both located
in OSCP. Ser226 and Thr229 of subunit-b, and Ser64 and
Ser65 (which are not conserved in other mammals) of the F6
subunit, all found in skeletal muscle (236), are also close to this
upper region where little functional effects can be expected. It
is not known whether any of the above-mentioned sites are
responsible for the labeling of the OSCP and d-subunits with
PhosTag, ProQ, or 32P detected in heart mitochondria (6, 83,
161). Little can be said about the functional significance of the
five phosphorylation sites located in mature Fo subunits because their structure and relation to other subunits is not
Review
MITOCHONDRIA PROTEIN PHOSPHORYLATION
those occurring in the matrix space. Thus the difference between mitochondrial association and true matrix localization
has been difficult to prove.
One of the most extensively studied systems associated with
the mitochondria is a cAMP-dependent PKA system that was
found in isolated membrane fractions by Papa’s group (185,
209) well before the recent developments in screening methodology. Subsequently, it has been demonstrated that PKA can
phosphorylate many mitochondrial proteins when combined in
vitro (17, 36, 136); however, the evidence that PKA is within
the matrix space is still controversial. For example, one of the
early studies on the topology of PKA evaluated cAMP-sensitive protein phosphorylation in intact deenergized oligomycintreated mitochondria using ␥[32P]ATP (186). The authors
found that the addition of the purified catalytic subunit of PKA
was more effective than the endogenous PKA activity at
phosphorylating the same protein fractions, suggesting that
PKA outside of the matrix could be influencing these results.
The authors suggested that this was due to the same molecular
weight proteins being fortuitously phosphorylated on the outer
surface of the inner membrane; however, the specific sites were
not determined in these early studies. An inhibitor of the
adenylate translocase blocked much of the native phosphorylation, but not the exogenously added PKA. This latter result
suggested that a cAMP-sensitive matrix pool of protein phosphorylation existed; however, the specific activity of the ATP
was not determined in these complicated experiments with
compromised mitochondria.
Electron microscopy immune-localization of gold particles
has been used to confirm the location of PKA on the mitochondrion inner membrane (184, 193); however, whether PKA
actually has access to the matrix is still controversial. Indeed,
recent reviews have models locating this enzyme in the intermembrane space (66), whereas others propose a matrix-PKA
(155). Most recently, Means et al. (134) provided convincing
evidence that one of prominent substrates for PKA is the
intermembrane helix protein ChChd3, supporting the notion of
its localization outside the matrix. Again, the evidence that
PKA plays a role in mitochondrial intermembrane space phosphorylation reactions and that a complex “transduceosome”
seems to exist in this space is compelling [reviewed in Feliciello et al. (66)]. However, the evidence that PKA is translocated into the matrix is still not as well developed. One of the
major issues is how this translocation can occur for this and
other kinases that lack a MTS. Sardanelli et al. (184) suggested
that dissociated PKA may diffuse through cellular membranes
just as it does across the nucleus (75); however, the nuclear
membrane is permeable to proteins up to ⬃40 kDa, making it
very different from the remarkably tight inner membrane of the
mitochondria.
To date, the prediction of Lubomir (127) that the cytoplasmic kinases effects seem to be limited to the region of the outer
membrane and intermembrane space is clearly correct for the
large majority of conventional protein kinases. How these
reactions influence the matrix function is a continuing area of
investigation. However, it is still unclear how 32P protein
phosphorylation is occurring in isolated mitochondria at numerous sites with very few demonstrated matrix kinases. To
illustrate this point, a computational search of classical kinase
recognition motifs in MOPC subunits that face to the matrix,
except complex IV, where a search yielding similar results has
AJP-Heart Circ Physiol • doi:10.1152/ajpheart.00077.2012 • www.ajpheart.org
Downloaded from http://ajpheart.physiology.org/ by 10.220.32.246 on May 5, 2017
on signaling mechanisms mediated by protein kinases residing
within the mitochondria. Instead, the nucleo-mitochondrial
communication system requiring protein phosphorylation may
be predominantly regulated by protein kinases, which are
cytosolic and/or anchored to the outer mitochondrial membrane.” We would modify this quote to include the intermembrane space and outer regions of the inner membrane based on
the observations presented below. Since many of these kinases
have not been found in the matrix or possess a MTS, many of
the phosphorylation sites in mitochondrial proteins might be
the result of cytosolic kinase reactions that take place before
being imported, as suggested by Lubomir. Cytosolic or intermembrane space protein phosphorylation has been demonstrated to influence the import of some proteins located both in
the cytosol and matrix such as mitochondrial-targeted cytochrome P-450, family 1, subfamily A, polypeptide 1 (51);
glutathione S-transferase (172); and complex 1 NDUFS4 (54),
supporting this view. We also found many examples of protein
phosphorylation in targeting (MTS) portions of the protein,
which is consistent with a possible role in protein translocation.
However, the extent of proteins that are transported into the
matrix with phosphorylations from the cytosol is still poorly
defined.
Besides the well-characterized PDH and BCKDH kinase
phosphatase system, the best evidence for the existence of
matrix kinase activity is protein phosphorylation with 32P in
isolated mitochondria. As originally shown by Sale and Randle
(179), the addition of 32P to energized mitochondria can be
demonstrated to directly phosphorylate PDH in the steady
state, suggesting a rapid turnover of these regulatory sites. This
high turnover confirmed the existence of an active kinase and
phosphatase in the matrix. Subsequently, this approach has
been used extensively in intact potato mitochondria (201),
isolated potato mitochondria membranes (164, 204), porcine
heart mitochondria (5, 6, 83, 161), as well as in blue native gel
resolved mitochondrial complexes (161). These studies demonstrate extensive 32P incorporation into matrix-exclusive proteins once proper controls are run for weak and tight metabolite
associations (5). Since 32P incorporation requires these sites to
turn over, that is, to dephosphorylate and rephosphorylate,
these data imply that there is a wide range of matrix kinase and
phosphatase reactions occurring that do not require cytosolic
elements, and this phosphorylation process is widely conserved
phylogenetically.
What are these matrix kinase reactions? There is no question
that cytosolic kinases are associated with purified mitochondrial preparations and that these kinases can be recruited to this
subcellular fraction under different experimental conditions
(23, 104, 115, 129, 139, 182). These later observations have
been supported by the association of several kinase anchoring
proteins including AKAP (86), SKIP (134), PICK1 (216),
Grb10 (143), Sab (221), OMP25 (44), and Rab32 (4). Thus, at
least on the outer membrane and outer surface of the inner
membrane, there is a growing body of evidence that many
kinases are sequestered on the mitochondrion at different times
to accomplish a variety of intriguing signaling chores. These
intermembrane space kinases include PINK (41), PKA (4,
134), Src kinase (182), and PKC␦ (3). However, how these
kinases can influence matrix phosphorylation events remains
elusive. One of the difficulties in this process is differentiating
kinase activities on the surface of the mitochondrion from
H957
Review
H958
MITOCHONDRIA PROTEIN PHOSPHORYLATION
B
Complex I
50
45
40
35
30
25
20
15
10
5
0
CK2
High
G k3b
Gsk3b
Gsk3b
CK2
PKA
p85 SH2
CK2
Medium
50
40
High
30
Medium
20
Low
10
Low
Complex V
18
5
4
PKCδ
3
High
Medium
2
Clk2
Low
1/Percentile
6
PKA
16
Akt
14
12
10
8
High
PKCζ
Medium
6
4
Akt
2
Low
AMPK
0
0
Fig. 7. Kinase recognition motifs in selected complex I (A), II (B), III (C), and V (D) subunits located in the mitochondrial matrix. The Scansite 2 program (147)
was used to analyze the subunits that face the matrix in the indicated MOPC to identify kinase consensus motif sequences. The confidence level is related to
the reciprocal of the percentile score provided by the Scansite program in the following manner: high (red), ⬎5; medium (blue), and 1–5; low (green), 0.2–1.
The kinase with the highest score for each site is indicated above the medium and high confidence sites. Residues in brackets are located in the import presequence
and are thus not present in the assembled complex. Reported sites that were not recognized at the lowest confidence threshold are not shown. See main text for
definitions of abbreviations.
already been reported elsewhere (92), is shown in Fig. 7. Out
of the ⬎140 reported sites in these subunits, only 19 were
identified in silico with at least medium confidence as substrates of known kinases, and almost half of these were in the
string of Ser residues located in the NDUFV3 (isoform 2) subunit of complex I (see Fig. 7A). Interestingly, two of the sites
with high similarity to consensus recognition sequences are
located in the import presequence of ETFA (Fig. 7B) and the
␤-subunit of complex V (Fig. 7D), clearly indicating that
phosphorylation of these residues occurred in the cytosol. Only
a handful of sites with medium to high confidence kinase
motifs appear in the mature proteins, some of which are
thought to be important only for assembly of the complexes
such as the accessory subunits UQCRC1, UQCRC2, and
UQCR10 of complex III (Fig. 7C) and NDUFA7 of complex I
(Fig. 7A). The few sites that are located in catalytically relevant
subunits are either in structurally uninteresting regions of the
protein, such as Tyr365 of SDHA in complex II, Ser76 and
Thr236 of subunit-␣, and Ser146 of -␥ in complex V, or have
only been found in cancer cells, such as Tyr141 of NDUFS2 in
complex I and Ser89 of COX4 in complex IV (92, 151). No
common kinase activation pathway in the matrix can therefore
be discerned in mitochondrial matrix proteins that could support the assumption that oxidative phosphorylation is regulated
by cytosolic kinases that are translocated into the matrix.
Kinase dependent protein phosphorylation relies on the main
product of oxidative phosphorylation, ATP, thus the possibility
that [ATP] could regulate protein phosphorylation is feasible
and an intriguing feedback mechanism. However, the fact that
a metabolic homeostasis exists in the heart, maintaining the
[ATP] essentially constant over wide work ranges (11), suggests that [ATP] is not regulatory under normal physiological
conditions. However, [ATP] regulation of kinase activity during metabolic compromises could occur if the ATP drops
sufficiently. Most classical protein kinases have affinities of 10
to 100 ␮M for ATP, depending on the [Mg2⫹], well below the
resting millimolar concentration of ATP (195, 206). Based on
these classical protein kinase ATP affinity values, the [ATP]
would need to drop roughly an order of magnitude to influence
protein kinase reaction rates.
Matrix Protein Phosphatases
As is the case for the kinase system, the number of documented phosphatases in the matrix is much lower than those
found associated with a mitochondrial enriched fraction from
tissues [for example, see Hopper et al. (83)]. Again, the best
characterized are the PDHPs where the Ca2⫹-sensitive PDHP
1 is the major isoform in heart (85), generating the sensitivity
of PDH activity to Ca2⫹. The BCKDH phosphatase protein has
also been characterized (50). Recently, a serine/threonine phosphatase was identified in the matrix (PP2Cm), with highsequence homology with yeast Ptc4, which was shown to be
critical in cell survival, possibly responsible for the dephos-
AJP-Heart Circ Physiol • doi:10.1152/ajpheart.00077.2012 • www.ajpheart.org
Downloaded from http://ajpheart.physiology.org/ by 10.220.32.246 on May 5, 2017
D
Complex III
7
1/Percentile
AMPK
0
C
1
Complex II and ETF system
70
60
Gsk3b
1/Percentile
1/Percen
ntile
A
Review
MITOCHONDRIA PROTEIN PHOSPHORYLATION
Nonconventional Protein Phosphorylation
Nonconventional protein phosphorylations are events not
associated with conventional serine, threonine, and tyrosine
sites. Of particular interest are the protein phosphorylation
events associated with the bacterial system that includes histidine, arginine, aspartate, cysteine, and lysine. Many of these
phospho-amino acids have high energy phosphate bonds and
permit a phosphotransfer relay between sites without a specific
kinase. Recently, it has become appreciated that “remnants” of
this signaling systems are present in mammalian cells (99, 110,
117). These are remarkably different from the conventional
protein phosphorylation systems in numerous ways. The first is
that these sites are highly labile at acid pH, making many
conventional proteomic procedures miss them entirely. In addition, the actual phosphorylation reactions are often done via
autophosphorylation mechanisms not requiring a specialized
secondary kinase. Finally, due to the labile nature of these
events, no antibodies are available for detection.
The reasons for suspecting these reactions in the mitochondrion are severalfold. First, many of the bacterial systems
involved in protein synthesis, DNA structure, and processing
are very similar to the mitochondrial system (153), supporting
the notion that the mitochondria originated from some type of
symbiotic relationship early in the development of the eukaryotic cell (130). Thus it is not unreasonable to assume that the
bacterial signaling systems might also be conserved in the
mitochondrion. Interestingly, the bacterial two-component system is particularly suited to generate cellular signals to adapt to
extracellular events, much like the mitochondrion samples and
“serves” the cytosol. Beyond this rationalization, what is the
evidence that these events are occurring in the mitochondria?
First, it has been known since the 1960s that most of the 32P
associated with mitochondrial proteins is labile at acid pH
(122, 160, 168, 190). One of these acid labile sites was
identified to be phosphohistidine by several investigators (160,
168, 198). This acid labile phosphorylation pool is also modulated by Ca2⫹ (146), which is interesting given the proposed
role of Ca2⫹ in regulating mitochondrial energy conversion
(10). Estimates by Pressman (168) suggests that up to 99% of
the initial 32P protein association is acid labile, implying not
only a widespread distribution but also a significant fraction of
the total protein phosphorylation in the matrix. This observation was recently confirmed by Aponte et al. (5) in blue native
electrophoresis gels of mitochondrial proteins labeled in the
intact mitochondrion after 20 min, where most of the 32P
labeling was removed by warm acid washing. Nonconventional protein phosphorylation sites are usually initially detected based on their acid sensitivity, since other methods of
detection are difficult using conventional proteomic methodologies. Finally, Phillips et al. (161) demonstrated protein phosphorylation by ␥-[32P]ATP in oxidative phosphorylation complexes resolved with blue native gel electrophoresis with no
detectable kinases associated within the protein complexes.
These later results are consistent with the autocatalytic processes in the bacterial phosphorylation pathways.
From a kinase sequence or motif searching approach, the two
protein kinases that are unequivocally shown to be in the matrix,
PDHK and BCKDH kinase (BCKDHK), have more sequence
similarity to bacterial histidine kinases motifs than to cytosolic
serine protein kinases (77). However, no evidence exists that these
kinases phosphorylate anything other than the E1␣ serines of their
target proteins. We and others have screened the mammalian
mitochondrial nuclear and mitochondrial genome for the limited
consensus eukaryotic phospho-histidine kinases (154) and other
nonconventional kinases motifs with little success. Specifically,
we have used the bacterial histidine kinase motifs (PF02895) from
PFAM (169) to BLAST against the mouse genome. A brief
overview of these searches with regard to mitochondrial proteins include 1) IPR003660 HAMP linker domain; no proteins;
2) IPR005467 signal transduction histidine kinase, core; no proteins; 3) IPR003594 ATPase-like, ATP-binding domain; the previously identified BCKDHK and PDH isoforms, as well as heat
shock protein 75 (TNF receptor-associated protein 1), and topoisomerase; 4) IPR010559 signal transduction histidine kinase,
internal region; no protein; and 5) IPR004358 signal transduction
histidine kinase-related protein, COOH-terminal; apparent isoforms of BCKDHK. This search confirms the sequence similarity
of PDHK and BCKDHK with bacterial kinases, whereas any
protein kinase activity associated with the mitochondrial heat
shock proteins and topoisomerase ATPase activity has to our
knowledge not been reported but might be an interesting area of
investigation. However, despite the considerable evidence for
labile phosphorylation events, the limited detection of bacterial
histidine kinase motifs, generated from bacteria or simple eukaryotic systems, suggests that either new motifs have evolved in
mammalia or completely different mechanisms for these labile
phosphorylation sites are in play. The evolution of new motifs is
not surprising given the different environmental pressures and
nuclear genomic recombination occurring in the mitochondria
versus bacterial systems.
Though definitive evidence is not available for the existence
of these nonconventional protein phosphorylations in the mitochondrial matrix, the circumstantial evidences discussed, as
well as the apparent origins of many mitochondrial matrix
reactions, suggest that a careful examination of this process
should be conducted. Recent developments in MS methods
AJP-Heart Circ Physiol • doi:10.1152/ajpheart.00077.2012 • www.ajpheart.org
Downloaded from http://ajpheart.physiology.org/ by 10.220.32.246 on May 5, 2017
phorylation of BCKDH (98, 126). Since the phosphorylation of
BCKDH was found to change opposite to that of PDH under a
number of conditions (24), the differences between these
process might be the regulation of PP2Cm. Outside of these
phosphatases, the only other confirmed phosphatase is PTMT1,
which was identified as a tyrosine phosphatase (152); however,
subsequent studies demonstrated it was a phosphatidylglycerophosphate phosphatase that plays a role in cardiolipin biosynthesis (234).
Other phosphatases have been localized to the outer membrane or intermembrane space, such as Aup1p (207), PPL2a
(178), Shp-2 (181), PHLPP-1 (138), and DSP-18 (171), consistent with the dominant localization of cytosolic protein
kinases in the same space. Thus, even though extensive phosphatase activity is present in the matrix based on 32P turnover
and quantitative MS studies (24), the phosphatases of the
matrix remain obscure. Added evidence that these phosphatases exist in the aqueous phase in the matrix was the demonstration of phosphatase activity on the inner membrane by a
matrix extract in potato mitochondria (204). Clearly, the protein phosphatase system within the mitochondria matrix is as ill
defined as are the matrix kinase components, and a clear
mechanism to explain the larger turnover of protein phosphorylation in the matrix is yet to be resolved.
H959
Review
H960
MITOCHONDRIA PROTEIN PHOSPHORYLATION
suitable for the identification of these sites should provide
useful tools in this effort (21, 117, 238).
signaling pathways exist, they have likely evolved new functional motifs.
Conclusions and Future Directions
DISCLOSURES
No conflicts of interest, financial or otherwise, are declared by the author (s).
AUTHOR CONTRIBUTIONS
R.C. and R.S.B. conception and design of research; R.C. and R.S.B.
analyzed data; R.C. and R.S.B. interpreted results of experiments; R.C. and
R.S.B. prepared figures; R.C. and R.S.B. drafted manuscript; R.C. and R.S.B.
edited and revised manuscript; R.C. and R.S.B. approved final version of
manuscript; R.S.B. performed experiments.
REFERENCES
1. Acin-Perez R, Gatti DL, Bai Y, Manfredi G. Protein phosphorylation
and prevention of cytochrome oxidase inhibition by ATP: coupled
mechanisms of energy metabolism regulation. Cell Metab 13: 712–719,
2011.
2. Acin-Perez R, Salazar E, Kamenetsky M, Buck J, Levin LR, Manfredi G. Cyclic AMP produced inside mitochondria regulates oxidative
phosphorylation. Cell Metab 9: 265–276, 2009.
3. Acin-Perez R, Hoyos B, Gong J, Vinogradov V, Fischman DA,
Leitges M, Borhan B, Starkov A, Manfredi G, Hammerling U.
Regulation of intermediary metabolism by the PKCdelta signalosome in
mitochondria. FASEB J 24: 5033–5042, 2010.
4. Alto NM, Soderling J, Scott JD. Rab32 is an A-kinase anchoring
protein and participates in mitochondrial dynamics. J Cell Biol 158:
659 –668, 2002.
5. Aponte AM, Phillips D, Harris RA, Blinova K, French S, Johnson
DT, Balaban RS. 32P labeling of protein phosphorylation and metabolite association in the mitochondria matrix. Methods Enzymol 457:
63–80, 2009.
6. Aponte AM, Phillips D, Hopper RK, Johnson DT, Harris RA,
Blinova K, Boja ES, French S, Balaban RS. Use of (32)P to study
dynamics of the mitochondrial phosphoproteome. J Proteome Res 8:
2679 –2695, 2009.
7. Aprille JR. Mechanism and regulation of the mitochondrial ATP-Mg/
P(i) carrier. J Bioenerg Biomembr 25: 473–481, 1993.
8. Arnold S, Goglia F, Kadenbach B. 3,5-Diiodothyronine binds to
subunit Va of cytochrome-c oxidase and abolishes the allosteric inhibition of respiration by ATP. Eur J Biochem 252: 325–330, 1998.
9. Arrell DK, Elliott ST, Kane LA, Guo Y, Ko YH, Pedersen PL,
Robinson J, Murata M, Murphy AM, Marbán E, Van Eyk JE.
Proteomic analysis of pharmacological preconditioning: novel protein
targets converge to mitochondrial metabolism pathways. Circ Res 99:
706 –714, 2006.
10. Balaban RS. Cardiac energy metabolism homeostasis: role of cytosolic
calcium. J Mol Cell Cardiol 34: 1259 –1271, 2002.
11. Balaban RS. Maintenance of the metabolic homeostasis of the heart:
developing a systems analysis approach. Ann NY Acad Sci 1080: 140 –
153, 2006.
12. Balaban RS. Domestication of the cardiac mitochondrion for energy
conversion. J Mol Cell Cardiol 46: 832–841, 2009.
13. Balaban RS. The role of Ca2⫹ signaling in the coordination of mitochondrial ATP production with cardiac work. Biochim Biophys Acta
1787: 1334 –1341, 2009.
14. Baranova EA, Holt PJ, Sazanov LA. Projection structure of the
membrane domain of Escherichia coli respiratory complex I at 8 A
resolution. J Mol Biol 366: 140 –154, 2007.
15. Bason JV, Runswick MJ, Fearnley IM, Walker JE. Binding of the
inhibitor protein IF(1) to bovine F(1)-ATPase. J Mol Biol 406: 443–453,
2011.
16. Belogrudov GI. Recent advances in structure-functional studies of
mitochondrial factor B. J Bioenerg Biomembr 41: 137–143, 2009.
17. Bender A, Hajieva P, Moosmann B. Adaptive antioxidant methionine
accumulation in respiratory chain complexes explains the use of a deviant
genetic code in mitochondria. Proc Natl Acad Sci USA 105: 16496 –
16501, 2008.
18. Bender E, Kadenbach B. The allosteric ATP-inhibition of cytochrome
c oxidase activity is reversibly switched on by cAMP-dependent phosphorylation. FEBS Lett 466: 130 –134, 2000.
AJP-Heart Circ Physiol • doi:10.1152/ajpheart.00077.2012 • www.ajpheart.org
Downloaded from http://ajpheart.physiology.org/ by 10.220.32.246 on May 5, 2017
It is clear that mitochondrial matrix protein phosphorylation
is extensive when detected at high sensitivity. However, the
functional impact of these phosphorylations and their kinase/
phosphatase system remain generally obscure for most of the
sites identified. We take the general position that most of the
sites detected with these high-sensitivity approaches are not
impacting enzyme function but are low level phosphorylation
“noise” from numerous protein kinase reactions. The issue in
the field is separating the noise from the regulatory sites. One
of the major limitations for the metabolic enzymes is the lack
of information on the fraction of the enzyme that is phosphorylated. The development of quantitative protein phosphorylation methods is underway around the world and, we predict,
will greatly reduce the number candidate sites for regulatory
activity in the energy metabolism enzyme network in two
ways: first, establishing the fraction of the enzymatic pool
phosphorylated and second, quantitatively relating the amount
of phosphorylation with the enzyme activity [for example, see
Boja et al. (24)]. It is important to note that cataloging the
phosphorylation sites is still very important for several reasons.
First, the detection of the phosphorylation event confirms that
the site can be modified by existing kinases. Second, the
limited functional analysis we perform and the conditions we
can create may miss the functional consequences of these
phosphorylations on processes such as assembly, transport, and
unknown functions that we cannot or know not to assay.
Many protein kinases and phosphatases are selectively targeted to the outer compartments of the mitochondria through
very sophisticated cell signaling processes; however, the functional significance of this localization is less defined. With
regard to the matrix, the currently established matrix kinase
and phosphatase system is inadequate to explain the extent of
protein phosphorylation detected. One possibility was that
most of the phosphorylation events occur in the cytosol or
kinase rich intermembrane space and outer mitochondrial
membrane before, or during, protein transport into the mitochondria. However, 32P studies on intact mitochondria clearly
establish extensive matrix protein kinase activity. In silico
surveys of the phosphorylation sites suggest no specific conventional protein kinases responsible for the phosphorylation
patterns observed, with some exceptions. Thus the mechanisms
for much of the matrix protein phosphorylation detected, especially with 32P labeling in intact mitochondria, remain unknown. The mitochondrion has an early bacterial origin, and
based on several lines of evidence including the pH and
temporal sensitivity of protein phosphorylation and the presence of autophosphorylation events within the complexes, it
can be suggested that bacterial nonconventional protein phosphorylation transfer events may be playing a role in protein
phosphorylation. The test of this hypothesis will require the
application of nonconventional methods to monitor these putative high-energy protein phosphorylation events. It is important to note that very little genetic evidence exists for conserved nonconventional kinase activity in the mitochondrial
proteome. These later results suggest that if these bacterial
Review
MITOCHONDRIA PROTEIN PHOSPHORYLATION
43. Couoh-Cardel SJ, Uribe-Carvajal S, Wilkens S, Garcia-Trejo JJ.
Structure of dimeric F1F0-ATP synthase. J Biol Chem 285: 36447–
36455, 2010.
44. Court N, Ingley E, Klinken SP, Bogoyevitch MA. Outer membrane
protein 25-a mitochondrial anchor and inhibitor of stress-activated protein kinase-3. Biochim Biophys Acta 1744: 68 –75, 2005.
45. Covian R, Trumpower BL. Regulatory interactions between ubiquinol
oxidation and ubiquinone reduction sites in the dimeric cytochrome bc1
complex. J Biol Chem 281: 30925–30932, 2006.
46. Crofts AR, Guergova-Kuras M, Huang L, Kuras R, Zhang Z, Berry
EA. Mechanism of ubiquinol oxidation by the bc(1) complex: role of the
iron sulfur protein and its mobility. Biochemistry 38: 15791–15806,
1999.
47. Cui Z, Hou J, Chen X, Li J, Xie Z, Xue P, Cai T, Wu P, Xu T, Yang
F. The profile of mitochondrial proteins and their phosphorylation
signaling network in INS-1 beta cells. J Proteome Res 9: 2898 –2908,
2010.
48. Dai J, Jin WH, Sheng QH, Shieh CH, Wu JR, Zeng R. Protein
phosphorylation and expression profiling by Yin-yang multidimensional
liquid chromatography (Yin-yang MDLC) mass spectrometry. J Proteome Res 6: 250 –262, 2007.
49. Damuni Z, Merryfield ML, Humphreys JS, Reed LJ. Purification and
properties of branched-chain alpha-keto acid dehydrogenase phosphatase
from bovine kidney. Proc Natl Acad Sci USA 81: 4335–4338, 1984.
50. Damuni Z, Reed LJ. Purification and properties of the catalytic subunit
of the branched-chain alpha-keto acid dehydrogenase phosphatase from
bovine kidney mitochondria. J Biol Chem 262: 5129 –5132, 1987.
51. Dasari VR, Anandatheerthavarada HK, Robin MA, Boopathi E,
Biswas G, Fang JK, Nebert DW, Avadhani NG. Role of protein kinase
C-mediated protein phosphorylation in mitochondrial translocation of
mouse CYP1A1, which contains a non-canonical targeting signal. J Biol
Chem 281: 30834 –30847, 2006.
52. Dave KR, Defazio RA, Raval AP, Torraco A, Saul I, Barrientos A,
Perez-Pinzon MA. Ischemic preconditioning targets the respiration of
synaptic mitochondria via protein kinase C epsilon. J Neurosci 28:
4172–4182, 2008.
53. Davie JR, Wynn RM, Meng M, Huang YS, Aalund G, Chuang DT,
Lau KS. Expression and characterization of branched-chain alphaketoacid dehydrogenase kinase from the rat. Is it a histidine-protein
kinase? J Biol Chem 270: 19861–19867, 1995.
54. De Rasmo D, Panelli D, Sardanelli AM, Papa S. cAMP-dependent
protein kinase regulates the mitochondrial import of the nuclear encoded
NDUFS4 subunit of complex I. Cell Signal 20: 989 –997, 2008.
55. De Rasmo D, Palmisano G, Scacco S, Technikova-Dobrova Z, Panelli
D, Cocco T, Sardanelli AM, Gnoni A, Micelli L, Trani A, Di Luccia
A, Papa S. Phosphorylation pattern of the NDUFS4 subunit of complex
I of the mammalian respiratory chain. Mitochondrion 10: 464 –471, 2010.
57. Deng K, Shenoy SK, Tso SC, Yu L, Yu CA. Reconstitution of
mitochondrial processing peptidase from the core proteins (subunits I and
II) of bovine heart mitochondrial cytochrome bc(1) complex. J Biol
Chem 276: 6499 –6505, 2001.
58. Deng N, Zhang J, Zong C, Wang Y, Lu H, Yang P, Wang W, Young
GW, Wang Y, Korge P, Lotz C, Doran P, Liem DA, Apweiler R,
Weiss JN, Duan H, Ping P. Phosphoproteome analysis reveals regulatory sites in major pathways of cardiac mitochondria. Mol Cell Proteomics. 2011 Feb; 10(2):M110.000117. Epub 2010 May 22.
59. Denton RM. Regulation of mitochondrial dehydrogenases by calcium
ions. Biochim Biophys Acta 1787: 1309 –1316, 2009.
60. Denton RM, Randle P, Martin BR. Stimulation by calciums ions of
pyruvate dehydrogenase phosphate phosphatase. Biochem J 128: 161–
163, 1972.
61. Dephoure N, Zhou C, Villen J, Beausoleil SA, Bakalarski CE,
Elledge SJ, Gygi SP. A quantitative atlas of mitotic phosphorylation.
Proc Natl Acad Sci USA 105: 10762–10767, 2008.
62. Depontieu FR, Qian J, Zarling AL, McMiller TL, Salay TM, Norris
A, English AM, Shabanowitz J, Engelhard VH, Hunt DF, Topalian
SL. Identification of tumor-associated, MHC class II-restricted phosphopeptides as targets for immunotherapy. Proc Natl Acad Sci USA 106:
12073–12078, 2009.
63. Di PF, Bisetto E, Alverdi V, Mavelli I, Esposito G, Lippe G. Differential steady-state tyrosine phosphorylation of two oligomeric forms of
mitochondrial F0F1ATPsynthase: a structural proteomic analysis. Proteomics 6: 921–926, 2006.
AJP-Heart Circ Physiol • doi:10.1152/ajpheart.00077.2012 • www.ajpheart.org
Downloaded from http://ajpheart.physiology.org/ by 10.220.32.246 on May 5, 2017
19. Bera AK, Ghosh S. Dual mode of gating of voltage-dependent anion
channel as revealed by phosphorylation. J Struct Biol 135: 67–72, 2001.
20. Berrisford JM, Sazanov LA. Structural basis for the mechanism of
respiratory complex I. J Biol Chem 284: 29773–29783, 2009.
21. Besant PG, Attwood PV. Detection and analysis of protein histidine
phosphorylation. Mol Cell Biochem 329: 93–106, 2009.
22. Blom N, Sicheritz-Ponten T, Gupta R, Gammeltoft S, Brunak S.
Prediction of post-translational glycosylation and phosphorylation of
proteins from the amino acid sequence. Proteomics 4: 1633–1649, 2004.
23. Boerner JL, Demory ML, Silva C, Parsons SJ. Phosphorylation of
Y845 on the epidermal growth factor receptor mediates binding to the
mitochondrial protein cytochrome c oxidase subunit II. Mol Cell Biol 24:
7059 –7071, 2004.
24. Boja ES, Phillips D, French SA, Harris RA, Balaban RS. Quantitative
mitochondrial phosphoproteomics using iTRAQ on an LTQ-Orbitrap
with high energy collision dissociation. J Proteome Res 8: 4665–4675,
2009.
25. Bowler MW, Montgomery MG, Leslie AG, Walker JE. Ground state
structure of F1-ATPase from bovine heart mitochondria at 1.9 A resolution. J Biol Chem 282: 14238 –14242, 2007.
26. Brandt U. Energy converting NADH: quinone oxidoreductase (complex
I). Annu Rev Biochem 75: 69 –92, 2006.
27. Brasseur G, Saribas AS, Daldal F. A compilation of mutations located
in the cytochrome b subunit of the bacterial and mitochondrial bc1
complex. Biochim Biophys Acta 1275: 61–69, 1996.
28. Buxton DB, Olson MS. Regulation of the branched chain alpha-keto
acid and pyruvate dehydrogenases in the perfused rat heart. J Biol Chem
257: 15026 –15029, 1982.
29. Bykova NV, Egsgaard H, Moller IM. Identification of 14 new phosphoproteins involved in important plant mitochondrial processes. FEBS
Lett 540: 141–146, 2003.
30. Cabezon E, Arechaga I, Jonathan P, Butler G, Walker JE. Dimerization of bovine F1-ATPase by binding the inhibitor protein, IF1. J Biol
Chem 275: 28353–28355, 2000.
31. Cabezon E, Montgomery MG, Leslie AG, Walker JE. The structure of
bovine F1-ATPase in complex with its regulatory protein IF1. Nat Struct
Biol 10: 744 –750, 2003.
32. Cabezon E, Runswick MJ, Leslie AG, Walker JE. The structure of
bovine IF(1), the regulatory subunit of mitochondrial F-ATPase. EMBO
J 20: 6990 –6996, 2001.
33. Cantin GT, Yi W, Lu B, Park SK, Xu T, Lee JD, Yates JR 3rd.
Combining protein-based IMAC, peptide-based IMAC, and MudPIT for
efficient phosphoproteomic analysis. J Proteome Res 7: 1346 –1351,
2008.
34. Carbajo RJ, Kellas FA, Yang JC, Runswick MJ, Montgomery MG,
Walker JE, Neuhaus D. How the N-terminal domain of the OSCP
subunit of bovine F1Fo-ATP synthase interacts with the N-terminal
region of an alpha subunit. J Mol Biol 368: 310 –318, 2007.
35. Castro L, Demicheli V, Tortora V, Radi R. Mitochondrial protein
tyrosine nitration. Free Radic Res 45: 37–52, 2011.
36. Chen R, Fearnley IM, Peak-Chew SY, Walker JE. The phosphorylation of subunits of complex I from bovine heart mitochondria. J Biol
Chem 279: 26036 –26045, 2004.
37. Chen Y, Lu B, Yang Q, Fearns C, Yates JR, III, Lee JD. Combined
integrin phosphoproteomic analyses and small interfering RNA-based
functional screening identify key regulators for cancer cell adhesion and
migration. Cancer Res 69: 3713–3720, 2009.
38. Chen Y, Gaczynska M, Osmulski P, Polci R, Riley DJ. Phosphorylation by Nek1 regulates opening and closing of voltage dependent anion
channel 1. Biochem Biophys Res Commun 394: 798 –803, 2010.
39. Choudhary C, Olsen JV, Brandts C, Cox J, Reddy PN, Bohmer FD,
Gerke V, Schmidt-Arras DE, Berdel WE, Muller-Tidow C, Mann M,
Serve H. Mislocalized activation of oncogenic RTKs switches downstream signaling outcomes. Mol Cell 36: 326 –339, 2009.
40. Christensen GL, Kelstrup CD, Lyngso C, Sarwar U, Bogebo R,
Sheikh SP, Gammeltoft S, Olsen JV, Hansen JL. Quantitative phosphoproteomics dissection of seven-transmembrane receptor signaling
using full and biased agonists. Mol Cell Proteomics 9: 1540 –1553, 2010.
41. Chu CT. Tickled PINK1: mitochondrial homeostasis and autophagy in
recessive Parkinsonism. Biochim Biophys Acta 1802: 20 –28, 2010.
42. Colombini M. VDAC: the channel at the interface between mitochondria
and the cytosol. Mol Cell Biochem 256 –257: 107–115, 2004.
H961
Review
H962
MITOCHONDRIA PROTEIN PHOSPHORYLATION
The mTOR-regulated phosphoproteome reveals a mechanism of
mTORC1-mediated inhibition of growth factor signaling. Science 332:
1317–1322, 2011.
85. Huang B, Gudi R, Wu P, Harris RA, Hamilton J, Popov KM.
Isoenzymes of pyruvate dehydrogenase phosphatase. J Biol Chem 273:
17680 –17688, 1998.
86. Huang LJ, Wang L, Ma Y, Durick K, Perkins G, Deerinck TJ,
Ellisman MH, Taylor SS. NH2-terminal targeting motifs direct dual
specificity A-kinase-anchoring protein 1 (D-AKAP1) to either mitochondria or endoplasmic reticulum. J Cell Biol 145: 951–959, 1999.
87. Huang LS, Cobessi D, Tung EY, Berry EA. Binding of the respiratory
chain inhibitor antimycin to the mitochondrial bc1 complex: a new
crystal structure reveals an altered intramolecular hydrogen-bonding
pattern. J Mol Biol 351: 573–597, 2005.
88. Huang SY, Tsai ML, Chen GY, Wu CJ, Chen SH. A systematic
MS-based approach for identifying in vitro substrates of PKA and PKG
in rat uteri. J Proteome Res 6: 2674 –2684, 2007.
89. Hughes WA, Halestrap AP. The regulation of branched-chain 2-oxo
acid dehydrogenase of liver, kidney and heart by phosphorylation.
Biochem J 196: 459 –469, 1981.
90. Hunte C, Palsdottir H, Trumpower BL. Protonmotive pathways and
mechanisms in the cytochrome bc1 complex. FEBS Lett 545: 39 –46,
2003.
91. Hurd TR, Costa NJ, Dahm CC, Beer SM, Brown SE, Filipovska A,
Murphy MP. Glutathionylation of mitochondrial proteins. Antioxid
Redox Signal 7: 999 –1010, 2005.
92. Huttemann M, Helling S, Sanderson TH, Sinkler C, Samavati L,
Mahapatra G, Varughese A, Lu G, Liu J, Ramzan R, Vogt S,
Grossman LI, Doan JW, Marcus K, Lee I. Regulation of mitochondrial
respiration and apoptosis through cell signaling: cytochrome c oxidase
and cytochrome c in ischemia/reperfusion injury and inflammation.
Biochim Biophys Acta 1817: 598 –609, 2012.
93. Huttemann M, Kadenbach B, Grossman LI. Mammalian subunit IV
isoforms of cytochrome c oxidase. Gene 267: 111–123, 2001.
94. Huttlin EL, Jedrychowski MP, Elias JE, Goswami T, Rad R, Beausoleil SA, Villen J, Haas W, Sowa ME, Gygi SP. A tissue-specific atlas
of mouse protein phosphorylation and expression. Cell 143: 1174 –1189,
2010.
95. Imami K, Sugiyama N, Kyono Y, Tomita M, Ishihama Y. Automated
phosphoproteome analysis for cultured cancer cells by two-dimensional
nanoLC-MS using a calcined titania/C18 biphasic column. Anal Sci 24:
161–166, 2008.
96. Iwata S, Lee JW, Okada K, Lee JK, Iwata M, Rasmussen B, Link
TA, Ramaswamy S, Jap BK. Complete structure of the 11-subunit
bovine mitochondrial cytochrome bc1 complex. Science 281: 64 –71,
1998.
97. Johnson ET, Parson WW. Electrostatic interactions in an integral
membrane protein. Biochemistry 41: 6483–6494, 2002.
98. Joshi M, Jeoung NH, Popov KM, Harris RA. Identification of a novel
PP2C-type mitochondrial phosphatase. Biochem Biophys Res Commun
356: 38 –44, 2007.
99. Jung K, Jung H. A new mechanism of phosphoregulation in signal
transduction pathways. Sci Signal 2: e71, 2009.
100. Kadenbach B, Napiwotzki J, Frank V, Arnold S, Exner S, Huttemann M. Regulation of energy transduction and electron transfer in
cytochrome c oxidase by adenine nucleotides. J Bioenerg Biomembr 30:
25–33, 1998.
101. Kane LA, Youngman MJ, Jensen RE, Van Eyk JE. Phosphorylation
of the F(1)F(o) ATP synthase beta subunit: functional and structural
consequences assessed in a model system. Circ Res 106: 504 –513, 2010.
102. Kerbey AL, Randle PJ, Cooper RH, Whitehouse S, Pask HT, Denton
RM. Regulation of pyruvate dehydrogenase in rat heart. Mechanism of
regulation of proportions of dephosphorylated and phosphorylated enzyme by oxidation of fatty acids and ketone bodies and of effects of
diabetes: role of coenzyme A, acetyl-coenzyme A and reduced and
oxidized nicotinamide-adenine dinucleotide. Biochem J 154: 327–348,
1976.
103. Kerner J, Distler AM, Minkler P, Parland W, Peterman SM, Hoppel
CL. Phosphorylation of rat liver mitochondrial carnitine palmitoyltransferase-I: effect on the kinetic properties of the enzyme. J Biol Chem 279:
41104 –41113, 2004.
104. Kharbanda S, Saxena S, Yoshida K, Pandey P, Kaneki M, Wang Q,
Cheng K, Chen YN, Campbell A, Sudha T, Yuan ZM, Narula J,
Weichselbaum R, Nalin C, Kufe D. Translocation of SAPK/JNK to
AJP-Heart Circ Physiol • doi:10.1152/ajpheart.00077.2012 • www.ajpheart.org
Downloaded from http://ajpheart.physiology.org/ by 10.220.32.246 on May 5, 2017
64. Efremov RG, Sazanov LA. Structure of the membrane domain of
respiratory complex I. Nature 476: 414 –420, 2011.
65. Fang JK, Prabu SK, Sepuri NB, Raza H, Anandatheerthavarada
HK, Galati D, Spear J, Avadhani NG. Site specific phosphorylation of
cytochrome c oxidase subunits I, IVi1 and Vb in rabbit hearts subjected
to ischemia/reperfusion. FEBS Lett 581: 1302–1310, 2007.
66. Feliciello A, Gottesman ME, Avvedimento EV. cAMP-PKA signaling
to the mitochondria: protein scaffolds, mRNA and phosphatases. Cell
Signal 17: 279 –287, 2005.
67. Feng J, Lucchinetti E, Enkavi G, Wang Y, Gehrig P, Roschitzki B,
Schaub MC, Tajkhorshid E, Zaugg K, Zaugg M. Tyrosine phosphorylation by Src within the cavity of the adenine nucleotide translocase 1
regulates ADP/ATP exchange in mitochondria. Am J Physiol Cell
Physiol 298: C740 –C748, 2010.
68. Feng J, Zhu M, Schaub MC, Gehrig P, Roschitzki B, Lucchinetti E,
Zaugg M. Phosphoproteome analysis of isoflurane-protected heart mitochondria: phosphorylation of adenine nucleotide translocator-1 on
Tyr194 regulates mitochondrial function. Cardiovasc Res 80: 20 –29,
2008.
69. Galante YM, Hatefi Y. Resolution of complex I and isolation of NADH
dehydrogenase and an iron-sulfur protein. Methods Enzymol 53: 15–21,
1978.
70. Garcia JJ, Morales-Rios E, Cortes-Hernandez P, Rodriguez-Zavala
JS. The inhibitor protein (IF1) promotes dimerization of the mitochondrial F1F0-ATP synthase. Biochemistry 45: 12695–12703, 2006.
71. Gledhill JR, Montgomery MG, Leslie AG, Walker JE. How the
regulatory protein, IF(1), inhibits F(1)-ATPase from bovine mitochondria. Proc Natl Acad Sci USA 104: 15671–15676, 2007.
72. Goodwin GW, Taylor CS, Taegtmeyer H. Regulation of energy metabolism of the heart during acute increase in heart work. J Biol Chem
273: 29530 –29539, 1998.
73. Gu TL, Deng X, Huang F, Tucker M, Crosby K, Rimkunas V, Wang
Y, Deng G, Zhu L, Tan Z, Hu Y, Wu C, Nardone J, Macneill J, Ren
J, Reeves C, Innocenti G, Norris B, Yuan J, Yu J, Haack H, Shen B,
Peng C, Li H, Zhou X, Liu X, Rush J, Comb MJ. Survey of tyrosine
kinase signaling reveals ROS kinase fusions in human cholangiocarcinoma. PLoS One 6: e15640, 2011.
74. Han G, Ye M, Liu H, Song C, Sun D, Wu Y, Jiang X, Chen R, Wang
C, Wang L, Zou H. Phosphoproteome analysis of human liver tissue by
long-gradient nanoflow LC coupled with multiple stage MS analysis.
Electrophoresis 31: 1080 –1089, 2010.
75. Harootunian AT, Adams SR, Wen W, Meinkoth JL, Taylor SS,
Tsien RY. Movement of the free catalytic subunit of cAMP-dependent
protein kinase into and out of the nucleus can be explained by diffusion.
Mol Biol Cell 4: 993–1002, 1993.
76. Harper AE, Miller RH, Block KP. Branched-chain amino acid metabolism. Annu Rev Nutr 4: 409 –454, 1984.
77. Harris RA, Popov KM, Zhao Y, Kedishvili NY, Shimomura Y,
Crabb DW. A new family of protein kinases—the mitochondrial protein
kinases. Adv Enzyme Regul 35: 147–162, 1995.
78. He L, Lemasters JJ. Dephosphorylation of the Rieske iron-sulfur
protein after induction of the mitochondrial permeability transition.
Biochem Biophys Res Commun 334: 829 –837, 2005.
79. Helling S, Vogt S, Rhiel A, Ramzan R, Wen L, Marcus K, Kadenbach B. Phosphorylation and kinetics of mammalian cytochrome c
oxidase. Mol Cell Proteomics 7: 1714 –1724, 2008.
80. Hoffert JD, Nielsen J, Yu MJ, Pisitkun T, Schleicher SM, Nielsen S,
Knepper MA. Dynamics of aquaporin-2 serine-261 phosphorylation in
response to short-term vasopressin treatment in collecting duct. Am J
Physiol Renal Physiol 292: F691–F700, 2007.
81. Hoffert JD, Pisitkun T, Wang G, Shen RF, Knepper MA. Quantitative
phosphoproteomics of vasopressin-sensitive renal cells: regulation of
aquaporin-2 phosphorylation at two sites. Proc Natl Acad Sci USA 103:
7159 –7164, 2006.
82. Hojlund K, Yi Z, Lefort N, Langlais P, Bowen B, Levin K, BeckNielsen H, Mandarino LJ. Human ATP synthase beta is phosphorylated
at multiple sites and shows abnormal phosphorylation at specific sites in
insulin-resistant muscle. Diabetologia 53: 541–551, 2010.
83. Hopper RK, Carroll S, Aponte AM, Johnson DT, French S, Shen RF,
Witzmann FA, Harris RA, Balaban RS. Mitochondrial matrix phosphoproteome: effect of extra mitochondrial calcium. Biochemistry 45:
2524 –2536, 2006.
84. Hsu PP, Kang SA, Rameseder J, Zhang Y, Ottina KA, Lim D,
Peterson TR, Choi Y, Gray NS, Yaffe MB, Marto JA, Sabatini DM.
Review
MITOCHONDRIA PROTEIN PHOSPHORYLATION
105.
106.
107.
108.
109.
111.
112.
113.
114.
115.
116.
117.
118.
119.
120.
121.
122.
123.
124.
125.
126.
127.
128. Lund AA, Rhoads DM, Lund AL, Cerny RL, Elthon TE. In vivo
modifications of the maize mitochondrial small heat stress protein,
HSP22. J Biol Chem 276: 29924 –29929, 2001.
129. Majumder PK, Mishra NC, Sun X, Bharti A, Kharbanda S, Saxena
S, Kufe D. Targeting of protein kinase C delta to mitochondria in the
oxidative stress response. Cell Growth Differ 12: 465–470, 2001.
130. Margulis L, Sagan D. Acquiring Genomes: A Yheory of the Origins of
Species. New York: Basic Books, 2002.
131. Mayya V, Lundgren DH, Hwang SI, Rezaul K, Wu L, Eng JK,
Rodionov V, Han DK. Quantitative phosphoproteomic analysis of T cell
receptor signaling reveals system-wide modulation of protein-protein
interactions. Sci Signal 2: ra46, 2009.
132. McCormack JG, Denton RM. The activation of pyruvate dehydrogenase in the perfused rat heart by adrenaline and other inotropic agents.
Biochem J 194: 639 –643, 1981.
133. McCormack JG, Halestrap AP, Denton RM. Role of calcium ions in
regulation of mammalian intramitochondrial metabolism. Physiol Rev
70: 391–425, 1990.
134. Means CK, Lygren B, Langeberg LK, Jain A, Dixon RE, Vega AL,
Gold MG, Petrosyan S, Taylor SS, Murphy AN, Ha T, Santana LF,
Tasken K, Scott JD. An entirely specific type I A-kinase anchoring
protein that can sequester two molecules of protein kinase A at mitochondria. Proc Natl Acad Sci USA 108: E1227–E1235, 2011.
135. Mehler EL, Guarnieri F. A self-consistent, microenvironment modulated screened coulomb potential approximation to calculate pH-dependent electrostatic effects in proteins. Biophys J 77: 3–22, 1999.
136. Miller JL, Cimen H, Koc H, Koc EC. Phosphorylated proteins of the
mammalian mitochondrial ribosome: implications in protein synthesis. J
Proteome Res 8: 4789 –4798, 2009.
137. Miura T, Tanno M, Sato T. Mitochondrial kinase signalling pathways
in myocardial protection from ischaemia/reperfusion-induced necrosis.
Cardiovasc Res 88: 7–15, 2010.
138. Miyamoto S, Purcell NH, Smith JM, Gao T, Whittaker R, Huang K,
Castillo R, Glembotski CC, Sussman MA, Newton AC, Brown JH.
PHLPP-1 negatively regulates akt activity and survival in the heart/
novelty and significance. Circ Res 107: 476 –484, 2010.
139. Miyazaki T, Neff L, Tanaka S, Horne WC, Baron R. Regulation of
cytochrome c oxidase activity by c-Src in osteoclasts. J Cell Biol 160:
709 –718, 2003.
140. Mootha VK, Arai AE, Balaban RS. Maximum oxidative phosphorylation capacity of the mammalian heart. Am J Physiol Heart Circ Physiol
272: H769 –H775, 1997.
141. Moritz A, Li Y, Guo A, Villen J, Wang Y, Macneill J, Kornhauser J,
Sprott K, Zhou J, Possemato A, Ren JM, Hornbeck P, Cantley LC,
Gygi SP, Rush J, Comb MJ. Akt-RSK-S6 kinase signaling networks
activated by oncogenic receptor tyrosine kinases. Sci Signal 3: ra64,
2010.
142. Murphy E, Kohr M, Sun J, Nguyen T, Steenbergen C. S-nitrosylation:
a radical way to protect the heart. J Mol Cell Cardiol 52: 568 –577, 2012.
143. Nantel A, Huber M, Thomas DY. Localization of endogenous Grb10 to
the mitochondria and its interaction with the mitochondrial-associated
Raf-1 pool. J Biol Chem 274: 35719 –35724, 1999.
144. Neubauer S. The failing heart—an engine out of fuel. N Engl J Med 356:
1140 –1151, 2007.
145. Neupert W, Brunner M. The protein import motor of mitochondria. Nat
Rev Mol Cell Biol 3: 555–565, 2002.
146. Norman AW, Bieber LL, Lindber GO, Boyer PD. Relationships of
Ca⫹⫹ to “protein-bound” phosphate fractions of mitochondria. J Biol
Chem 240: 2855–2862, 1965.
147. Obenauer JC, Cantley LC, Yaffe MB. Scansite 2.0: Proteome-wide
prediction of cell signaling interactions using short sequence motifs.
Nucleic Acids Res 31: 3635–3641, 2003.
148. Ogbi M, Chew CS, Pohl J, Stuchlik O, Ogbi S, Johnson JA. Cytochrome c oxidase subunit IV as a marker of protein kinase Cepsilon
function in neonatal cardiac myocytes: implications for cytochrome c
oxidase activity. Biochem J 382: 923–932, 2004.
149. Ogbi M, Johnson JA. Protein kinase Cepsilon interacts with cytochrome
c oxidase subunit IV and enhances cytochrome c oxidase activity in
neonatal cardiac myocyte preconditioning. Biochem J 393: 191–199,
2006.
150. Old WM, Shabb JB, Houel S, Wang H, Couts KL, Yen CY, Litman
ES, Croy CH, Meyer-Arendt K, Miranda JG, Brown RA, Witze ES,
Schweppe RE, Resing KA, Ahn NG. Functional proteomics identifies
AJP-Heart Circ Physiol • doi:10.1152/ajpheart.00077.2012 • www.ajpheart.org
Downloaded from http://ajpheart.physiology.org/ by 10.220.32.246 on May 5, 2017
110.
mitochondria and interaction with Bcl-xL in response to DNA damage. J
Biol Chem 275: 322–327, 2000.
Kim BG, Lee JH, Ahn JM, Park SK, Cho JH, Hwang D, Yoo JS,
Yates JR 3rd, Ryoo HM, Cho JY. ‘Two-stage double-technique hybrid
(TSDTH)’ identification strategy for the analysis of BMP2-induced
transdifferentiation of premyoblast C2C12 cells to osteoblast. J Proteome
Res 8: 4441–4454, 2009.
Kim CH, Yencha AJ, Bunker G, Zhang G, Chance B, King TE.
Effect of the hinge protein on the heme iron site of cytochrome c1.
Biochemistry 28: 1439 –1441, 1989.
Kim SC, Chen Y, Mirza S, Xu Y, Lee J, Liu P, Zhao Y. A clean, more
efficient method for in-solution digestion of protein mixtures without
detergent or urea. J Proteome Res 5: 3446 –3452, 2006.
King TE, Kim CH. Preparation of hinge protein and its requirement for
interaction of cytochrome c with cytochrome c1. Methods Enzymol 126:
238 –253, 1986.
Klingenberg M. The ADP and ATP transport in mitochondria and its
carrier. Biochim Biophys Acta 1778: 1978 –2021, 2008.
Klumpp S, Krieglstein J. Reversible phosphorylation of histidine residues in proteins from vertebrates. Sci Signal 2: e13, 2009.
Knebel A, Morrice N, Cohen P. A novel method to identify protein
kinase substrates: eEF2 kinase is phosphorylated and inhibited by
SAPK4/p38delta. EMBO J 20: 4360 –4369, 2001.
Ko YH, Pan W, Inoue C, Pedersen PL. Signal transduction to mitochondrial ATP synthase: evidence that PDGF-dependent phosphorylation of the delta-subunit occurs in several cell lines, involves tyrosine,
and is modulated by lysophosphatidic acid. Mitochondrion 1: 339 –348,
2002.
Kobayashi K, Neely JR. Mechanism of pyruvate dehydrogenase activation by increased cardiac work. J Mol Cell Cardiol 15: 369 –382, 1983.
Krebs EG, Fischer EH. The phosphorylase b to a converting enzyme of
rabbit skeletal muscle. Biochim Biophys Acta 20: 150 –157, 1956.
Kumar S, Bharti A, Mishra NC, Raina D, Kharbanda S, Saxena S,
Kufe D. Targeting of the c-Abl tyrosine kinase to mitochondria in the
necrotic cell death response to oxidative stress. J Biol Chem 276:
17281–17285, 2001.
Lange C, Hunte C. Crystal structure of the yeast cytochrome bc1
complex with its bound substrate cytochrome c. Proc Natl Acad Sci USA
99: 2800 –2805, 2002.
Lapek JD Jr, Tombline G, Friedman AE. Mass spectrometry detection
of histidine phosphorylation on NM23-H1. J Proteome Res 10: 751–755,
2011.
Lee I, Salomon AR, Ficarro S, Mathes I, Lottspeich F, Grossman LI,
Huttemann M. cAMP-dependent tyrosine phosphorylation of subunit I
inhibits cytochrome c oxidase activity. J Biol Chem 280: 6094 –6100,
2005.
Lee I, Salomon AR, Yu K, Doan JW, Grossman LI, Huttemann M.
New prospects for an old enzyme: mammalian cytochrome c is tyrosinephosphorylated in vivo. Biochemistry 45: 9121–9128, 2006.
Lee I, Salomon AR, Yu K, Samavati L, Pecina P, Pecinova A,
Huttemann M. Isolation of regulatory-competent, phosphorylated cytochrome C oxidase. Methods Enzymol 457: 193–210, 2009.
Lee J, Xu Y, Chen Y, Sprung R, Kim SC, Xie S, Zhao Y. Mitochondrial phosphoproteome revealed by an improved IMAC method and
MS/MS/MS. Mol Cell Proteomics 6: 669 –676, 2007.
Linberg O, Duffy JJ, Norman AW, Boyer PD. Characteristics of
bound phosphohistidine labeling in mitochondria. J Biol Chem 240:
2850 –2854, 1965.
Linn TC, Pettit FH, Reed LJ. Alpha-keto acid dehydrogenase complexes. X. Regulation of the activity of the pyruvate dehydrogenase
complex from beef kidney mitochondria by phosphorylation and dephosphorylation. Proc Natl Acad Sci USA 62: 234 –241, 1969.
Louro RO, Catarino T, Paquete CM, Turner DL. Distance dependence of interactions between charged centres in proteins with common
structural features. FEBS Lett 576: 77–80, 2004.
Lu G, Sun H, Korge P, Koehler CM, Weiss JN, Wang Y. Functional
characterization of a mitochondrial Ser/Thr protein phosphatase in cell
death regulation. Methods Enzymol 457: 255–273, 2009.
Lu G, Sun H, She P, Youn JY, Warburton S, Ping P, Vondriska TM,
Cai H, Lynch CJ, Wang Y. Protein phosphatase 2Cm is a critical
regulator of branched-chain amino acid catabolism in mice and cultured
cells. J Clin Invest 119: 1678 –1687, 2009.
Lubomir T. Mitochondrial protein phosphorylation: lessons from yeasts.
Gene 255: 59 –64, 2000.
H963
Review
H964
151.
152.
153.
154.
155.
157.
158.
159.
160.
161.
162.
163.
164.
165.
166.
167.
168.
169.
170.
targets of phosphorylation by B-Raf signaling in melanoma. Mol Cell 34:
115–131, 2009.
Olsen JV, Vermeulen M, Santamaria A, Kumar C, Miller ML,
Jensen LJ, Gnad F, Cox J, Jensen TS, Nigg EA, Brunak S, Mann M.
Quantitative phosphoproteomics reveals widespread full phosphorylation
site occupancy during mitosis. Sci Signal 3: ra3, 2010.
Pagliarini DJ, Wiley SE, Kimple ME, Dixon JR, Kelly P, Worby CA,
Casey PJ, Dixon JE. Involvement of a mitochondrial phosphatase in the
regulation of ATP production and insulin secretion in pancreatic beta
cells. Mol Cell 19: 197–207, 2005.
Pallen MJ. Time to recognise that mitochondria are bacteria? Trends
Microbiol 19: 58 –64, 2011.
Pao GM, Saier MH Jr. Nonplastid eukaryotic response regulators have
a monophyletic origin and evolved from their bacterial precursors in
parallel with their cognate sensor kinases. J Mol Evol 44: 605–613, 1997.
Papa S, Rasmo DD, Technikova-Dobrova Z, Panelli D, Signorile A,
Scacco S, Petruzzella V, Papa F, Palmisano G, Gnoni A, Micelli L,
Sardanelli AM. Respiratory chain complex I, a main regulatory target of
the cAMP/PKA pathway is defective in different human diseases. FEBS
Lett 586: 568 –577, 2012.
Paradela A, Albar JP. Advances in the analysis of protein phosphorylation. J Proteome Res 7: 1809 –1818, 2008.
Pastorino JG, Hoek JB, Shulga N. Activation of glycogen synthase
kinase 3beta disrupts the binding of hexokinase II to mitochondria by
phosphorylating voltage-dependent anion channel and potentiates chemotherapy-induced cytotoxicity. Cancer Res 65: 10545–10554, 2005.
Pebay-Peyroula E, Dahout-Gonzalez C, Kahn R, Trezeguet V, Lauquin GJM, Brandolin G. Structure of mitochondrial ADP/ATP carrier
in complex with carboxyatractyloside. Nature 426: 39 –44, 2003.
Pecina P, Borisenko GG, Belikova NA, Tyurina YY, Pecinova A, Lee
I, Samhan-Arias AK, Przyklenk K, Kagan VE, Huttemann M.
Phosphomimetic substitution of cytochrome C tyrosine 48 decreases
respiration and binding to cardiolipin and abolishes ability to trigger
downstream caspase activation. Biochemistry 49: 6705–6714, 2010.
Peter JB, Boyer PD. The formation of bound phosphohistidine from
adenosine triphosphate-P32 in mitochondria. J Biol Chem 238: 1180 –
1182, 1963.
Phillips D, Aponte AM, Covian RG, Balaban RS. Intrinsic protein
kinase activity of mitochondrial oxidative phosphorylation complexes.
Biochemistry 50: 2515–2529, 2011.
Phillips D, Aponte AM, French SA, Chess DJ, Balaban RS. SuccinylCoA synthetase is a phosphate target for the activation of mitochondrial
metabolism. Biochemistry 48: 7140 –7149, 2009.
Phillips D, Reilley MJ, Aponte AM, Wang G, Boja E, Gucek M,
Balaban RS. Stoichiometry of STAT3 and mitochondrial proteins:
implications for the regulation of oxidative phosphorylation by proteinprotein interactions. J Biol Chem 285: 23532–23536, 2010.
Pical C, Fredlund KM, Petit PX, Sommarin M, Moller IM. The outer
membrane of plant mitochondria contains a calcium-dependent protein
kinase and multiple phosphoproteins. FEBS Lett 336: 347–351, 1993.
Pocsfalvi G, Cuccurullo M, Schlosser G, Scacco S, Papa S, Malorni
A. Phosphorylation of B14.5a subunit from bovine heart complex I
identified by titanium dioxide selective enrichment and shotgun proteomics. Mol Cell Proteomics 6: 231–237, 2007.
Popov KM, Zhao Y, Shimomura Y, Kuntz MJ, Harris RA. Branchedchain alpha-ketoacid dehydrogenase kinase. Molecular cloning, expression, and sequence similarity with histidine protein kinases. J Biol Chem
267: 13127–13130, 1992.
Prabu SK, Anandatheerthavarada HK, Raza H, Srinivasan S, Spear
JF, Avadhani NG. Protein kinase A-mediated phosphorylation modulates cytochrome c oxidase function and augments hypoxia and myocardial ischemia-related injury. J Biol Chem 281: 2061–2070, 2006.
Pressman BC. Metabolic function of phosphohistidine. Biochem Biophys Res Commun 15: 556 –561, 1964.
Punta M, Coggill PC, Eberhardt RY, Mistry J, Tate J, Boursnell C,
Pang N, Forslund K, Ceric G, Clements J, Heger A, Holm L,
Sonnhammer EL, Eddy SR, Bateman A, Finn RD. The Pfam protein
families database. Nucleic Acids Res 40: D290 –D301, 2012.
Rao S, Schmidt O, Harbauer AB, Schonfisch B, Guiard B, Pfanner
N, Meisinger C. Biogenesis of the preprotein translocase of the outer
mitochondrial membrane: protein kinase A phosphorylates the precursor
of Tom40 and impairs its import. Mol Biol Cell 23: 1618 –1627, 2012.
171. Rardin MJ, Wiley SE, Murphy AN, Pagliarini DJ, Dixon JE. Dual
specificity phosphatases 18 and 21 target to opposing sides of the
mitochondrial inner membrane. J Biol Chem 283: 15440 –15450, 2008.
172. Raza H. Dual localization of glutathione S-transferase in the cytosol and
mitochondria: implications in oxidative stress, toxicity and disease.
FEBS J 278: 4243–4251, 2011.
173. Reed LJ, Damuni Z, Merryfield ML. Regulation of mammalian pyruvate and branched-chain alpha-keto acid dehydrogenase complexes by
phosphorylation-dephosphorylation. Curr Top Cell Regul 27: 41–49,
1985.
174. Rees DM, Leslie AG, Walker JE. The structure of the membrane
extrinsic region of bovine ATP synthase. Proc Natl Acad Sci USA 106:
21597–21601, 2009.
175. Reinders J, Sickmann A. State-of-the-art in phosphoproteomics. Proteomics 5: 4052–4061, 2005.
176. Rikova K, Guo A, Zeng Q, Possemato A, Yu J, Haack H, Nardone J,
Lee K, Reeves C, Li Y, Hu Y, Tan Z, Stokes M, Sullivan L, Mitchell
J, Wetzel R, Macneill J, Ren JM, Yuan J, Bakalarski CE, Villen J,
Kornhauser JM, Smith B, Li D, Zhou X, Gygi SP, Gu TL, Polakiewicz RD, Rush J, Comb MJ. Global survey of phosphotyrosine signaling identifies oncogenic kinases in lung cancer. Cell 131: 1190 –1203,
2007.
177. Roberts DL, Frerman FE, Kim JJ. Three-dimensional structure of
human electron transfer flavoprotein to 2.1-A resolution. Proc Natl Acad
Sci USA 93: 14355–14360, 1996.
178. Ruvolo PP, Deng X, Ito T, Carr BK, May WS. Ceramide induces Bcl2
dephosphorylation via a mechanism involving mitochondrial PP2A. J
Biol Chem 274: 20296 –20300, 1999.
179. Sale GJ, Randle PJ. Analysis of site occupancies in [32P]phosphorylated pyruvate dehydrogenase complexes by aspartyl-prolyl cleavage of
tryptic phosphopeptides. Eur J Biochem 120: 535–540, 1981.
180. Salvi M, Morrice NA, Brunati AM, Toninello A. Identification of the
flavoprotein of succinate dehydrogenase and aconitase as in vitro mitochondrial substrates of Fgr tyrosine kinase. FEBS Lett 581: 5579 –5585,
2007.
181. Salvi M, Stringaro A, Brunati AM, Agostinelli E, Arancia G, Clari G,
Toninello A. Tyrosine phosphatase activity in mitochondria: presence of
Shp-2 phosphatase in mitochondria. Cell Mol Life Sci 61: 2393–2404,
2004.
182. Salvi M, Brunati AM, Bordin L, La Rocca N, Clari G, Toninello A.
Characterization and location of Src-dependent tyrosine phosphorylation
in rat brain mitochondria. Biochim Biophys Acta 1589: 181–195, 2002.
183. Samavati L, Lee I, Mathes I, Lottspeich F, Huttemann M. Tumor
necrosis factor alpha inhibits oxidative phosphorylation through tyrosine
phosphorylation at subunit I of cytochrome c oxidase. J Biol Chem 283:
21134 –21144, 2008.
184. Sardanelli AM, Signorile A, Nuzzi R, Rasmo DD, TechnikovaDobrova Z, Drahota Z, Occhiello A, Pica A, Papa S. Occurrence of
A-kinase anchor protein and associated cAMP-dependent protein kinase
in the inner compartment of mammalian mitochondria. FEBS Lett 580:
5690 –5696, 2006.
185. Sardanelli AM, Technikova-Dobrova Z, Scacco SC, Speranza F,
Papa S. Characterization of proteins phosphorylated by the cAMPdependent protein kinase of bovine heart mitochondria. FEBS Lett 377:
470 –474, 1995.
186. Sardanelli AM, Technikova-Dobrova Z, Speranza F, Mazzocca A,
Scacco S, Papa S. Topology of the mitochondrial cAMP-dependent
protein kinase and its substrates. FEBS Lett 396: 276 –278, 1996.
187. Sawicki G, Jugdutt BI. Valsartan reverses post-translational modifications of the delta-subunit of ATP synthase during in vivo canine reperfused myocardial infarction. Proteomics 7: 2100 –2110, 2007.
188. Sazanov LA, Hinchliffe P. Structure of the hydrophilic domain of
respiratory complex I from Thermus thermophilus. Science 311: 1430 –
1436, 2006.
189. Schieven G, Martin GS. Nonenzymatic phosphorylation of tyrosine and
serine by ATP is catalyzed by manganese but not magnesium. J Biol
Chem 263: 15590 –15593, 1988.
190. Schilling B, Aggeler R, Schulenberg B, Murray J, Row RH, Capaldi
RA, Gibson BW. Mass spectrometric identification of a novel phosphorylation site in subunit NDUFA10 of bovine mitochondrial complex I.
FEBS Lett 579: 2485–2490, 2005.
191. Schmidt O, Harbauer AB, Rao S, Eyrich B, Zahedi RP, Stojanovski
D, Schönfisch B, Guiard B, Sickmann A, Pfanner N, Meisinger C.
AJP-Heart Circ Physiol • doi:10.1152/ajpheart.00077.2012 • www.ajpheart.org
Downloaded from http://ajpheart.physiology.org/ by 10.220.32.246 on May 5, 2017
156.
MITOCHONDRIA PROTEIN PHOSPHORYLATION
Review
MITOCHONDRIA PROTEIN PHOSPHORYLATION
192.
193.
194.
195.
196.
198.
199.
200.
201.
202.
203.
204.
205.
206.
207.
208.
209.
210.
211.
212.
213.
214. Vosseller K, Hansen KC, Chalkley RJ, Trinidad JC, Wells L, Hart
GW, Burlingame AL. Quantitative analysis of both protein expression
and serine / threonine post-translational modifications through stable
isotope labeling with dithiothreitol. Proteomics 5: 388 –398, 2005.
215. Walker JE, Dickson VK. The peripheral stalk of the mitochondrial ATP
synthase. Biochim Biophys Acta 1757: 286 –296, 2006.
216. Wang WL, Yeh SF, Chang YI, Hsiao SF, Lian WN, Lin CH, Huang
CY, Lin WJ. PICK1, an anchoring protein that specifically targets
protein kinase Calpha to mitochondria selectively upon serum stimulation in NIH 3T3 cells. J Biol Chem 278: 37705–37712, 2003.
217. Wang Z, Gucek M, Hart GW. Cross-talk between GlcNAcylation and
phosphorylation: site-specific phosphorylation dynamics in response to
globally elevated O-GlcNAc. Proc Natl Acad Sci USA 105: 13793–
13798, 2008.
218. Weishaupt A, Kadenbach B. Selective removal of subunit VIb increases the activity of cytochrome c oxidase. Biochemistry 31: 11477–
11481, 1992.
219. Wieland O, Jagow-Westermann B. ATP-dependent inactivation of
heart muscle pyruvate dehydrogenase and reactivation by Mg⫹⫹. FEBS
Lett 3: 271–274, 1969.
220. Wieland O, Siess E. Interconversion of phospho- and dephospho- forms
of pig heart pyruvate dehydrogenase. Proc Natl Acad Sci USA 65:
947–954, 1970.
221. Wiltshire C, Matsushita M, Tsukada S, Gillespie DA, May GH. A
new c-Jun N-terminal kinase (JNK)-interacting protein, Sab (SH3BP5),
associates with mitochondria. Biochem J 367: 577–585, 2002.
222. Wisniewski JR, Nagaraj N, Zougman A, Gnad F, Mann M. Brain
phosphoproteome obtained by a FASP-based method reveals plasma
membrane protein topology. J Proteome Res 9: 3280 –3289, 2010.
223. Wu QZ, Li DM, Kuang WH, Zhang TJ, Lui S, Huang XQ, Chan RC,
Kemp GJ, Gong QY. Abnormal regional spontaneous neural activity in
treatment-refractory depression revealed by resting-state fMRI. Hum
Brain Mapp 32: 1290 –1299, 2011.
224. Xue Y, Ren J, Gao X, Jin C, Wen L, Yao X. GPS 2.0: a tool to predict
kinase-specific phosphorylation sites in hierarchy. Mol Cell Proteomics
7: 1598 –1608, 2008.
225. Yadava N, Potluri P, Scheffler IE. Investigations of the potential effects
of phosphorylation of the MWFE and ESSS subunits on complex I
activity and assembly. Int J Biochem Cell Biol 40: 447–460, 2008.
226. Yaffe MB, Leparc GG, Lai J, Obata T, Volinia S, Cantley LC. A
motif-based profile scanning approach for genome-wide prediction of
signaling pathways. Nat Biotechnol 19: 348 –353, 2001.
227. Yang F, Stenoien DL, Strittmatter EF, Wang J, Ding L, Lipton MS,
Monroe ME, Nicora CD, Gristenko MA, Tang K, Fang R, Adkins
JN, Camp DG, Chen DJ, Smith RD. Phosphoproteome profiling of
human skin fibroblast cells in response to low- and high-dose irradiation.
J Proteome Res 5: 1252–1260, 2006.
228. Yu CA, Xia D, Kim H, Deisenhofer J, Zhang L, Kachurin AM, Yu L.
Structural basis of functions of the mitochondrial cytochrome bc1 complex. Biochim Biophys Acta 1365: 151–158, 1998.
229. Yu H, Lee I, Salomon AR, Yu K, Huttemann M. Mammalian liver
cytochrome c is tyrosine-48 phosphorylated in vivo, inhibiting mitochondrial respiration. Biochim Biophys Acta 1777: 1066 –1071, 2008.
230. Yu PH, Deng YL. Endogenous formaldehyde as a potential factor of
vulnerability of atherosclerosis: involvement of semicarbazide-sensitive
amine oxidase-mediated methylamine turnover. Atherosclerosis 140:
357–363, 1998.
231. Zanivan S, Gnad F, Wickstrom SA, Geiger T, Macek B, Cox J,
Fassler R, Mann M. Solid tumor proteome and phosphoproteome
analysis by high resolution mass spectrometry. J Proteome Res 7:
5314 –5326, 2008.
232. Zara V, Conte L, Trumpower BL. Biogenesis of the yeast cytochrome
bc1 complex. Biochim Biophys Acta 1793: 89 –96, 2009.
233. Zhang J, Duncker DJ, Ya X, Zhang Y, Pavek T, Wei H, Merkle H,
Ugurbil K, From AH, Bache RJ. Effect of left ventricular hypertrophy
secondary to chronicX pressure overload on transmural myocardial
2-deoxyglucose uptake. X A 31P NMR spectroscopic study. Circulation
92: 1274 –1283, 1995.
234. Zhang J, Murphy AN, Wiley SE, Perkins G, Worby CA, Engel JL,
Heacock P, Nguyen OK, Wang JH, Raetz CRH, Dowhan W, Dixon
JE. Mitochondrial phosphatase PTPMT1 is essential for cardiolipin
biosynthesis. Cell Metab 13: 690 –700, 2012.
AJP-Heart Circ Physiol • doi:10.1152/ajpheart.00077.2012 • www.ajpheart.org
Downloaded from http://ajpheart.physiology.org/ by 10.220.32.246 on May 5, 2017
197.
Regulation of mitochondrial protein import by cytosolic kinases. Cell
144: 227–239, 2011.
Schmitt ME, Trumpower BL. Subunit 6 regulates half-of-the-sites
reactivity of the dimeric cytochrome bc1 complex in Saccharomyces
cerevisiae. J Biol Chem 265: 17005–17011, 1990.
Schwoch G, Trinczek B, Bode C. Localization of catalytic and regulatory subunits of cyclic AMP-dependent protein kinases in mitochondria
from various rat tissues. Biochem J 270: 181–188, 1990.
Segal HL. Enzymatic interconversion of active and inactive forms of
enzymes. Science 180: 25–32, 1973.
Shaffer J, Adams JA. Detection of conformational changes along the
kinetic pathway of protein kinase A using a catalytic trapping technique.
Biochemistry 38: 12072–12079, 1999.
Sharma V, Abraham T, So A, Allard MF, McNeill JH. Functional
effects of protein kinases and peroxynitrite on cardiac carnitine palmitoyltransferase-1 in isolated mitochondria. Mol Cell Biochem 337: 223–
237, 2010.
Sheldon KL, Maldonado EN, Lemasters JJ, Rostovtseva TK, Bezrukov SM. Phosphorylation of voltage-dependent anion channel by serine/
threonine kinases governs its interaction with tubulin. PLoS One 6:
e25539, 2011.
Slater E, Kemp A. Rate of labelling of mitochondrial phosphohistidine
by radioactive inorganic phosphate. Nature 204: 1268 –1271, 1964.
Snyder CH, Merbitz-Zahradnik T, Link TA, Trumpower BL. Role of
the Rieske iron-sulfur protein midpoint potential in the protonmotive
Q-cycle mechanism of the cytochrome bc1 complex. J Bioenerg
Biomembr 31: 235–242, 1999.
Solmaz SR, Hunte C. Structure of complex III with bound cytochrome
c in reduced state and definition of a minimal core interface for electron
transfer. J Biol Chem 283: 17542–17549, 2008.
Sommarin M, Petit PX, Moller IM. Endogenous protein phosphorylation in purified plant mitochondria. Biochim Biophys Acta 1052: 195–
203, 1990.
Steinberg RA. Cyclic AMP-dependent phosphorylation of the precursor
to beta subunit of mitochondrial F1-ATPase: a physiological mistake? J
Cell Biol 98: 2174 –2178, 1984.
Stock D, Gibbons C, Arechaga I, Leslie AG, Walker JE. The rotary
mechanism of ATP synthase. Curr Opin Struct Biol 10: 672–679, 2000.
Struglics A, Fredlund KM, Konstantinov YM, Allen JF, Moller IM.
Protein phosphorylation/dephosphorylation in the inner membrane of
potato tuber mitochondria. FEBS Lett 475: 213–217, 2000.
Sun F, Huo X, Zhai Y, Wang A, Xu J, Su D, Bartlam M, Rao Z.
Crystal structure of mitochondrial respiratory membrane protein complex
II. Cell 121: 1043–1057, 2005.
Sun G, Budde RJ. Requirement for an additional divalent metal cation
to activate protein tyrosine kinases. Biochemistry 36: 2139 –2146, 1997.
Tal R, Winter G, Ecker N, Klionsky DJ, Abeliovich H. Aup1p, a yeast
mitochondrial protein phosphatase homolog, is required for efficient
stationary phase mitophagy and cell survival. J Biol Chem 282: 5617–
5624, 2007.
Tao WA, Wollscheid B, O’Brien R, Eng JK, Li XJ, Bodenmiller B,
Watts JD, Hood L, Aebersold R. Quantitative phosphoproteome analysis using a dendrimer conjugation chemistry and tandem mass spectrometry. Nat Methods 2: 591–598, 2005.
Technikova-Dobrova Z, Sardanelli AM, Papa S. Phosphorylation of
mitochondrial proteins in bovine heart. Characterization of kinases and
substrates. FEBS Lett 322: 51–55, 1993.
Technikova-Dobrova Z, Sardanelli AM, Speranza F, Scacco S, Signorile A, Lorusso V, Papa S. Cyclic adenosine monophosphatedependent phosphorylation of mammalian mitochondrial proteins: enzyme and substrate characterization and functional role. Biochemistry 40:
13941–13947, 2001.
Toogood HS, van Thiel A, Basran J, Sutcliffe MJ, Scrutton NS, Leys
D. Extensive domain motion and electron transfer in the human electron
transferring flavoprotein.medium chain Acyl-CoA dehydrogenase complex. J Biol Chem 279: 32904 –32912, 2004.
Tsukihara T, Aoyama H, Yamashita E, Tomizaki T, Yamaguchi H,
Shinzawa-Itoh K, Nakashima R, Yaono R, Yoshikawa S. The whole
structure of the 13-subunit oxidized cytochrome c oxidase at 2.8 A.
Science 272: 1136 –1144, 1996.
Villen J, Beausoleil SA, Gerber SA, Gygi SP. Large-scale phosphorylation analysis of mouse liver. Proc Natl Acad Sci USA 104: 1488 –
1493, 2007.
H965
Review
H966
MITOCHONDRIA PROTEIN PHOSPHORYLATION
235. Zhang J, Frerman FE, Kim JJ. Structure of electron transfer flavoprotein-ubiquinone oxidoreductase and electron transfer to the mitochondrial ubiquinone pool. Proc Natl Acad Sci USA 103: 16212–16217, 2006.
236. Zhao X, León IR, Bak S, Mogensen M, Wrzesinski K, Højlund K,
Jensen ON. Phosphoproteome analysis of functional mitochondria isolated
from resting human muscle reveals extensive phosphorylation of inner membrane protein complexes and enzymes. Mol Cell Proteomics 10: 1–14, 2011.
237. Zhou L, Cabrera ME, Huang H, Yuan CL, Monika DK, Sharma N,
Bian F, Stanley WC. Parallel activation of mitochondrial oxidative
metabolism with increased cardiac energy expenditure is not dependent
on fatty acid oxidation in pigs. J Physiol 579: 811–821, 2007.
238. Zu XL, Besant PG, Imhof A, Attwood PV. Mass spectrometric
analysis of protein histidine phosphorylation. Amino Acids 32: 347–
357, 2007.
Downloaded from http://ajpheart.physiology.org/ by 10.220.32.246 on May 5, 2017
AJP-Heart Circ Physiol • doi:10.1152/ajpheart.00077.2012 • www.ajpheart.org