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Impact of Mitochondrial Fatty Acid Synthesis on Mitochondrial Biogenesis
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Minireview Impact of Mitochondrial Fatty Acid Synthesis on Mitochondrial Biogenesis Sara M. Nowinski1, Jonathan G. Van Vranken1, Katja K. Dove1, and Jared Rutter1,2,* 1Department
of Biochemistry, University of Utah School of Medicine, Salt Lake City, UT 84132, USA Hughes Medical Institute, University of Utah School of Medicine, Salt Lake City, UT 84132, USA *Correspondence: [email protected] https://doi.org/10.1016/j.cub.2018.08.022 2Howard
Biology students today are taught that mitochondria are ‘the powerhouse of the cell’. This gross over-simplification of their cellular role has arguably led to a paucity of knowledge concerning the many other tasks carried out by this multifunctional organelle. Mitochondrial fatty acid synthesis (mtFAS) is one such underappreciated pathway that is crucial for mitochondrial function, although even mitochondrial experts are often surprised to learn of its existence. For many years, the only function of mtFAS was thought to be the production of lipoic acid, an important co-factor for several mitochondrial enzymes. However, recent advances have revealed a far wider role for mtFAS in mitochondrial physiology. The discovery of human patients with mutations in mtFAS enzymes has brought renewed interest in understanding the full significance of this novel mode of mitochondrial metabolic regulation. We now appreciate that mtFAS is a nutrient-sensitive pathway that provides an elegant mechanism whereby acetyl-CoA regulates its own consumption via coordination of lipoic acid synthesis and tricarboxylic acid (TCA) cycle activity, iron–sulfur (FeS) cluster biogenesis, assembly of oxidative phosphorylation complexes, and mitochondrial translation. In this minireview, we describe and build upon the important discoveries that led to our current understanding of this elegant mechanism of coordination of nutrient status and metabolism. Introduction Fatty acid synthesis (FAS) is an essential process in organisms from bacteria to mammals. FAS has a number of cellular roles, including the production of triglycerides as lipid storage units, the generation of phospholipids to build cell membranes, and many more. However, it is a little-known fact that eukaryotes actually harbor two distinct FAS systems — one in the cytoplasm, and one in mitochondria. Although mitochondrial fatty acid synthesis (mtFAS) was discovered decades ago, the importance of this pathway has been underestimated compared with its more famous cytoplasmic cousin FASN. The recent finding that some patients with neurogenerative disease harbor mutations in a mtFAS enzyme has generated new interest in understanding the cellular role(s) played by this ‘old’ pathway [1]. Unlike FASN, mtFAS does not appear to contribute significantly to cellular triglyceride stores or phospholipids, but is instead responsible for the generation of lipoic acid, an important lipid cofactor for multiple mitochondrial dehydrogenases. Recent studies have now implicated mtFAS in a range of mitochondrial processes beyond its classical role in lipoic acid production, from mitochondrial translation to the support and coordination of mitochondrial oxidative phosphorylation. Herein, we follow the evolution of our understanding of the mtFAS pathway and the processes it governs. Discovering mtFAS FAS is a complex process whereby saturated fatty acids are built stepwise in two carbon units, the addition of which is mediated by several enzymatic activities. Most bacteria utilize a dissociated pathway in which separate enzymes catalyze each step in the elongation of fatty acyl chains on a soluble acyl carrier protein
(ACP) scaffold. In contrast, eukaryotes evolved more consolidated systems that encode all of the necessary enzymatic activities on one (mammals) or two (yeast) peptides (Figure 1 and Table 1). These systems are termed type II and type I fatty acid synthesis, respectively, and all organisms were historically thought to exclusively utilize one pathway or the other, with few exceptions. However, reports claiming to observe fatty acid synthesis in mammalian mitochondria (mtFAS) were published in the 1960s to much skepticism. Real progress towards understanding this pathway did not truly come until the late 1980s, when the eukaryotic mitochondrial ACP was discovered in Neurospora crassa by Stuart Brody and Steve Mikolajczyk while they were looking for pantetheine-modified proteins [2]. A highly conserved serine residue on ACP is modified with a 4’-phosphopantetheine (4’-PP) cofactor, which supplies the terminal thiol group upon which fatty acids are built. This seminal study identified the primary acyl chain on ACP as the 14-carbon lipid 3-hydroxymyristate; however, the source of the acyl chain was at first unknown. Was it made by cytoplasmic FASN or by a novel synthesis pathway in the mitochondria? De novo mtFAS in N. crassa was described two years later, with a series of in vitro experiments that led the researchers to conclude that myristic acid (14:0) was the ‘product’ of mtFAS, while in vivo experiments again yielded ACP with labelled hydroxymyristic acid (14:0-OH) [3]. Initially, the primary hypothesis regarding the significance of this pathway was that mtFAS likely contributed to the synthesis of mitochondrial phospholipids. Soon after the initial discovery of ACP and mtFAS, efforts by Hanns Weiss’s lab to characterize complex I of the mitochondrial electron transport chain (ETC) in N. crassa also discovered ACP as a stable subunit of this complex [4]. Around the same time,
R1212 Current Biology 28, R1212–R1219, October 22, 2018 ª 2018 Elsevier Ltd.
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Minireview FAS I (yeast) FAS II (mtFAS)
O
ACT
ER
DH
AT
ACP
KR
ACT?
ER
DH
AT
ACP
KR
ACC
OH
CH3 C CoA
CH3 (CH2)N CH CH2 C S
Malonyl-CoA
Initiation
AT
H2O
O
O
HO C CH2 C S
ACP
KR
ACT/KS
CO2 O
O
NADPH
CoA CoA
PPT
3-hydroxyacyl-ACP
NADP
+
acetyl-CoA
PPT
KS
O
O
HO C CH2 C CoA
KS
O
O
CH3 (CH2)N C CH2 C S
ACP
Malonyl-ACP
DH
CH3 (CH2)N CH CH C S
ACP
ACP
trans -2-enoyl-ACP
3-ketoacyl-ACP
ER HS
HS
ACP
O
ACP CO2
Holo-ACP
Holo-ACP
Elongation
KS
Apo-ACP O C
S
CH2 CH2
O
N
C
CH2 CH2
CH3 (CH2)N C S
ACP
Octanoyl-ACP H
CH3
C
C
C
O
OH CH3
H
H
O
Acyl chain
NADP+
O CH3 (CH2)6 C S
Lipoic acid and lipoylated proteins
ACP
CH3 (CH2)N
ACP
Acyl-ACP
4’-phosphopantetheine
PPT
NADPH
CH3 (CH2)N C S
N
ACP
Acyl-ACP
OCH2 O P O
4’-phosphopantetheine
ACP
Substrate
O
Ser
ACP
Apo-ACP No 4’-PP
Products
HS
ACP
Holo-ACP 4’-PP Current Biology
Figure 1. Fatty acid synthesis is catalyzed by a series of conserved enzymatic activities in the eukaryotic cytoplasm and mitochondria. (Top) A schematic representing the different subunits of yeast FAS I and FAS II. In mammals, components of FAS I are contained on one polypeptide, whereas yeast FAS I comprises two polypeptides (black boxes). On the contrary, members of FAS II are distinct entities in bacteria, yeast and mammals. (Middle) Mitochondrial fatty acid synthesis pathway. Enzymes corresponding to each of the domain/protein abbreviations can be found in Table 1. (Bottom left) Chemical structure of the ACP prosthetic group 4’-phosphopantetheine covalently bound to a conserved serine is shown. (Bottom right) Key for the FAS pathway shown in the middle section.
ACP was found in complex I from bovine heart, marking its first description in mammalian mitochondria [5]. Both studies suggested that the majority of ACP in complex I was acylated, while unbound ACP was mostly de-acylated, and it was therefore postulated that complex I was required for mtFAS. Many other studies describing mitochondrially localized ACP in various organisms followed [6–9], culminating with its eventual discovery in humans [10]. Despite many advances in our understanding of this system, several pertinent questions have remained unanswered until today. What is the acyl chain modification on ACP and/or the major product of mtFAS? Why is ACP associated with complex I and what are the implications of this interaction? What is the purpose of mtFAS, and why has this pathway been evolutionarily conserved when it is known that fatty acids can be imported into mitochondria from the endoplasmic reticulum or cytosol? Recent publications discussed below have made significant advances towards answering these questions and have identified important trajectories for future research.
Defining the Product(s) of mtFAS In the decade following the initial discovery of mtFAS and ACP, a major research goal was to define the ultimate product(s) of mtFAS. Using an N. crassa strain with defective cytoplasmic FAS, the Weiss lab again found 3-hydroxymyristate (14:0) and 3-hydroxylaurate (12:0) as the major products [11]. While these early studies reproducibly identified a 14-carbon lipid as the major product of mtFAS, there was mixed opinion over the cellular role that this lipid was likely to play. While earlier studies had proposed that mtFAS was expected to contribute generally to mitochondrial membrane phospholipids, Weiss and colleagues disagreed, citing the existing knowledge that most mitochondrial phospholipids are imported from the endoplasmic reticulum [12]. They proposed a narrower hypothesis: that mtFAS supplied acyl chains specifically for cardiolipin biosynthesis within the mitochondria. The identification of genes encoding ACP (acp-1, ACP1) in a number of organisms enabled knockout studies aimed at answering this question [13]. The initial Current Biology 28, R1212–R1219, October 22, 2018 R1213
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Minireview Table 1. Fatty acid synthesis is a conserved pathway in bacteria, yeast, and mammals. FAS II gene names Domain/protein
Function
E. coli
S. cerevisiae
H. sapiens
ACC
Acetyl-CoA carboxylase
accA, accB/fabE, accC, accD
HFA1?
ACC1? (ACSF3?)
ACP
Acyl carrier protein
accP
ACP1
NDUFAB1
ACT
Acetyltransferase
fabH
?
?
AT
Acyltransferase
fabD
MCT1
MCAT
DH
Dehydratase
fabA, fabZ
HTD2
HTD2
ER
Enoyl reductase
fabI, fabK, fabL, fabV
ETR1
MECR
KR
Ketoreductase
fabG
OAR1
HSD17B8, CBR4
KS
Ketosynthase
fabB, fabF
CEM1
OXSM
PPT
Phosphopantetheinyl-transferase
acpS
PPT2
AASDHPPT?
Enzyme activities and the corresponding genes that encode this activity in E. coli, S. cerevisiae, and H. sapiens are listed. ? indicates areas of uncertainty and/or ongoing research. ACSF3 is listed in parentheses next to ACC1 because it has been implicated in the generation of malonyl-CoA in mammalian mitochondria but is a malonyl-CoA synthetase rather than an acetyl-CoA carboxylase.
knockout studies were performed simultaneously in N. crassa and Saccharomyces cerevisiae [14]. Knockout of acp-1 in N. crassa ablated the peripheral module of complex I and resulted in improper assembly of the membrane module and a lack of incorporation of iron–sulfur (FeS) clusters into complex I. However, respiration was otherwise unaffected, and no differences in mitochondrial phospholipids were observed. In contrast, knockout of ACP1 in S. cerevisiae resulted in a much more severe ‘‘pleiotropic respiratory-deficient phenotype’’. In these mutants, mitochondrial phospholipids did show some changes, though none that are not observed in other respiratory-deficient mutants (rho– or rho0 ‘petite’ mutants). The severity of the phenotype and similarity to other rho0 mutants led the researchers to conclude that these cells were rho– or rho0, and that the observed loss of ETC complexes and changes in mitochondrial lipids were likely indirect. Because mitochondrial phospholipids were largely unaffected by knockout of ACP in N. crassa, these first genetic studies seeded the idea that perhaps ACP was involved in the delivery of a specific product essential for the proper formation of the membrane module of complex I. However, this way of thinking changed a few years later in 1997, when it was discovered that ACP was required for the synthesis of lipoic acid, an important cofactor for several mitochondrial dehydrogenases, including pyruvate dehydrogenase (PDH), a-ketoglutarate dehydrogenase (a-KGDH), and the glycine cleavage system [15–17]. Shedding some doubt on the phenotypic findings of earlier ACP-knockout studies, the Schweizer lab reported that ACP1 loss was often lethal in S. cerevisiae and proposed that surviving mutants had activated some compensatory mechanism [15]. Since lipoic acid is only an eight-carbon lipid, and previous studies had all identified 3-hydroxymyristic acid as the major acyl chain found covalently linked to ACP, it remained likely that longer acyl products of mtFAS played some other cellular role [15]. However, with these findings the focus of the mtFAS field completely shifted toward the study of lipoic acid production. The possible role(s) of longer fatty acids synthesized on ACP and of ACP in ETC assembly and function were largely forgotten and would not be revisited for nearly two decades. R1214 Current Biology 28, R1212–R1219, October 22, 2018
Identifying the mtFAS Enzymes The discovery of ACP and de novo mitochondrial fatty acid biosynthesis started a race to identify the mtFAS enzymes and the genes that encode them. Prokaryotic (type II) FAS is catalyzed by at least five enzymes. For each two-carbon increase in acyl chain length, these enzymes catalyze the addition of malonyl-CoA to ACP, the condensation of malonyl-ACP with a growing acyl chain forming a keto-acyl-ACP product, and the subsequent reduction of that keto group to a methylene group, completing the elongation cycle (Figure 1). The first eukaryotic mtFAS gene to be discovered in yeast was CEM1, which was identified as the enzyme that performs the condensation reaction in 1993 by virtue of its sequence homology to the bacterial b-keto-acyl synthase, at a time before the identification of lipoic acid as a product of mtFAS [18]. Null cem1 mutants are respiratory deficient but show no major differences in mitochondrial lipids and fatty acids, an observation that is consistent with findings from non-lethal ACP1 knockouts. The genes encoding two more yeast mtFAS enzymes, MCT1 and OAR1, were also identified via homology to their respective bacterial acyltransferase and ketoreductase counterparts, and mutation of these genes led to similar phenotypes as those of acp1 and cem1 mutants [19]. The last two S. cerevisiae genes proved to be more elusive. ETR1 was originally identified as MRF1, a protein that was purified from nuclear extracts and shown to bind to autonomously replicating sequences of nuclear DNA [20]. It was eventually identified as an enoyl thioester reductase by homology to an enzyme from Candida tropicalis and by complementation of mutants by a mitochondria-targeted version of the Escherichia coli FabI enzyme. Re-investigation of its localization revealed that it was found predominantly in mitochondria and that mitochondrial localization was essential for rescue of the petite phenotype in etr1 mutants [21]. The dehydratase encoded by HTD2, on the other hand, was discovered via a screen that used a mitochondria-targeted form of the E. coli FabA gene (Table 1) [22]. htd2 mutant phenotypes were identical to those observed for other mtFAS mutants, indicating that the resultant defects were all likely the result of a failure to acylate ACP. In support of this idea, re-localization of a peroxisomal fatty acyl-CoA synthetase to mitochondria rescued cem1 mutants, presumably by
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Minireview re-acylating ACP [23]. Identification of most of the corresponding mammalian genes came about a decade later and added a few mechanistic insights, showing that these enzymes can accommodate chain lengths out to C14–C16 in vitro [24–29]. While the major catalytic enzymes of the mtFAS pathway are now defined, a number of accessory enzymes important for mtFAS are less well studied. One such enzyme is Ppt2, the mitochondrial phosphopantetheinyl transferase that catalyzes the addition of the 4’-PP moiety from coenzyme A to the invariant serine residue on ACP [30]. Interestingly, mammals appear to have only a cytoplasmic version of phosphopantetheinyl transferase [24], raising the question of whether the 4-phosphopantetheine is added in the cytosol and translocated into the mitochondria, or whether there is an undiscovered mitochondrial phosphopantetheinyl transferase. Similarly, mitochondrial malonyl-CoA is a required substrate of mtFAS, but the identity of a mitochondrial acetyl-CoA carboxylase (ACC) enzyme that would synthesize malonyl-CoA from acetyl-CoA is still uncertain. Originally, researchers proposed that the cytosolic ACC enzyme Acc1 might fulfil this function, but no mechanism for malonylCoA import has been described. Despite this, mutants with defects in Acc1 activity are respiratory deficient and reportedly have a 50% reduction in lipoic acid levels [31]. The same lab later identified Hfa1 as the mitochondrial ACC; however, hfa1D strains are respiration competent under normal growth conditions, indicating that mtFAS is still functional and that there is at least an additional source of mitochondrial malonyl-CoA in these cells ([32,33] and our unpublished data, J.G. Van Vranken). In mammals, there is some evidence that ACC1 and ACSF3 (a mammalian malonyl-CoA synthetase) might work in a coordinated fashion [34]. Regardless, the source of mitochondrial malonyl-CoA remains up for debate. Beginning to Put It Together Although early studies largely agreed on the phenotypes of mtFAS knockouts, it was unclear what mechanisms underlied their pleiotropic respiratory-deficient phenotype, and whether these effects were a result of the loss of lipoic acid, or some other function of mtFAS. Several possibilities have been proposed. While some studies claimed that ACP-deficient cells were rho0, these cells were genetically rescuable, indicating that they retained at least some mitochondrial DNA [14]. Subsequent reports clarified that mtFAS mutants in fact had no defects in mitochondrial DNA replication under normal growth conditions [18,20,23,33]. Another possibility was defective mitochondrial translation. Although early studies identifying mtFAS enzymes reported no defects in mitochondrial translation, our lab and others have measured translational deficiencies in mtFAS mutants ([33,35] and our unpublished data, Sara M. Nowinski and Olga Zurita Rendon). One study identified a requirement for mtFAS in the processing of the 5’ ends of mitochondrial tRNAs by RNase P, with mtFAS mutants having profound defects in the synthesis of mitochondrial tRNAs [33]. Moreover, the authors show that lipoic acid synthesis mutants do not exhibit the same defects as mtFAS mutants, implying that the activation of RNase P may be one role for the longer acyl chains synthesized on ACP. However, the mechanism(s) whereby mtFAS is required for RNase P function remain unknown. Yeast RNase P is composed of one nuclear-encoded
peptide (Rpm2) and one mitochondria-encoded RNA (RPM1), which is itself processed by RNase P to its mature form, while human mitochondrial RNase P is a heterotrimer of three protein subunits, none of which resembles Rpm2. Thus, it is unlikely that the mechanisms underlying RNase P activity are conserved in mammalian systems, although other mechanisms to connect mtFAS and translation may have evolved. The mammalian HTD2 and RPP14 (a mammalian nuclear RNase P subunit) are encoded by the same mRNA transcript [28]. Furthermore, a recent structure of a mitochondrial ribosome assembly intermediate identified ACP bound directly to the mitochondrial ribosome [36]; however, the functional significance of these observations remains to be seen. To this point, previous studies had identified two major routes whereby mtFAS regulates mitochondrial metabolism: via lipoic acid production and mitochondrial translation. This creates two positive feedback loops in mitochondria. In the first, PDH converts pyruvate to acetyl-CoA using its lipoylated domain; some of this acetyl-CoA enters the tricarboxylic acid (TCA) cycle, while some becomes the substrate for the mtFAS pathway, ultimately producing more lipoic acid. In the second, RNase P processes its own RPM1 precursor RNA to form more active RNase P and enable mitochondrial translation. Both of these processes are tied to the requirement for a mtFAS product (lipoic acid in the case of PDH, unknown in the case of RNase P) for their activity. Hiltunen and Kastaniotis therefore proposed the insightful speculation in the absence of confirmatory data that these pathways might regulate mitochondrial function in response to nutrient status [37]. A subsequent publication from the same lab focused on etr1D cells and showed that these mutants have low respiration, low activities of ETC complexes, and low membrane potential [35]. They attributed this phenotype to a reduction in steady-state levels of ETC complexes II, III, and IV subunits but not complex V. However, their proposed model hinged on observed effects on mitochondrial RNA splicing, which cannot explain the effects of mtFAS loss on complex II assembly and function because it is encoded entirely in the nucleus, indicating that some other mechanism must still be at play. Finally, this publication also postulated that because acetyl-CoA is the substrate of the TCA cycle and mtFAS, this pathway is a mechanism by which mitochondria sense cellular metabolic status and control mitochondrial biogenesis [35]. Although studies had speculated that ETC assembly defects might underlie the profound mitochondrial defects observed in mtFAS mutants, none of these phenotypes could explain the observation first made by Brody et al. that ACP1 is an essential gene in many genetic backgrounds [15,33]. Therefore, our lab set out to identify the essential function of ACP. Proteomics studies had identified physical interactions between ACP and Isd11, a protein that is part of the mitochondrial FeS cluster biogenesis machinery [38–41]. We found that ACP is an essential and stable subunit of the FeS cluster biosynthetic complex, where it physically interacts with Isd11 and enables FeS cluster biosynthesis in an acylation-dependent fashion [41]. Subsequently, two structures confirmed this interaction, and showed an acyl modification on ACP that is embedded in the three-helix bundle formed by Isd11 [42,43] (Figure 2). These studies reported density in the crystal structure that could correspond to at least a 12-carbon acyl chain, but Isd11 could potentially accommodate longer Current Biology 28, R1212–R1219, October 22, 2018 R1215
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Minireview ACP–ISD11 iron sulfur cluster biogenesis
ACP–NDUFB9 complex I
ACP–NDUFA6 complex I
ACP is acylated
ACP–L0R8F8 (novel LYR) ribosome assembly intermediate
ACP is not acylated Current Biology
Figure 2. ACP–LYR protein interactions suggest an important role for ACP acylation. Cartoon representations of ACP bound to LYR proteins in different contexts. ACP is colored blue in all structures. (Left) The interaction between ACP and Isd11 (yellow, PDB 5USR), a LYR protein involved in iron–sulfur cluster biogenesis, intimately involves an acyl chain (grey spheres) covalently bound to ACP through 4’-phosphopantetheine (4’-PP, grey spheres). ACP–LYR interactions involved in electron transport chain (ETC) assembly also show acylation of ACP, as depicted by interactions with two LYR proteins in ETC complex I, NDUFB9 (green, PDB 5XTH) and NDUFA6 (pink, PDB 5XTH). (Right) A recent structure of the mitochondrial ribosome revealed a novel LYR protein L0R8F8 (light blue, PDB 5OOL) to which ACP is bound, although in this structure ACP is not acylated.
chains, and purification of the complex followed by fatty acid methyl ester gas chromatography mass spectrometry (FAME GC-MS) analysis found primarily 16-carbon acyl chains. Acylated ACP Directly Coordinates Mitochondrial Metabolism with Nutrient Status Concurrent with the identification of the ACP–Isd11 interaction and corresponding structures, several proteomic and structural studies identified interactions between ACP and a family of (mostly) late-stage ETC assembly factors known as the LYR proteins, of which Isd11 is a member [36,38–47]. The LYR proteins are so-called because each member contains a degenerate Leu–Tyr–Arg (LYR) motif, which is required for interaction of these proteins with ACP [48]. In addition to Isd11, the yeast LYR family includes Sdh6, Sdh7, Mzm1, and Fmc1, which facilitate the last steps in the assembly of ETC complexes II (Sdh6/ Sdh7), III (Mzm1), and V (Fmc1) (Figure 3; reviewed in [38]). Mammals and fungi that have ETC complex I also express two additional LYR protein–ACP dimers that are stable subunits of ETC complex I [38]. Additionally, mammalian genomes encode at least five additional LYR proteins that are largely uncharacterized. In agreement with the very early studies identifying ACP in complex I, recent studies show that ACP is nearly uniformly acylated when bound to LYR proteins, and moreover that these acyl chains appear to be longer (C12–C14) than those used as precursors for lipoic acid synthesis (C8), although the exact identity of the acyl chain is still debated [49,50]. Beyond providing a potential mechanistic link to ETC assembly, the structural studies that identified ACP–LYR interactions hinted at an intimate role for ACP acylation in these complexes, because like Isd11, the acyl chain threads into the middle of a three-helix bundle in each LYR (Figure 2) [42–45]. Interestingly, a recent structure of ACP bound to a LYR protein as part of a ribosome assembly intermediate reveals that ACP is not acylated, implying that the acyl chain is not favored in all ACP–LYR interactions (Figure 2) [36,48]. The obvious question that followed these obR1216 Current Biology 28, R1212–R1219, October 22, 2018
servations was whether acylation was important for the function of these interactions. A recent publication from our group describes the mechanism whereby mtFAS, via ACP acylation and interaction with LYR proteins, controls the assembly of mitochondrial ETC complexes in S. cerevisiae [48] (Figure 3). LYR proteins preferentially interact with acylated ACP (Figure 2), and this interaction is crucial for the ability of the LYR proteins to function in ETC complex assembly. Moreover, we showed that this process is dependent on mitochondrial acetyl-CoA availability. mtFAS and ACP acylation therefore indeed can be thought of as a ‘sensor’ for mitochondrial acetyl-CoA levels as Hiltunen and Kastaniotis proposed [35,37]. This system theoretically provides a mechanism to integrate most major mitochondrial processes with nutrient levels, as ACP acylation affects mitochondrial translation via RNase P, FeS cluster biogenesis via Isd11, ETC assembly via the other LYRs, and finally, TCA cycle and 1-carbon metabolism via lipoic acid production. Mitochondrial acetyl-CoA is, of course, not just the substrate for mtFAS, but is much better known for its role as a major substrate for the TCA cycle, where it is oxidized to generate the reduced forms of nicotinamide adenine dinucleotide (NADH) and flavin adenine dinucleotide (FADH2), which feed reducing equivalents to the ETC, ultimately fueling the production of mitochondrial ATP. As such, acetyl-CoA elegantly regulates its own metabolism through mtFAS, tying mitochondrial acetyl-CoA availability to the activity of TCA cycle enzymes and ETC oxidative capacity (Figure 3). Conclusions The story of mtFAS has evolved slowly over the course of several decades, leading to our current appreciation of its multifaceted role in mitochondrial metabolism. This pathway, largely ignored by the mitochondrial research community, has now returned to the limelight of mitochondrial metabolism. From the moment of the discovery of mitochondrial ACP and de novo mtFAS, the purpose and cellular function of this pathway has
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Minireview Figure 3. The mtFAS pathway is a nutrientsensitive coordinator of mitochondrial oxidative phosphorylation.
ACP Acetyl-CoA
mtFAS AcylACP
OctanoylACP
?
Lipoic acid
TCA cycle
Mitochondrial translation
LYR network
NADH/FADH 2
AcylACP
LYR
FeS
II
III
IV
I
V
OXPHOS IMS
(Top) Schematic overview of the integrated role of mtFAS in the coordination of mitochondrial nutrient supply and metabolism. Acetyl-CoA feeds the TCA cycle and mtFAS pathway, where it fuels the generation of octanoyl-ACP as well as longer acyl chains (acyl-ACP). Octanoyl-ACP is the precursor for the synthesis of lipoic acid, which is an important cofactor for TCA cycle and other mitochondrial enzymes. Acyl-ACP supports oxidative phosphorylation via two mechanisms: an undefined role in mitochondrial translation, as well as its interaction with the LYR protein network. (Bottom) In complex with Acyl-ACP, LYRs support the assembly of ETC complexes and the iron–sulfur (FeS) cluster biogenesis machinery. Specifically, Sdh6 and Sdh7 (SDHAF1 and SDHAF3) facilitate the assembly of complex II, Mzm1 (LYRM7) assembles complex III, Fmc1 supports assembly of complex V, and Isd11 (LYRM4) is an essential subunit of an FeS complex biosynthetic complex. In mammals, two additional LYR proteins (NDUFA6 and NDUFB9) in complex with acylACP are stable subunits of complex I.
C Q
I
II
III
IV
NDUFB9
Matrix
NDUFA6
AcylACP Sdh6
AcylACP AcylACP FeS Isd11 complex
AcylACP
Mzm1 Sdh7 AcylACP
AcylACP
been under debate. Although early publications speculated about a function for mtFAS in mitochondrial phospholipid and/or cardiolipin production, it appears that fatty acids produced by this pathway play much more specific roles. The discoveries of Brody et al. [15] and Wada et al. [17] defined the role of ACP in lipoic acid production, arguably one of its most important cellular functions. However, it has been clear from the beginning that longer acyl chains are synthesized on ACP. In the last five years, research advances have allowed us to finally understand the cellular role of these other mtFAS products in the regulation of mitochondrial translation, FeS cluster biogenesis, and ETC assembly. Through its specific interaction with the LYR proteins, acylation of ACP provides the cell with a means to coordinate acetyl-CoA concentration with the ability to metabolize acetyl-CoA. While this elegant regulatory mechanism and the purpose of longer acyl chains on ACP have now been described, many interesting questions remain. What is the utility of maintaining such a complex system for monitoring acetyl-CoA and coordinating mitochondrial function throughout evolution, especially given that targeting a single fatty acyl-CoA synthetase (Faa2) to the mitochondrial matrix can circumvent this process [23,48]? While mitochondrial acetyl-CoA is clearly required for mtFAS and ACP
acylation, the link between acetyl-CoA and malonyl-CoA in the mitochondria is not well established. To what extent is the acyl-ACP–LYR regulatory network conserved in higher eukaryotes? Given Fmc1 that two mammalian LYR proteins are Acylstable subunits of ETC complex I, ACP Fmc1 insights into the functional significance and regulation of these interactions may prove particularly interesting. And finally, Current Biology beyond the level of substrate regulation, how is this pathway controlled? It will be exciting to follow the discoveries as these and other questions guide future studies aimed at understanding the importance of this elegant mechanism of metabolic regulatory control.
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ACKNOWLEDGEMENTS We would like to acknowledge the many groups who contributed to the studies defining the functions of the LYR proteins reviewed in [38], but could not be cited due to space constraints. Related work in the lab of the authors was partially supported by grants to J.R. from the NIH (GM115174 and GM115129), the Nora Eccles Treadwell Foundation, and the Howard Hughes Medical Institute. S.M.N. was also supported by postdoctoral fellowships from UMDF and ACS. REFERENCES 1. Heimer, G., Keratar, J.M., Riley, L.G., Balasubramaniam, S., Eyal, E., Pietikainen, L.P., Hiltunen, J.K., Marek-Yagel, D., Hamada, J., Gregory, A., et al. (2016). MECR mutations cause childhood-onset dystonia and optic atrophy, a mitochondrial fatty acid synthesis disorder. Am. J. Hum. Genet. 99, 1229–1244. 2. Brody, S., and Mikolajczyk, S. (1988). Neurospora mitochondria contain an acyl-carrier protein. Eur. J. Biochem. 173, 353–359. 3. Mikolajczyk, S., and Brody, S. (1990). De novo fatty acid synthesis mediated by acyl-carrier protein in Neurospora crassa mitochondria. Eur. J. Biochem. 187, 431–437.
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Minireview 4. Sackmann, U., Zensen, R., Rohlen, D., Jahnke, U., and Weiss, H. (1991). The acyl-carrier protein in Neurospora crassa mitochondria is a subunit of NADH:ubiquinone reductase (complex I). Eur. J. Biochem. 200, 463–469.
22. Kastaniotis, A.J., Autio, K.J., Sormunen, R.T., and Hiltunen, J.K. (2004). Htd2p/Yhr067p is a yeast 3-hydroxyacyl-ACP dehydratase essential for mitochondrial function and morphology. Mol. Mcirobiol. 53, 1407–1421.
5. Runswick, M.J., Fearnley, I.M., Skehel, J.M., and Walker, J.E. (1991). Presence of an acyl carrier protein in NADH:ubiquinone oxidoreductase from bovine heart mitochondria. FEBS Lett. 286, 121–124.
23. Harington, A., Schwarz, E., Slonimski, P.P., and Herbert, C.J. (1994). Subcellular relocalization of a long-chain fatty acid CoA ligase by a suppressor mutation alleviates a respiration deficiency in Saccharomyces cerevisiae. EMBO J. 13, 5531–5538.
6. Chuman, L., and Brody, S. (1989). Acyl carrier protein is present in the mitochondria of plants and eucaryotic micro-organisms. Eur. J. Biochem. 184, 643–649. 7. Wilson, R., Ainscough, R., Anderson, K., Baynes, C., Berks, M., Bonfield, J., Burton, J., Connell, M., Copsey, T., Cooper, J., et al. (1994). 2.2 Mb of contiguous nucleotide sequence from chromosome III of C. elegans. Nature 368, 32–38. 8. Shintani, D.K., and Ohlrogge, J.B. (1994). The characterization of a mitochondrial acyl carrier protein isoform isolated from Arabidopsis thaliana. Plant Physiol. 104, 1221–1229. 9. Ragone, G., Caizzi, R., Moschetti, R., Barsanti, P., De Pinto, V., and Caggese, C. (1999). The Drosophila melanogaster gene for the NADH:ubiquinone oxidoreductase acyl carrier protein: developmental expression analysis and evidence for alternatively spliced forms. Mol. Gen. Genet. 261, 690–697. 10. Triepels, R., Smeitink, J., Loeffen, J., Smeets, R., Buskens, C., Trijbels, F., and van den Heuvel, L. (1999). The human nuclear-encoded acyl carrier subunit (NDUFAB1) of the mitochondrial complex I in human pathology. J. Inherit. Metab. Dis. 22, 163–173. 11. Schneider, R., Brors, B., Massow, M., and Weiss, H. (1997). Mitochondrial fatty acid synthesis: a relic of endosymbiontic origin and a specialized means for respiration. FEBS Lett. 407, 249–252. 12. Daum, G. (1985). Lipids of mitochondria. Biochim. Biophys. Acta 822, 1–42. 13. Cheret, G., Mattheakis, L.C., and Sor, F. (1993). DNA sequence analysis of the YCN2 region of chromosome XI in Saccharomyces cerevisiae. Yeast 9, 661–667. 14. Schneider, R., Massow, M., Lisowsky, T., and Weiss, H. (1995). Different respiratory-defective phenotypes of Neurospora crassa and Saccharomyces cerevisiae after inactivation of the gene encoding the mitochondrial acyl carrier protein. Curr. Genet. 29, 10–17. 15. Brody, S., Oh, C., Hoja, U., and Schweizer, E. (1997). Mitochondrial acyl carrier protein is involved in lipoic acid synthesis in Saccharomyces cerevisiae. FEBS Lett. 408, 217–220. 16. Jordan, S.W., and Cronan, J.E., Jr. (1997). A new metabolic link. The acyl carrier protein of lipid synthesis donates lipoic acid to the pyruvate dehydrogenase complex in Escherichia coli and mitochondria. J. Biol. Chem. 272, 17903–17906. 17. Wada, H., Shintani, D., and Ohlrogge, J. (1997). Why do mitochondria synthesize fatty acids? Evidence for involvement in lipoic acid production. Proc. Natl. Acad. Sci. USA 94, 1591–1596. 18. Harington, A., Herbert, C.J., Tung, B., Getz, G.S., and Slonimski, P.P. (1993). Identification of a new nuclear gene (CEM1) encoding a protein homologous to a beta-keto-acyl synthase which is essential for mitochondrial respiration in Saccharomyces cerevisiae. Mol. Microbiol. 9, 545–555. 19. Schneider, R., Brors, B., Burger, F., Camrath, S., and Weiss, H. (1997). Two genes of the putative mitochondrial fatty acid synthase in the genome of Saccharomyces cerevisiae. Curr. Genet. 32, 384–388. 20. Yamazoe, M., Shirahige, K., Rashid, M.B., Kaneko, Y., Nakayama, T., Ogasawara, N., and Yoshikawa, H. (1994). A protein which binds preferentially to single-stranded core sequence of autonomously replicating sequence is essential for respiratory function in mitochondrial of Saccharomyces cerevisiae. J. Biol. Chem. 269, 15244–15252. 21. Torkko, J.M., Koivuranta, K.T., Miinalainen, I.J., Yagi, A.I., Schmitz, W., Kastaniotis, A.J., Airenne, T.T., Gurvitz, A., and Hiltunen, K.J. (2001). Candida tropicalis Etr1p and Saccharomyces cerevisiae Ybr026p (Mrf1’p), 2-enoyl thioester reductases essential for mitochondrial respiratory competence. Mol. Cell. Biol. 21, 6243–6253.
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24. Joshi, A.K., Zhang, L., Rangan, V.S., and Smith, S. (2003). Cloning, expression, and characterization of a human 40 -phosphopantetheinyl transferase with broad substrate specificity. J. Biol. Chem. 278, 33142– 33149. 25. Zhang, L., Joshi, A.K., Hofmann, J., Schweizer, E., and Smith, S. (2005). Cloning, expression, and characterization of the human mitochondrial beta-ketoacyl synthase. Complementation of the yeast CEM1 knock-out strain. J. Biol. Chem. 280, 12422–12429. 26. Zhang, L., Joshi, A.K., and Smith, S. (2003). Cloning, expression, characterization, and interaction of two components of a human mitochondrial fatty acid synthase: malonyltransferase and acyl carrier protein. J. Biol. Chem. 278, 40067–40074. €, P.L., Sormunen, R.T., 27. Miinalainen, I.J., Chen, Z.-J., Torkko, J.M., Pirila Bergmann, U., Qin, Y.-M., and Hiltunen, J.K. (2003). Characterization of 2-enoyl thioester reductase from mammals: an ortholog of Ybr026p/ Mrf10 p of the yeast mitochondrial fatty acid synthesis type II. J. Biol. Chem. 278, 20154–20161. 28. Autio, K.J., Kastaniotis, A.J., Pospiech, H., Miinalainen, I.J., Schonauer, M.S., Dieckmann, C.L., and Hiltunen, J.K. (2008). An ancient genetic link between vertebrate mitochondrial fatty acid synthesis and RNA processing. FASEB J. 22, 569–578. 29. Chen, Z., Kastaniotis, A.J., Miinalainen, I.J., Rajaram, V., Wierenga, R.K., and Hiltunen, J.K. (2009). 17beta-hydroxysteroid dehydrogenase type 8 and carbonyl reductase type 4 assemble as a ketoacyl reductase of human mitochondrial FAS. FASEB J. 23, 3682–3691. 30. Stuible, H.P., Meier, S., Wagner, C., Hannappel, E., and Schweizer, E. (1998). A novel phosphopantetheine:protein transferase activating yeast mitochondrial acyl carrier protein. J. Biol. Chem. 273, 22334–22339. 31. Hoja, U., Wellein, C., Greiner, E., and Schweizer, E. (1998). Pleiotropic phenotype of acetyl-CoA-carboxylase-defective yeast cells–viability of a BPL1-amber mutation depending on its readthrough by normal tRNA(Gln)(CAG). Eur. J. Biochem. 254, 520–526. 32. Hoja, U., Marthol, S., Hofmann, J., Stegner, S., Schulz, R., Meier, S., Greiner, E., and Schweizer, E. (2004). HFA1 encoding an organellespecific acetyl-CoA carboxylase controls mitochondrial fatty acid synthesis in Saccharomyces cerevisiae. J. Biol. Chem. 279, 21779– 21786. 33. Schonauer, M.S., Kastaniotis, A.J., Hiltunen, J.K., and Dieckmann, C.L. (2008). Intersection of RNA processing and the type II fatty acid synthesis pathway in yeast mitochondria. Mol. Cell. Biol. 28, 6646–6657. 34. Monteuuis, G., Suomi, F., Keratar, J.M., Masud, A.J., and Kastaniotis, A.J. (2017). A conserved mammalian mitochondrial isoform of acetylCoA carboxylase ACC1 provides the malonyl-CoA essential for mitochondrial biogenesis in tandem with ACSF3. Biochem. J. 474, 3783– 3797. 35. Kursu, V.A., Pietikainen, L.P., Fontanesi, F., Aaltonen, M.J., Suomi, F., Raghavan Nair, R., Schonauer, M.S., Dieckmann, C.L., Barrientos, A., Hiltunen, J.K., et al. (2013). Defects in mitochondrial fatty acid synthesis result in failure of multiple aspects of mitochondrial biogenesis in Saccharomyces cerevisiae. Mol. Microbiol. 90, 824–840. 36. Brown, A., Rathore, S., Kimanius, D., Aibara, S., Bai, X.C., Rorbach, J., Amunts, A., and Ramakrishnan, V. (2017). Structures of the human mitochondrial ribosome in native states of assembly. Nat. Struct. Mol. Biol. 24, 866–869. 37. Hiltunen, J.K., Autio, K.J., Schonauer, M.S., Kursu, V.A., Dieckmann, C.L., and Kastaniotis, A.J. (2010). Mitochondrial fatty acid synthesis and respiration. Biochim. Biophys. Acta 1797, 1195–1202.
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44. Fiedorczuk, K., Letts, J.A., Degliesposti, G., Kaszuba, K., Skehel, M., and Sazanov, L.A. (2016). Atomic structure of the entire mammalian mitochondrial complex I. Nature 538, 406–410. 45. Zhu, J., Vinothkumar, K.R., and Hirst, J. (2016). Structure of mammalian respiratory complex I. Nature 536, 354–358. 46. Guo, R., Zong, S., Wu, M., Gu, J., and Yang, M. (2017). Architecture of human mitochondrial respiratory megacomplex I2III2IV2. Cell 170, 1247– 1257.e1212. 47. Wu, M., Gu, J., Guo, R., Huang, Y., and Yang, M. (2016). Structure of mammalian respiratory supercomplex I1III2IV1. Cell 167, 1598– 1609.e1510. 48. Van Vranken, J.G., Nowinski, S.M., Clowers, K.J., Jeong, M.Y., Ouyang, Y., Berg, J.A., Gygi, J.P., Gygi, S.P., Winge, D.R., and Rutter, J. (2018). ACP acylation is an acetyl-CoA-dependent modification required for electron transport chain assembly. Mol. Cell 71, 567–580.e564. 49. Carroll, J., Fearnley, I.M., Shannon, R.J., Hirst, J., and Walker, J.E. (2003). Analysis of the subunit composition of complex I from bovine heart mitochondria. Mol. Cell. Proteomics 2, 117–126. 50. Angerer, H., Schonborn, S., Gorka, J., Bahr, U., Karas, M., Wittig, I., Heidler, J., Hoffmann, J., Morgner, N., and Zickermann, V. (2017). Acyl modification and binding of mitochondrial ACP to multiprotein complexes. Biochim. Biophys. Acta 1864, 1913–1920.
Current Biology 28, R1212–R1219, October 22, 2018 R1219
Minireview Impact of Mitochondrial Fatty Acid Synthesis on Mitochondrial Biogenesis Sara M. Nowinski1, Jonathan G. Van Vranken1, Katja K. Dove1, and Jared Rutter1,2,* 1Department
of Biochemistry, University of Utah School of Medicine, Salt Lake City, UT 84132, USA Hughes Medical Institute, University of Utah School of Medicine, Salt Lake City, UT 84132, USA *Correspondence: [email protected] https://doi.org/10.1016/j.cub.2018.08.022 2Howard
Biology students today are taught that mitochondria are ‘the powerhouse of the cell’. This gross over-simplification of their cellular role has arguably led to a paucity of knowledge concerning the many other tasks carried out by this multifunctional organelle. Mitochondrial fatty acid synthesis (mtFAS) is one such underappreciated pathway that is crucial for mitochondrial function, although even mitochondrial experts are often surprised to learn of its existence. For many years, the only function of mtFAS was thought to be the production of lipoic acid, an important co-factor for several mitochondrial enzymes. However, recent advances have revealed a far wider role for mtFAS in mitochondrial physiology. The discovery of human patients with mutations in mtFAS enzymes has brought renewed interest in understanding the full significance of this novel mode of mitochondrial metabolic regulation. We now appreciate that mtFAS is a nutrient-sensitive pathway that provides an elegant mechanism whereby acetyl-CoA regulates its own consumption via coordination of lipoic acid synthesis and tricarboxylic acid (TCA) cycle activity, iron–sulfur (FeS) cluster biogenesis, assembly of oxidative phosphorylation complexes, and mitochondrial translation. In this minireview, we describe and build upon the important discoveries that led to our current understanding of this elegant mechanism of coordination of nutrient status and metabolism. Introduction Fatty acid synthesis (FAS) is an essential process in organisms from bacteria to mammals. FAS has a number of cellular roles, including the production of triglycerides as lipid storage units, the generation of phospholipids to build cell membranes, and many more. However, it is a little-known fact that eukaryotes actually harbor two distinct FAS systems — one in the cytoplasm, and one in mitochondria. Although mitochondrial fatty acid synthesis (mtFAS) was discovered decades ago, the importance of this pathway has been underestimated compared with its more famous cytoplasmic cousin FASN. The recent finding that some patients with neurogenerative disease harbor mutations in a mtFAS enzyme has generated new interest in understanding the cellular role(s) played by this ‘old’ pathway [1]. Unlike FASN, mtFAS does not appear to contribute significantly to cellular triglyceride stores or phospholipids, but is instead responsible for the generation of lipoic acid, an important lipid cofactor for multiple mitochondrial dehydrogenases. Recent studies have now implicated mtFAS in a range of mitochondrial processes beyond its classical role in lipoic acid production, from mitochondrial translation to the support and coordination of mitochondrial oxidative phosphorylation. Herein, we follow the evolution of our understanding of the mtFAS pathway and the processes it governs. Discovering mtFAS FAS is a complex process whereby saturated fatty acids are built stepwise in two carbon units, the addition of which is mediated by several enzymatic activities. Most bacteria utilize a dissociated pathway in which separate enzymes catalyze each step in the elongation of fatty acyl chains on a soluble acyl carrier protein
(ACP) scaffold. In contrast, eukaryotes evolved more consolidated systems that encode all of the necessary enzymatic activities on one (mammals) or two (yeast) peptides (Figure 1 and Table 1). These systems are termed type II and type I fatty acid synthesis, respectively, and all organisms were historically thought to exclusively utilize one pathway or the other, with few exceptions. However, reports claiming to observe fatty acid synthesis in mammalian mitochondria (mtFAS) were published in the 1960s to much skepticism. Real progress towards understanding this pathway did not truly come until the late 1980s, when the eukaryotic mitochondrial ACP was discovered in Neurospora crassa by Stuart Brody and Steve Mikolajczyk while they were looking for pantetheine-modified proteins [2]. A highly conserved serine residue on ACP is modified with a 4’-phosphopantetheine (4’-PP) cofactor, which supplies the terminal thiol group upon which fatty acids are built. This seminal study identified the primary acyl chain on ACP as the 14-carbon lipid 3-hydroxymyristate; however, the source of the acyl chain was at first unknown. Was it made by cytoplasmic FASN or by a novel synthesis pathway in the mitochondria? De novo mtFAS in N. crassa was described two years later, with a series of in vitro experiments that led the researchers to conclude that myristic acid (14:0) was the ‘product’ of mtFAS, while in vivo experiments again yielded ACP with labelled hydroxymyristic acid (14:0-OH) [3]. Initially, the primary hypothesis regarding the significance of this pathway was that mtFAS likely contributed to the synthesis of mitochondrial phospholipids. Soon after the initial discovery of ACP and mtFAS, efforts by Hanns Weiss’s lab to characterize complex I of the mitochondrial electron transport chain (ETC) in N. crassa also discovered ACP as a stable subunit of this complex [4]. Around the same time,
R1212 Current Biology 28, R1212–R1219, October 22, 2018 ª 2018 Elsevier Ltd.
Current Biology
Minireview FAS I (yeast) FAS II (mtFAS)
O
ACT
ER
DH
AT
ACP
KR
ACT?
ER
DH
AT
ACP
KR
ACC
OH
CH3 C CoA
CH3 (CH2)N CH CH2 C S
Malonyl-CoA
Initiation
AT
H2O
O
O
HO C CH2 C S
ACP
KR
ACT/KS
CO2 O
O
NADPH
CoA CoA
PPT
3-hydroxyacyl-ACP
NADP
+
acetyl-CoA
PPT
KS
O
O
HO C CH2 C CoA
KS
O
O
CH3 (CH2)N C CH2 C S
ACP
Malonyl-ACP
DH
CH3 (CH2)N CH CH C S
ACP
ACP
trans -2-enoyl-ACP
3-ketoacyl-ACP
ER HS
HS
ACP
O
ACP CO2
Holo-ACP
Holo-ACP
Elongation
KS
Apo-ACP O C
S
CH2 CH2
O
N
C
CH2 CH2
CH3 (CH2)N C S
ACP
Octanoyl-ACP H
CH3
C
C
C
O
OH CH3
H
H
O
Acyl chain
NADP+
O CH3 (CH2)6 C S
Lipoic acid and lipoylated proteins
ACP
CH3 (CH2)N
ACP
Acyl-ACP
4’-phosphopantetheine
PPT
NADPH
CH3 (CH2)N C S
N
ACP
Acyl-ACP
OCH2 O P O
4’-phosphopantetheine
ACP
Substrate
O
Ser
ACP
Apo-ACP No 4’-PP
Products
HS
ACP
Holo-ACP 4’-PP Current Biology
Figure 1. Fatty acid synthesis is catalyzed by a series of conserved enzymatic activities in the eukaryotic cytoplasm and mitochondria. (Top) A schematic representing the different subunits of yeast FAS I and FAS II. In mammals, components of FAS I are contained on one polypeptide, whereas yeast FAS I comprises two polypeptides (black boxes). On the contrary, members of FAS II are distinct entities in bacteria, yeast and mammals. (Middle) Mitochondrial fatty acid synthesis pathway. Enzymes corresponding to each of the domain/protein abbreviations can be found in Table 1. (Bottom left) Chemical structure of the ACP prosthetic group 4’-phosphopantetheine covalently bound to a conserved serine is shown. (Bottom right) Key for the FAS pathway shown in the middle section.
ACP was found in complex I from bovine heart, marking its first description in mammalian mitochondria [5]. Both studies suggested that the majority of ACP in complex I was acylated, while unbound ACP was mostly de-acylated, and it was therefore postulated that complex I was required for mtFAS. Many other studies describing mitochondrially localized ACP in various organisms followed [6–9], culminating with its eventual discovery in humans [10]. Despite many advances in our understanding of this system, several pertinent questions have remained unanswered until today. What is the acyl chain modification on ACP and/or the major product of mtFAS? Why is ACP associated with complex I and what are the implications of this interaction? What is the purpose of mtFAS, and why has this pathway been evolutionarily conserved when it is known that fatty acids can be imported into mitochondria from the endoplasmic reticulum or cytosol? Recent publications discussed below have made significant advances towards answering these questions and have identified important trajectories for future research.
Defining the Product(s) of mtFAS In the decade following the initial discovery of mtFAS and ACP, a major research goal was to define the ultimate product(s) of mtFAS. Using an N. crassa strain with defective cytoplasmic FAS, the Weiss lab again found 3-hydroxymyristate (14:0) and 3-hydroxylaurate (12:0) as the major products [11]. While these early studies reproducibly identified a 14-carbon lipid as the major product of mtFAS, there was mixed opinion over the cellular role that this lipid was likely to play. While earlier studies had proposed that mtFAS was expected to contribute generally to mitochondrial membrane phospholipids, Weiss and colleagues disagreed, citing the existing knowledge that most mitochondrial phospholipids are imported from the endoplasmic reticulum [12]. They proposed a narrower hypothesis: that mtFAS supplied acyl chains specifically for cardiolipin biosynthesis within the mitochondria. The identification of genes encoding ACP (acp-1, ACP1) in a number of organisms enabled knockout studies aimed at answering this question [13]. The initial Current Biology 28, R1212–R1219, October 22, 2018 R1213
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Minireview Table 1. Fatty acid synthesis is a conserved pathway in bacteria, yeast, and mammals. FAS II gene names Domain/protein
Function
E. coli
S. cerevisiae
H. sapiens
ACC
Acetyl-CoA carboxylase
accA, accB/fabE, accC, accD
HFA1?
ACC1? (ACSF3?)
ACP
Acyl carrier protein
accP
ACP1
NDUFAB1
ACT
Acetyltransferase
fabH
?
?
AT
Acyltransferase
fabD
MCT1
MCAT
DH
Dehydratase
fabA, fabZ
HTD2
HTD2
ER
Enoyl reductase
fabI, fabK, fabL, fabV
ETR1
MECR
KR
Ketoreductase
fabG
OAR1
HSD17B8, CBR4
KS
Ketosynthase
fabB, fabF
CEM1
OXSM
PPT
Phosphopantetheinyl-transferase
acpS
PPT2
AASDHPPT?
Enzyme activities and the corresponding genes that encode this activity in E. coli, S. cerevisiae, and H. sapiens are listed. ? indicates areas of uncertainty and/or ongoing research. ACSF3 is listed in parentheses next to ACC1 because it has been implicated in the generation of malonyl-CoA in mammalian mitochondria but is a malonyl-CoA synthetase rather than an acetyl-CoA carboxylase.
knockout studies were performed simultaneously in N. crassa and Saccharomyces cerevisiae [14]. Knockout of acp-1 in N. crassa ablated the peripheral module of complex I and resulted in improper assembly of the membrane module and a lack of incorporation of iron–sulfur (FeS) clusters into complex I. However, respiration was otherwise unaffected, and no differences in mitochondrial phospholipids were observed. In contrast, knockout of ACP1 in S. cerevisiae resulted in a much more severe ‘‘pleiotropic respiratory-deficient phenotype’’. In these mutants, mitochondrial phospholipids did show some changes, though none that are not observed in other respiratory-deficient mutants (rho– or rho0 ‘petite’ mutants). The severity of the phenotype and similarity to other rho0 mutants led the researchers to conclude that these cells were rho– or rho0, and that the observed loss of ETC complexes and changes in mitochondrial lipids were likely indirect. Because mitochondrial phospholipids were largely unaffected by knockout of ACP in N. crassa, these first genetic studies seeded the idea that perhaps ACP was involved in the delivery of a specific product essential for the proper formation of the membrane module of complex I. However, this way of thinking changed a few years later in 1997, when it was discovered that ACP was required for the synthesis of lipoic acid, an important cofactor for several mitochondrial dehydrogenases, including pyruvate dehydrogenase (PDH), a-ketoglutarate dehydrogenase (a-KGDH), and the glycine cleavage system [15–17]. Shedding some doubt on the phenotypic findings of earlier ACP-knockout studies, the Schweizer lab reported that ACP1 loss was often lethal in S. cerevisiae and proposed that surviving mutants had activated some compensatory mechanism [15]. Since lipoic acid is only an eight-carbon lipid, and previous studies had all identified 3-hydroxymyristic acid as the major acyl chain found covalently linked to ACP, it remained likely that longer acyl products of mtFAS played some other cellular role [15]. However, with these findings the focus of the mtFAS field completely shifted toward the study of lipoic acid production. The possible role(s) of longer fatty acids synthesized on ACP and of ACP in ETC assembly and function were largely forgotten and would not be revisited for nearly two decades. R1214 Current Biology 28, R1212–R1219, October 22, 2018
Identifying the mtFAS Enzymes The discovery of ACP and de novo mitochondrial fatty acid biosynthesis started a race to identify the mtFAS enzymes and the genes that encode them. Prokaryotic (type II) FAS is catalyzed by at least five enzymes. For each two-carbon increase in acyl chain length, these enzymes catalyze the addition of malonyl-CoA to ACP, the condensation of malonyl-ACP with a growing acyl chain forming a keto-acyl-ACP product, and the subsequent reduction of that keto group to a methylene group, completing the elongation cycle (Figure 1). The first eukaryotic mtFAS gene to be discovered in yeast was CEM1, which was identified as the enzyme that performs the condensation reaction in 1993 by virtue of its sequence homology to the bacterial b-keto-acyl synthase, at a time before the identification of lipoic acid as a product of mtFAS [18]. Null cem1 mutants are respiratory deficient but show no major differences in mitochondrial lipids and fatty acids, an observation that is consistent with findings from non-lethal ACP1 knockouts. The genes encoding two more yeast mtFAS enzymes, MCT1 and OAR1, were also identified via homology to their respective bacterial acyltransferase and ketoreductase counterparts, and mutation of these genes led to similar phenotypes as those of acp1 and cem1 mutants [19]. The last two S. cerevisiae genes proved to be more elusive. ETR1 was originally identified as MRF1, a protein that was purified from nuclear extracts and shown to bind to autonomously replicating sequences of nuclear DNA [20]. It was eventually identified as an enoyl thioester reductase by homology to an enzyme from Candida tropicalis and by complementation of mutants by a mitochondria-targeted version of the Escherichia coli FabI enzyme. Re-investigation of its localization revealed that it was found predominantly in mitochondria and that mitochondrial localization was essential for rescue of the petite phenotype in etr1 mutants [21]. The dehydratase encoded by HTD2, on the other hand, was discovered via a screen that used a mitochondria-targeted form of the E. coli FabA gene (Table 1) [22]. htd2 mutant phenotypes were identical to those observed for other mtFAS mutants, indicating that the resultant defects were all likely the result of a failure to acylate ACP. In support of this idea, re-localization of a peroxisomal fatty acyl-CoA synthetase to mitochondria rescued cem1 mutants, presumably by
Current Biology
Minireview re-acylating ACP [23]. Identification of most of the corresponding mammalian genes came about a decade later and added a few mechanistic insights, showing that these enzymes can accommodate chain lengths out to C14–C16 in vitro [24–29]. While the major catalytic enzymes of the mtFAS pathway are now defined, a number of accessory enzymes important for mtFAS are less well studied. One such enzyme is Ppt2, the mitochondrial phosphopantetheinyl transferase that catalyzes the addition of the 4’-PP moiety from coenzyme A to the invariant serine residue on ACP [30]. Interestingly, mammals appear to have only a cytoplasmic version of phosphopantetheinyl transferase [24], raising the question of whether the 4-phosphopantetheine is added in the cytosol and translocated into the mitochondria, or whether there is an undiscovered mitochondrial phosphopantetheinyl transferase. Similarly, mitochondrial malonyl-CoA is a required substrate of mtFAS, but the identity of a mitochondrial acetyl-CoA carboxylase (ACC) enzyme that would synthesize malonyl-CoA from acetyl-CoA is still uncertain. Originally, researchers proposed that the cytosolic ACC enzyme Acc1 might fulfil this function, but no mechanism for malonylCoA import has been described. Despite this, mutants with defects in Acc1 activity are respiratory deficient and reportedly have a 50% reduction in lipoic acid levels [31]. The same lab later identified Hfa1 as the mitochondrial ACC; however, hfa1D strains are respiration competent under normal growth conditions, indicating that mtFAS is still functional and that there is at least an additional source of mitochondrial malonyl-CoA in these cells ([32,33] and our unpublished data, J.G. Van Vranken). In mammals, there is some evidence that ACC1 and ACSF3 (a mammalian malonyl-CoA synthetase) might work in a coordinated fashion [34]. Regardless, the source of mitochondrial malonyl-CoA remains up for debate. Beginning to Put It Together Although early studies largely agreed on the phenotypes of mtFAS knockouts, it was unclear what mechanisms underlied their pleiotropic respiratory-deficient phenotype, and whether these effects were a result of the loss of lipoic acid, or some other function of mtFAS. Several possibilities have been proposed. While some studies claimed that ACP-deficient cells were rho0, these cells were genetically rescuable, indicating that they retained at least some mitochondrial DNA [14]. Subsequent reports clarified that mtFAS mutants in fact had no defects in mitochondrial DNA replication under normal growth conditions [18,20,23,33]. Another possibility was defective mitochondrial translation. Although early studies identifying mtFAS enzymes reported no defects in mitochondrial translation, our lab and others have measured translational deficiencies in mtFAS mutants ([33,35] and our unpublished data, Sara M. Nowinski and Olga Zurita Rendon). One study identified a requirement for mtFAS in the processing of the 5’ ends of mitochondrial tRNAs by RNase P, with mtFAS mutants having profound defects in the synthesis of mitochondrial tRNAs [33]. Moreover, the authors show that lipoic acid synthesis mutants do not exhibit the same defects as mtFAS mutants, implying that the activation of RNase P may be one role for the longer acyl chains synthesized on ACP. However, the mechanism(s) whereby mtFAS is required for RNase P function remain unknown. Yeast RNase P is composed of one nuclear-encoded
peptide (Rpm2) and one mitochondria-encoded RNA (RPM1), which is itself processed by RNase P to its mature form, while human mitochondrial RNase P is a heterotrimer of three protein subunits, none of which resembles Rpm2. Thus, it is unlikely that the mechanisms underlying RNase P activity are conserved in mammalian systems, although other mechanisms to connect mtFAS and translation may have evolved. The mammalian HTD2 and RPP14 (a mammalian nuclear RNase P subunit) are encoded by the same mRNA transcript [28]. Furthermore, a recent structure of a mitochondrial ribosome assembly intermediate identified ACP bound directly to the mitochondrial ribosome [36]; however, the functional significance of these observations remains to be seen. To this point, previous studies had identified two major routes whereby mtFAS regulates mitochondrial metabolism: via lipoic acid production and mitochondrial translation. This creates two positive feedback loops in mitochondria. In the first, PDH converts pyruvate to acetyl-CoA using its lipoylated domain; some of this acetyl-CoA enters the tricarboxylic acid (TCA) cycle, while some becomes the substrate for the mtFAS pathway, ultimately producing more lipoic acid. In the second, RNase P processes its own RPM1 precursor RNA to form more active RNase P and enable mitochondrial translation. Both of these processes are tied to the requirement for a mtFAS product (lipoic acid in the case of PDH, unknown in the case of RNase P) for their activity. Hiltunen and Kastaniotis therefore proposed the insightful speculation in the absence of confirmatory data that these pathways might regulate mitochondrial function in response to nutrient status [37]. A subsequent publication from the same lab focused on etr1D cells and showed that these mutants have low respiration, low activities of ETC complexes, and low membrane potential [35]. They attributed this phenotype to a reduction in steady-state levels of ETC complexes II, III, and IV subunits but not complex V. However, their proposed model hinged on observed effects on mitochondrial RNA splicing, which cannot explain the effects of mtFAS loss on complex II assembly and function because it is encoded entirely in the nucleus, indicating that some other mechanism must still be at play. Finally, this publication also postulated that because acetyl-CoA is the substrate of the TCA cycle and mtFAS, this pathway is a mechanism by which mitochondria sense cellular metabolic status and control mitochondrial biogenesis [35]. Although studies had speculated that ETC assembly defects might underlie the profound mitochondrial defects observed in mtFAS mutants, none of these phenotypes could explain the observation first made by Brody et al. that ACP1 is an essential gene in many genetic backgrounds [15,33]. Therefore, our lab set out to identify the essential function of ACP. Proteomics studies had identified physical interactions between ACP and Isd11, a protein that is part of the mitochondrial FeS cluster biogenesis machinery [38–41]. We found that ACP is an essential and stable subunit of the FeS cluster biosynthetic complex, where it physically interacts with Isd11 and enables FeS cluster biosynthesis in an acylation-dependent fashion [41]. Subsequently, two structures confirmed this interaction, and showed an acyl modification on ACP that is embedded in the three-helix bundle formed by Isd11 [42,43] (Figure 2). These studies reported density in the crystal structure that could correspond to at least a 12-carbon acyl chain, but Isd11 could potentially accommodate longer Current Biology 28, R1212–R1219, October 22, 2018 R1215
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Minireview ACP–ISD11 iron sulfur cluster biogenesis
ACP–NDUFB9 complex I
ACP–NDUFA6 complex I
ACP is acylated
ACP–L0R8F8 (novel LYR) ribosome assembly intermediate
ACP is not acylated Current Biology
Figure 2. ACP–LYR protein interactions suggest an important role for ACP acylation. Cartoon representations of ACP bound to LYR proteins in different contexts. ACP is colored blue in all structures. (Left) The interaction between ACP and Isd11 (yellow, PDB 5USR), a LYR protein involved in iron–sulfur cluster biogenesis, intimately involves an acyl chain (grey spheres) covalently bound to ACP through 4’-phosphopantetheine (4’-PP, grey spheres). ACP–LYR interactions involved in electron transport chain (ETC) assembly also show acylation of ACP, as depicted by interactions with two LYR proteins in ETC complex I, NDUFB9 (green, PDB 5XTH) and NDUFA6 (pink, PDB 5XTH). (Right) A recent structure of the mitochondrial ribosome revealed a novel LYR protein L0R8F8 (light blue, PDB 5OOL) to which ACP is bound, although in this structure ACP is not acylated.
chains, and purification of the complex followed by fatty acid methyl ester gas chromatography mass spectrometry (FAME GC-MS) analysis found primarily 16-carbon acyl chains. Acylated ACP Directly Coordinates Mitochondrial Metabolism with Nutrient Status Concurrent with the identification of the ACP–Isd11 interaction and corresponding structures, several proteomic and structural studies identified interactions between ACP and a family of (mostly) late-stage ETC assembly factors known as the LYR proteins, of which Isd11 is a member [36,38–47]. The LYR proteins are so-called because each member contains a degenerate Leu–Tyr–Arg (LYR) motif, which is required for interaction of these proteins with ACP [48]. In addition to Isd11, the yeast LYR family includes Sdh6, Sdh7, Mzm1, and Fmc1, which facilitate the last steps in the assembly of ETC complexes II (Sdh6/ Sdh7), III (Mzm1), and V (Fmc1) (Figure 3; reviewed in [38]). Mammals and fungi that have ETC complex I also express two additional LYR protein–ACP dimers that are stable subunits of ETC complex I [38]. Additionally, mammalian genomes encode at least five additional LYR proteins that are largely uncharacterized. In agreement with the very early studies identifying ACP in complex I, recent studies show that ACP is nearly uniformly acylated when bound to LYR proteins, and moreover that these acyl chains appear to be longer (C12–C14) than those used as precursors for lipoic acid synthesis (C8), although the exact identity of the acyl chain is still debated [49,50]. Beyond providing a potential mechanistic link to ETC assembly, the structural studies that identified ACP–LYR interactions hinted at an intimate role for ACP acylation in these complexes, because like Isd11, the acyl chain threads into the middle of a three-helix bundle in each LYR (Figure 2) [42–45]. Interestingly, a recent structure of ACP bound to a LYR protein as part of a ribosome assembly intermediate reveals that ACP is not acylated, implying that the acyl chain is not favored in all ACP–LYR interactions (Figure 2) [36,48]. The obvious question that followed these obR1216 Current Biology 28, R1212–R1219, October 22, 2018
servations was whether acylation was important for the function of these interactions. A recent publication from our group describes the mechanism whereby mtFAS, via ACP acylation and interaction with LYR proteins, controls the assembly of mitochondrial ETC complexes in S. cerevisiae [48] (Figure 3). LYR proteins preferentially interact with acylated ACP (Figure 2), and this interaction is crucial for the ability of the LYR proteins to function in ETC complex assembly. Moreover, we showed that this process is dependent on mitochondrial acetyl-CoA availability. mtFAS and ACP acylation therefore indeed can be thought of as a ‘sensor’ for mitochondrial acetyl-CoA levels as Hiltunen and Kastaniotis proposed [35,37]. This system theoretically provides a mechanism to integrate most major mitochondrial processes with nutrient levels, as ACP acylation affects mitochondrial translation via RNase P, FeS cluster biogenesis via Isd11, ETC assembly via the other LYRs, and finally, TCA cycle and 1-carbon metabolism via lipoic acid production. Mitochondrial acetyl-CoA is, of course, not just the substrate for mtFAS, but is much better known for its role as a major substrate for the TCA cycle, where it is oxidized to generate the reduced forms of nicotinamide adenine dinucleotide (NADH) and flavin adenine dinucleotide (FADH2), which feed reducing equivalents to the ETC, ultimately fueling the production of mitochondrial ATP. As such, acetyl-CoA elegantly regulates its own metabolism through mtFAS, tying mitochondrial acetyl-CoA availability to the activity of TCA cycle enzymes and ETC oxidative capacity (Figure 3). Conclusions The story of mtFAS has evolved slowly over the course of several decades, leading to our current appreciation of its multifaceted role in mitochondrial metabolism. This pathway, largely ignored by the mitochondrial research community, has now returned to the limelight of mitochondrial metabolism. From the moment of the discovery of mitochondrial ACP and de novo mtFAS, the purpose and cellular function of this pathway has
Current Biology
Minireview Figure 3. The mtFAS pathway is a nutrientsensitive coordinator of mitochondrial oxidative phosphorylation.
ACP Acetyl-CoA
mtFAS AcylACP
OctanoylACP
?
Lipoic acid
TCA cycle
Mitochondrial translation
LYR network
NADH/FADH 2
AcylACP
LYR
FeS
II
III
IV
I
V
OXPHOS IMS
(Top) Schematic overview of the integrated role of mtFAS in the coordination of mitochondrial nutrient supply and metabolism. Acetyl-CoA feeds the TCA cycle and mtFAS pathway, where it fuels the generation of octanoyl-ACP as well as longer acyl chains (acyl-ACP). Octanoyl-ACP is the precursor for the synthesis of lipoic acid, which is an important cofactor for TCA cycle and other mitochondrial enzymes. Acyl-ACP supports oxidative phosphorylation via two mechanisms: an undefined role in mitochondrial translation, as well as its interaction with the LYR protein network. (Bottom) In complex with Acyl-ACP, LYRs support the assembly of ETC complexes and the iron–sulfur (FeS) cluster biogenesis machinery. Specifically, Sdh6 and Sdh7 (SDHAF1 and SDHAF3) facilitate the assembly of complex II, Mzm1 (LYRM7) assembles complex III, Fmc1 supports assembly of complex V, and Isd11 (LYRM4) is an essential subunit of an FeS complex biosynthetic complex. In mammals, two additional LYR proteins (NDUFA6 and NDUFB9) in complex with acylACP are stable subunits of complex I.
C Q
I
II
III
IV
NDUFB9
Matrix
NDUFA6
AcylACP Sdh6
AcylACP AcylACP FeS Isd11 complex
AcylACP
Mzm1 Sdh7 AcylACP
AcylACP
been under debate. Although early publications speculated about a function for mtFAS in mitochondrial phospholipid and/or cardiolipin production, it appears that fatty acids produced by this pathway play much more specific roles. The discoveries of Brody et al. [15] and Wada et al. [17] defined the role of ACP in lipoic acid production, arguably one of its most important cellular functions. However, it has been clear from the beginning that longer acyl chains are synthesized on ACP. In the last five years, research advances have allowed us to finally understand the cellular role of these other mtFAS products in the regulation of mitochondrial translation, FeS cluster biogenesis, and ETC assembly. Through its specific interaction with the LYR proteins, acylation of ACP provides the cell with a means to coordinate acetyl-CoA concentration with the ability to metabolize acetyl-CoA. While this elegant regulatory mechanism and the purpose of longer acyl chains on ACP have now been described, many interesting questions remain. What is the utility of maintaining such a complex system for monitoring acetyl-CoA and coordinating mitochondrial function throughout evolution, especially given that targeting a single fatty acyl-CoA synthetase (Faa2) to the mitochondrial matrix can circumvent this process [23,48]? While mitochondrial acetyl-CoA is clearly required for mtFAS and ACP
acylation, the link between acetyl-CoA and malonyl-CoA in the mitochondria is not well established. To what extent is the acyl-ACP–LYR regulatory network conserved in higher eukaryotes? Given Fmc1 that two mammalian LYR proteins are Acylstable subunits of ETC complex I, ACP Fmc1 insights into the functional significance and regulation of these interactions may prove particularly interesting. And finally, Current Biology beyond the level of substrate regulation, how is this pathway controlled? It will be exciting to follow the discoveries as these and other questions guide future studies aimed at understanding the importance of this elegant mechanism of metabolic regulatory control.
V
ACKNOWLEDGEMENTS We would like to acknowledge the many groups who contributed to the studies defining the functions of the LYR proteins reviewed in [38], but could not be cited due to space constraints. Related work in the lab of the authors was partially supported by grants to J.R. from the NIH (GM115174 and GM115129), the Nora Eccles Treadwell Foundation, and the Howard Hughes Medical Institute. S.M.N. was also supported by postdoctoral fellowships from UMDF and ACS. REFERENCES 1. Heimer, G., Keratar, J.M., Riley, L.G., Balasubramaniam, S., Eyal, E., Pietikainen, L.P., Hiltunen, J.K., Marek-Yagel, D., Hamada, J., Gregory, A., et al. (2016). MECR mutations cause childhood-onset dystonia and optic atrophy, a mitochondrial fatty acid synthesis disorder. Am. J. Hum. Genet. 99, 1229–1244. 2. Brody, S., and Mikolajczyk, S. (1988). Neurospora mitochondria contain an acyl-carrier protein. Eur. J. Biochem. 173, 353–359. 3. Mikolajczyk, S., and Brody, S. (1990). De novo fatty acid synthesis mediated by acyl-carrier protein in Neurospora crassa mitochondria. Eur. J. Biochem. 187, 431–437.
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