Fatty acid activation in thermogenic adipose tissue

Fatty acid activation in thermogenic adipose tissue

BBA - Molecular and Cell Biology of Lipids 1864 (2019) 79–90 Contents lists available at ScienceDirect BBA - Molecular and Cell Biology of Lipids jo...

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BBA - Molecular and Cell Biology of Lipids 1864 (2019) 79–90

Contents lists available at ScienceDirect

BBA - Molecular and Cell Biology of Lipids journal homepage: www.elsevier.com/locate/bbalip

Review

Fatty acid activation in thermogenic adipose tissue☆,☆☆ ⁎

T

Sandra Steensels, Baran A. Ersoy

Department of Medicine, Division of Gastroenterology and Hepatology, Weill Cornell Medical College, New York, NY, USA

A R T I C LE I N FO

A B S T R A C T

Keywords: Thermogenesis Acyl-CoA synthetase Acyl-CoA thioesterase Obesity

Channeling carbohydrates and fatty acids to thermogenic tissues, including brown and beige adipocytes, have garnered interest as an approach for the management of obesity-related metabolic disorders. Mitochondrial fatty acid oxidation (β-oxidation) is crucial for the maintenance of thermogenesis. Upon cellular fatty acid uptake or following lipolysis from triglycerides (TG), fatty acids are esterified to coenzyme A (CoA) to form active acyl-CoA molecules. This enzymatic reaction is essential for their utilization in β-oxidation and thermogenesis. The activation and deactivation of fatty acids are regulated by two sets of enzymes called acyl-CoA synthetases (ACS) and acyl-CoA thioesterases (ACOT), respectively. The expression levels of ACS and ACOT family members in thermogenic tissues will determine the substrate availability for β-oxidation, and consequently the thermogenic capacity. Although the role of the majority of ACS and ACOT family members in thermogenesis remains unclear, recent proceedings link the enzymatic activities of ACS and ACOT family members to metabolic disorders and thermogenesis. Elucidating the contributions of specific ACS and ACOT family members to trafficking of fatty acids towards thermogenesis may reveal novel targets for modulating thermogenic capacity and treating metabolic disorders.

1. Introduction Fatty acids are major building blocks of life and are essential for a wide array of cellular functions in mammals. The cellular fatty acid supply can be maintained via a number of mechanisms [1]; 1) dietary triglycerides (TG) can be hydrolyzed in the lumen of the intestine followed by uptake by enterocytes. These lipids can be re-synthesized in the endoplasmic reticulum (ER) and secreted with chylomicrons and high density lipoproteins that can be absorbed by the peripheral cells, 2) adipocytes can release fatty acids into the circulation after lipolysis of TG that are stored in lipid droplets (LD) or 3) cellular fatty acids can be supplemented within the cell through lipolysis or by de novo lipogenesis from acetate. These cellular fatty acids can be processed further by elongation, desaturation and oxidation or converted into acylcarnitines, esterified to form glycerolipids, retinol esters and cholesterol

esters, acylated to proteins or incorporated into ceramide to form sphingolipids. The regulation of the cellular fatty acid landscape is highly dynamic and depends on the metabolic demands of the cell and organism, which can be affected by disease, fasting and feeding, exercise and environmental temperature. These factors determine pathways of fatty acid utilization, including mitochondrial fatty acid oxidation (β-oxidation) as an energy source or TG storage in LD. Thermogenesis is a process that produces heat and helps to maintain body temperature. Every organ and metabolic process contributes to heat production since nearly every enzymatic reaction is thermogenic. Heat is produced during digestion and absorption of food and during exercise. Heat is also produced as energy is dissipated in response to environmental changes such as diet and cold temperatures [2]. These latter forms are referred to as adaptive thermogenesis and can be divided into three subtypes; 1) cold exposure induces shivering

Abbreviations: −/−, knockout; β-oxidation, fatty acid oxidation; β3-AR, β3-adrenergic receptor; ACOT, acyl-CoA thioesterase; ACS, acyl-CoA synthetase; ACSL, long-chain acyl-CoA synthetase; ACSM, medium-chain acyl-CoA synthetase; ACSS, short-chain acyl-CoA synthetase; ACSVL, very long-chain acyl Co-A synthetase; ATGL, desnutrin/adipose triglyceride lipase; cAMP, cyclic adenosinemonophosphate; BAT, brown adipose tissue; BFIT, brown-fat-inducible thioesterase; CD36, cluster of differentiation 36; CoA, coenzyme A; CPT, carnitine palmitoyl transferase; CTMP, C-terminal modulator protein; DGAT, diacylglycerol O-acyltransferase; DG, diacylglycerol; ER, endoplasmic reticulum; ETC, electron transport chain; FA, fatty acid; FATP, fatty acid transport protein; GPAT, glycerol-3-phosphate acyltransferase; GLUT1, glucose transporter 1; GLUT4, glucose transporter 4; HCM, hypertrophic cardiomyopathy; HSL, hormone-sensitive lipase; LAL, lysosomal acid lipase; LD, lipid droplet; LPL, lipoprotein lipase; LVWT, left ventricle wall thickness; MCFA, medium-chain fatty acid; MG, monoacylglycerol; MGL, monoacylglycerol lipase; mTOR, mammalian target of rapamycin; NEFA, non-esterified fatty acid; PET, positron-emission tomography; PPAR, peroxisomal proliferator activated receptor; PI3K, phosphoinositide 3-kinase; PKA, protein kinase A; ROS, reactive oxygen species; Sc, subcutaneous; START, steroid acute regulatory-related transfer; TCA, tricarboxylic acid; TG, triglyceride; THEM, Thiosterase superfamily member; TRL, triglyceride-rich lipoprotein; UCP1, uncoupled protein 1; WAT, white adipose tissue ☆ This article is part of a Special Issue entitled Brown and Beige Fat: From Molecules to Physiology Guest Editor: Paul Cohen. ☆☆ The authors have declared that no conflict of interest exists. ⁎ Corresponding author at: Weill Cornell Medical College, 413 East 69th St, Belfer Research Building, Room BB-628, New York, NY 10021, USA. E-mail address: [email protected] (B.A. Ersoy). https://doi.org/10.1016/j.bbalip.2018.05.008 Received 4 November 2017; Received in revised form 10 March 2018; Accepted 17 May 2018

Available online 21 May 2018 1388-1981/ © 2018 Elsevier B.V. All rights reserved.

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Fig. 1. Key metabolic and signaling pathways in brown and beige adipocytes. This model is largely derived from [14,53,105,106,109–112,140–144]. Subcellular compartments are not drawn to proportion in order to emphasize on the lipid droplet and the mitochondria. Cold exposure induces noradrenalin release resulting in the activation of β3-adrenergic receptors (β3-AR) in brown and beige adipocytes. This activation results in increased intracellular cAMP levels, which stimulate GLUT1-mediated glucose uptake via the mTOR signaling pathway and lipolysis via activation of the protein kinase A (PKA) signaling pathway. Fatty acids (FA) in the brown and beige adipocytes can originate from three different pathways; cluster of differentiation 36 (CD36) can mediate uptake of lipoprotein in addition to channeling long chain fatty acids or FA can originate from BAT lipolysis. Lipolysis is a stepwize process and involves triacylglycerol (TAG) hydrolysis by desnutrin/adipose triglyceride lipase (ATGL) to form diacylglycerol (DAG) which is hydrolyzed by hormone-sensitive lipase (HSL) to monoacylglycerol (MG) and subsequently glycerol, with a fatty acid (FA) being released at every step. These fatty acids undergo β-oxidation in the mitochondria which will result in UCP-1 mediated heat production. The cold-depleted energy stores of the brown adipocytes are replenished by de novo lipogenesis and glycogen synthases after cellular uptake of circulating FAs by CD36 and glucose by glucose transporter 1 and 4 (GLUT1/GLUT4). After uptake, glucose is stored as glycogen or converted to acetyl-CoA which is converted into free fatty acids de novo by acetyl-CoA carboxylase (ACC) and fatty acid synthase (FASN) in the ER. TAGs are synthesized through re-esterification by several enzymes, including glycerol-3-phosphate acyltransferase (GPAT) and diacylglycerol O-acyltransferases (DGAT). Arrows indicate traficking, blunt-ends indicate inhibition and the plus sign indicates activation. Abbreviations: β3-AR, β3-adrenergic receptor; ATGL, desnutrin/adipose triglyceride lipase; cAMP, cyclic adenosinemonophosphate; CD36, cluster of differentiation 36; DGAT, diacylglycerol O-acyltransferase; DG, diacylglycerol; FA, fatty acid (represents NEFA and acyl-CoA pool); GPAT, glycerol-3-phosphate acyltransferase; GLUT, glucose transporter; HSL, hormone-sensitive lipase; LAL, lysosomal acid lipase; LP, lipoprotein; LPL, lipoprotein lipase; MG, monoacylglycerol; MGL, monoacylglycerol lipase; mTOR, mammalian target of rapamycin; NEFA, non-esterified fatty acid; PI3K, phosphoinositide 3-kinase; PKA, protein kinase A; TG, triglyceride; TCA, tricarboxylic acid; UCP1, uncoupled protein 1.

tissues lacking UCP1 expression, UCP1-expressing thermogenic tissues uncouple ATP production from β-oxidation to produce heat. In mice, the brown adipose tissue (BAT) and subcutaneous (also referred to as inguinal or beige) white adipose tissue (scWAT) exhibit high abundance of UCP1 expression, which can be further increased following cold exposure [9,10]. On the other hand, in humans, brown and beige adipocytes interspersed within the white adipose tissue (WAT) in the suprascapular region contribute to non-shivering thermogenesis [10]. In addition to these thermogenic adipose tissues, browning can be induced in nonthermogenic tissues such as WAT in mice and scWAT in humans [11,12] in response to cold exposure and seasonal changes. The maintenance of core body temperature requires fatty acid utilization. This has been recently demonstrated by indirect calorimetry in humans, whereby subjects with increased BAT depots exhibited a lower respiratory quotient, which reflects utilization of fatty acids as energy source, compared to those who had minimal to undetectable BAT depots regardless of food intake or activity [13]. This is because BAT uses fatty acids that are channeled from the lipolysis of TG to fuel heat production for the maintenance of core body temperature (Fig. 1)

thermogenesis, a function of the mitochondria in the skeletal muscle, 2) cold exposure induces non-shivering thermogenesis, a function of the mitochondria in brown fat and 3) brown fat activation can be triggered by overfeeding, inducing diet-induced thermogenesis. This review will emphasize on two sets of enzymes that play a role in the activation and deactivation of fatty acids in the regulation of non-shivering and dietinduced thermogenesis. Non-shivering thermogenesis occurs when the proton gradient established in the mitochondrial intermembrane space by β-oxidation becomes uncoupled from ATP-synthase-mediated oxidative phosphorylation [3–5] (Fig. 1). This is controlled by a tissue specific proton leak channel called uncoupling protein 1 (UCP1) [6]. In addition to UCP1, two other uncoupling proteins have been identified. UCP2 is expressed at low levels in many tissues, while UCP3 is expressed preferentially in skeletal muscle. Although a case for their involvement in thermogenesis is generally lacking [7], a recent study has shown that the loss of UCP2 impairs cold-induced non-shivering thermogenesis via mechanisms that are independent from proton uncoupling [8]. Whereas β-oxidation leads to the production of ATP from ADP through ATP-synthase in 80

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Fig. 2. The role of acyl-CoA synthetases (ACS) and acyl-CoA thioesterases (ACOT) in thermogenic adipose tissues. Cold-induced noradrenalin release increases lipolysis resulting in increased intracellular fatty acid (FA) concentrations. These FAs fulfill two principal metabolic fates; 1) they can bind and activate uncoupling protein 1 (UCP1), switching on UCP1 mediated proton (H+) conductance, or 2) activated FA-CoAs can be transported into the mitochondria via the carnitine palmitoyl transferase (CPT) system. After entering the mitochondria FA-CoAs can be used for fatty acid oxidation (βoxidation), potentiating anaplerosis and providing reducing equivalents for the electron transport chain (ETC). Activity of the ETC and ATP synthase can be coupled by the proton gradient. However, after UCP1 activation, UCP1 induces a basal leak of protons across the membrane, uncoupling ATP synthesis from the proton gradient. Proton translocation by this leak pathway prevents fixation of energy and releases it as heat. The ACS and ACOT family can influence brown adipocyte thermogenesis by 1) affecting the intracellular concentration of FAs versus fatty acid-CoAs, which will affect UCP1 activation and CPT1 mediated Acyl-CoA transport, and 2) ACS and ACOT members in the mitochondrial membrane may affect acyl-CoA substrate availability for mitochondrial β-oxidation. The ACS and ACOT family members that have been shown to directly influence adipose tissue thermogenesis are shown in the upper right panels. The enzymes that have not been directly implied in thermogenesis but might play a role in this process are listed in the lower sections of these panels. Arrows indicate trafficking, and plus signs indicate activation. Abbreviations: β3-AR, β3-adrenergic receptor; ACOT, acyl-CoA thioesterase; ACS, acyl-CoA synthetase; ATGL, desnutrin/adipose triglyceride lipase; cAMP, cyclic adenosinemonophosphate; CPT, carnitine palmitoyl transferase; DG, diacylglycerol; ETC, electron transport chain; NEFA, non-esterified fatty acid; HSL, hormonesensitive lipase; MG, monoacylglycerol; MGL, monoacylglycerol lipase; PKA, protein kinase A; TCA, tricarboxylic acid; TG, triglyceride; UCP1, uncoupled protein 1.

[23]. Secondly, fatty acids directly bind and activate UCP1 to induce heat production without contributing to mitochondrial β-oxidation as substrate [24]. Finally, the concentrations of NEFA and fatty acyl-CoA may activate transcription factors that regulate BAT thermogenesis and differentiation such as members of the peroxisomal proliferator activated receptor (PPAR) family [23]. In this review, we will focus on the ACS and ACOT family of enzymes that can play a role in the interplay between fatty acid activation and thermogenesis (Fig. 2).

[14–16]. However, following lipolysis, non-esterified fatty acids (NEFA) as such are chemically inert and need to be activated to form fatty acyl-CoAs by the acyl-CoA synthetase (ACS) family members before they can be taken up by the mitochondria and directed towards thermogenesis [17]. The activation of fatty acids is counter-regulated by another set of enzymes, namely acyl-CoA thioesterases (ACOT), which hydrolyze and deactivate fatty acyl-CoA into NEFA and CoA [18]. The metabolic impact of ACS and ACOT family members is influenced by a wide array of tissue expression patterns, subcellular localizations and substrate specificities (degree of saturation, fatty acid chain length) [17,19]. The characteristics of ACS and ACOT family members in substrate specificity, tissue expression and overall metabolic function have been eloquently reviewed elsewhere [19–22]. Here, we will elaborate on their potential involvement in thermogenesis (Fig. 2). The expression of these enzymes is highly regulated by metabolic demand as well as cold exposure whereby the balance between fatty acids and their CoA esters can influence the thermogenic regulation through multiple pathways. First, the conversion rate of NEFA to their CoA esters determines the substrate availability for mitochondrial β-oxidation

2. Fatty acid activation: Acyl-CoA synthetase family Fatty acid metabolism is initiated by the ACS family, which is comprised of 26 distinct members [25], and catalyzes the esterification of fatty acids to acyl-CoA with the use of a single ATP molecule (Table 1) [26]. The carbon chain length of the fatty acid species defines the substrate specificity for the different ACS, dividing them into five subfamilies: 1) the very long-chain synthetases (ACSVL) (> 21 carbon atoms), also known as fatty acid transport proteins (FATP), 2) the longchain acyl-CoA synthases (ACSL) (13–21 carbon atoms), 3) the medium-chain acyl-CoA synthetases (ACSM) (6–12 carbon atoms) and 81

Substrate

Catalyzes VLCFA and LCFA-CoA synthesis [47]

Acsl6

82

Catalyzes MCFA-CoA synthesis [154]

Catalyzes MCFA-CoA synthesis [154]

Catalyzes MCFA-CoA synthesis [154]

Acsm3

Acsm4

Acsm5

Catalyzes acetyl-CoA synthesis [157]

Acss3

Acyl-CoA synthetase family Acsf1 Activates ketone body acetoacetate to acetoacetyl-CoA synthetase Acsf2 Is a propionyl-CoA synthetase Acsf3 Is a malonyl-CoA synthetase

Catalyzes acetyl-CoA synthesis [22]

Acss2

Short-chain acyl-CoA synthetases Acss1 Catalyzes acetyl-CoA synthesis [155]

Liver [154]

Catalyzes MCFA-CoA synthesis [154]

Acsm2b

BAT BAT

Kidney, heart and brain [59]

Placenta, kidney, testis, BAT [156] Mammary gland, subcutaneous fat and liver [22] Adipose tissue, liver, kidney [157]

Olfactory epithelium [154]

Kidney [154]

Liver and kidney [22]

Medium-chain acyl-CoA synthetases Acsm1 Catalyzes MCFA-CoA synthesis [22] Acsm2a Catalyzes MCFA-CoA synthesis [154]

Acsl5

Catalyzes LCFA-CoA synthesis with a substrate preference for arachidonic acid [152] Catalyzes LCFA-CoA synthesis [153]

Acsl4

Adrenal glands and other steroid producing organs [45] Small intestine, liver and BAT [153] Brain [47]

Cerebrum and LDs [151]

Catalyzes LCFA-CoA synthesis [151]

Acsl3

Heart [148]

BAT, WAT, skeletal muscle, heart and skin [148] Liver [148]

Liver, adipose tissue [54]

Increase LCFA uptake and catalyzes VLCFA-CoA synthesis [20] Increase LCFA uptake, catalyzes VLCFA-CoA synthesis and exhibits bile acid-CoA synthetase activity [20] Increase LCFA uptake and catalyzes VLCFA-CoA synthesis [20]

Liver and testis [148]

Adipose tissue, heart and skeletal muscle [146] Liver and kidney [148]

Tissue expression

Long-chain acyl-CoA synthetases Acsl1 Catalyzes LCFA-CoA synthesis [54]

Fatp6

Fatp5

Fatp4

Fatty acid transport proteins Fatp1 Increases LCFA uptake and catalyzes VLCFA-CoA synthesis [145] Fatp2 Increases LCFA uptake, has bile acid-CoA ligase activity and catalyzes VLCFA-CoA synthesis [147] Fatp3 Catalyzes VLCFA-CoA synthesis [20]

Gene

Acyl-CoA synthetase family

Mitochondria [57]

Mitochondria (protein atlas)

Cytoplasma [22]

Mitochondria [156]

Mitochondria [22] Mitochondria (protein atlas) Mitochondria (protein atlas) Mitochondria (protein atlas) Mitochondria (protein atlas) Mitochondria (protein atlas)

Plasma membrane [47]

Mitochondria [43]

ER and cytosolic lipid droplets [151]

Mitochondria [54]

Sarcolemma of cardiomyocytes

ER and mitochondria (protein atlas) Plasma membrane or ER [149] Plasma membrane [150]

ER and peroxisome [147]

Plasma membrane

Subcellular localization

Table 1 Substrate specificity, tissue expression, subcellular localization and thermogenic role of acyl-CoA synthetase (ACS) and acyl-CoA thioesterase (ACOT) family members.

(continued on next page)

Among the top cold-regulated BAT proteins in mice (ref)

ASCL1A−/− mice showed lower fatty acid oxidation and intolerance to cold temperatures [23] Modulates lipogenic transcription factors such as PPARγ, ChREB and SREBP-1c [39–41]

Reduced TG accumulation in 3 T3-L1 adipocytes [30] Fatp5−/− mice showed higher O2 consumption and CO2 production rates and increased body temperature [31]

Essential for fatty acid uptake in response to β3adrenergic stimulation in mice [28]

Role in thermogenesis

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BBA - Molecular and Cell Biology of Lipids 1864 (2019) 79–90

Is a aminoadipate-semialdehyde dehydrogenase [60]

Acsf4

Acetyl-CoAs to LCFA-CoAs [72,162]

Acetyl-CoA and to lesser extent propionyl-, butyryl- and acetoacetyl-CoAs [163] LCFA-CoAs and short-chain aromatic acyl-CoAs [74,164]

MCFA-CoA and LCFA-CoAs [165]

Medium- and long-chain saturated and unsaturated acyl-CoAs [89]

Acot9–10

Acot11/Them1

Acot12

Acot13/Them2

Acot14/Them4

Acot15/Them5

LCFA-CoAs [80,160]

Type II Acots Acot7

Medium- to long-chain and methyl branched acyl-CoAs, intermediates of β-oxidation and bile acid-CoAs [84] LCFA, saturated and branched SCFA-CoAs [81,82]

MCFA-CoAs [86] Methyl-branched acyl-CoAs [86]

Acot5 Acot6

Acot8

LCFA-CoAs [158] Short dicarboxylic acyl-CoAs [158]

Long-chain saturated- and mono-unsaturated acyl-CoAs [158,159] LCFA-CoAs [158]

Substrate

Acot3 Acot4

Acot2

Type I Acots Acot1

Gene

Acyl-CoA thioesterase family

Substrate

Gene

Acyl-CoA synthetase family

Table 1 (continued)

83 Liver [89]

Liver, kidney and small intestine [163] Liver, heart, kidney and BAT [74,164]

Kidney, adipose tissue, brain and liver [81,82] BAT [72,162]

Brain, testis, pancreas and BAT [160,161] Liver and lung [84]

Liver, kidney, lung and heart [158,159] BAT, liver, heart and skeletal muscle [158] Kidney and liver Liver, kidney and proximal intestine [158] WAT, brain and intestine [86] WAT and kidney

Tissue expression

Tissue expression

Mitochondria MCFA-CoA and LCFA-CoAs [165] Mitochondria [89]

Cytosol, mitochondria and ER [72,162] Cytosol and a minor fraction in peroxisomes Mitochondria [74,164]

Mitochondria [81,82]

Peroxisome [84]

Cytosol [160,161]

Peroxisome [158] Peroxisome [158]

Peroxisome [158] Peroxisome [158]

Mitochondria [158]

Cytosol [158,159]

Subcellular localization

Role in thermogenesis

Suppresses cold-induced thermogenesis [77]

Suppresses thermogenesis in mice by reducing BAT oxygen consumption rates through decreased mitochondrial oxidation of endogenous fatty acids [70]

Role in thermogenesis

Subcellular localization

S. Steensels, B.A. Ersoy

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ACSL3–5 mRNA [36] and mitochondrial protein [37] levels are upregulated after cold exposure. ACSL3 is primarily localized to LD [38] where it promotes LD formation. ACSL3 may also indirectly influence thermogenesis through the modulation of several lipogenic transcription factors such as PPARγ, ChREB and SREBP-1c [39–41]. However, a direct link between thermogenesis and ACSL3 is yet to be investigated. TG synthesis is promoted by ACSL5 overexpression and decreased after ACSL5 genetic ablation, suggesting a role in TG homeostasis [39,42]. Although ACSL5 overexpression did not alter β-oxidation in rat hepatoma cells [43], its localization to the BAT mitochondria suggests that it might play a role in thermogenesis whether by contributing to the expansion of the TG pool or by activating FAs for β-oxidation. Indicative of a possible thermogenic function, several genomics studies (compiled in [44]) and a proteomics study have shown that the expression of ACSL5 increases up to 10-fold following chronic cold exposure, which ranks among the top cold-regulated BAT mitochondrial proteins in mice [37]. Future studies modulating ACSL5 activity in BAT should clarify whether it is essential in BAT fatty acid metabolism and thermogenesis. Due to their different localization, the other ACSLs have been implicated in other non-thermogenic metabolic pathways; ACSL4 expression is highest in adrenal glands and is related to steroid hormones [45] and prostaglandins [46]. ASCL6 is highly expressed in the brain [36], and its overexpression increases total cellular phospholipid production [47].

the 4) short-chain acyl-CoA synthetases (ACSS) (< 6 carbon atoms). A fifth subfamily was coined the acetyl-CoA synthetase family (ACSF) because of distant sequence homology compared to other ACS family members. 2.1. FATP The FATP family consists of 6 members (FATP1–6) which can both increase fatty acid uptake of long- and very long-chain fatty acids and exert ACSVL activity on only very long-chain fatty acids. Murine FATP1 (ACSVL4), FATP2 (ACSVL1), and FATP4 (ACSVL5) can increase longchain fatty acid (LCFA) uptake and catalyze the formation of very longchain fatty acyl CoAs, while FATP3 (ACSVL3) and FATP6 (ACSVL2) only exhibit ACSVL activity [27]. FATP1, FATP4 and FATP5 (ACSVL6) have been shown to play a role in thermogenesis while the other FATP family members have other functions in various tissues. FATP1 is expressed on the BAT plasma membrane, and its mRNA expression is increased after cold exposure (Table 1) [28]. Furthermore, genetic ablation of FATP1 decreases cold-induced fatty acid uptake into the BAT of mice, resulting in smaller LD and impairing their ability to defend core body temperature at 4 °C [28]. Experiments using the BATderived cell line HIB-1B confirmed that FATP1 is essential for fatty acid uptake in response to β3-adrenergic receptor (β3-AR) stimulation [28]. FATP4 is a closely related paralogue of FATP1 which is expressed in both BAT and WAT [29]. Whereas the lethality of a homozygous FATP4 knockout mouse model (Fatp4−/−) has complicated studies designed to characterize its biological function, knockdown of either FATP1 or FATP4 similarly reduced TG accumulation in 3 T3-L1 adipocytes [30]. FATP5 predominantly regulates fatty acid uptake in the liver, where it also exhibits bile acid-CoA synthetase activity [31]. Furthermore, FATP5 knockout (Fatp5−/−) mice showed higher O2 consumption and CO2 production rates, suggesting an increase in diet-induced thermogenesis. This was paralleled by an increase in body temperature but was not reflected in differences in β-oxidation, suggesting that FATP5 may regulate thermogenesis through alternate pathways rather than direct substrate availability [31]. The other FATP family members have not been linked to thermogenesis. FATP2 is highly expressed in the liver and kidneys and has been shown to modulate TG accumulation in renal cells [32]. FATP3 is expressed in the lungs, adrenal cortex, ovaries and testis [29,33], but little is known about its function. FATP6 is the major FATP family member expressed in the heart, where it is involved in heart LCFA transport [34].

2.3. ACSM MCFAs are effective mitochondrial substrates because unlike LCFA they can enter the mitochondrial matrix without the assistance of a membrane transporter, where they can be activated by ACSMs. However, they exert weak protonophoric and lytic activities in mitochondria and do not impair the electron transport in the respiratory chain [48]. Nonetheless, there is not a substantial amount of literature on the ACSM family, and much is left to be discovered about the role of MCFAs in thermogenesis. One study suggests that diets rich in mediumchain fatty acids (MCFA) reduce adiposity and increase thermogenesis in rats [49]. Although similar results were observed in overweight men, the role of BAT thermogenesis in MCFA-mediated regulation of energy homeostasis remains unclear [50]. Of interest is ACSM2, which preferentially activates MCFA [25]. ACSM2 protein is uniquely expressed in BAT as opposed to WAT in humans [51], and its protein and mRNA expression levels increase in response to cold exposure in BAT mitochondria of mice [37,52]. Obesity, which is associated with reduced diet-induced thermogenesis (discussed in more detail below), results in the downregulation of ACSM2 expression in the scWAT of obese men and women [53] as well as in diet-induced obese mice [54].

2.2. ACSL Mitochondria mostly utilize LCFA-CoA esters for β-oxidation. Therefore, the importance of ACSL members in thermogenesis has been of greater interest. The ACSL family consists of 5 members (ACSL1, 3–6), which are characterized by tissue and substrate specificity (Table 1). ACSL1 is the only ACSL that has been shown to play a role in thermogenesis. ACSL1 is responsible for up to 80% of the total ACSL activity in adipose tissue [23]. Adipose tissue-specific ablation of ACSL1 (ACSL1A−/−) in mice resulted in lower β-oxidation and intolerance to cold temperatures. Supporting a role in thermogenesis, brown adipocytes that were isolated from ACSL1A−/− mice failed to increase oxygen consumption in response to β3-AR stimulation despite exhibiting full UCP1 activity [23]. Taken together, these data suggest that the cold intolerance in these mice resulted from impaired activation of fatty acids by ACSL1 in BAT and demonstrate that ACSL1 is required for non-shivering thermogenesis [23]. In humans, an association with body temperature is lacking. Although single nucleotide polymorphisms of ACSL1 have been linked to increased fasting glucose in a large cohort study [35], contribution from impaired glucose clearance by BAT remains to be determined.

2.4. ACSS The fatty acids used for β-oxidation do not only originate from dietary lipids and lipolysis of cellular TG stores but can also be derived from de novo lipogenesis, which requires acetyl-CoA as substrate. This brings attention to one specific ACS group of proteins, ACSS1–3, that do not fall in the same category as the other ACS family members because they specifically facilitate the synthesis of acetyl-CoA from acetate and CoA (Table 1). Expression of ACSS1, 2 and 3 increases by 1.6-, 1.6- and 2.2-fold, respectively, during cold exposure in the BAT mitochondria of mice [37]. Furthermore, ACSS1 is specifically expressed in the mitochondria of BAT but not WAT in mice, further suggesting a possible involvement in thermogenesis [37]. The expression of ACSS2 and ACSS3 in scWAT is suppressed in obese individuals [53]. Nonetheless, to date, only ACSS2 has been directly linked to thermogenesis in mice whereby the loss of ACSS2 expression led to hypothermia, especially in the setting of low energy state such as fasting [55]. 84

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the top scWAT-specific proteins [51]. ACOT2 has also been shown to affect β-oxidation and increase the proton leak pathway in hepatic mitochondria although the effect of ACOT2 on UCP1-induced proton leak in BAT mitochondria has not been investigated [66]. In contrast to ACOT1 and ACOT2, mouse ACOT3–6 and human ACOT4 are peroxisomal thioesterases and only play a minor role in mitochondrial β-oxidation and UCP1-induced proton leak, and there is no evidence supporting a role in thermogenesis. The contribution of peroxisomal β-oxidation to thermogenesis is generally unknown. However, peroxisomal β-oxidation could indirectly contribute to thermogenesis by shortening VLCFA and facilitating acetyl-CoAs to mitochondria [67]. In this consideration, we will not focus on peroxisomal regulation of fatty acid activation in BAT in this review.

2.5. ACSF The ACSF family members acetoacetyl-CoA synthetase (ACSF1), acyl-CoA synthetase family member 2 (ACSF2), acyl-CoA synthetase family member 3 (ACSF3) and aminoadipate-semialdehyde dehydrogenase (ACSF4) are not assigned to other ACS subfamilies because their sequence differs substantially from other members [56]. ACSF2 and ACSF3 act as propionyl-CoA and malonyl-CoA synthetases, respectively, and are highly expressed in BAT. The propionyl-CoA generated by ACSF2 is normally metabolized to methylmalonyl-CoA, which can be converted into succinyl-CoA, a substrate for the citric acid cycle. Furthermore, Ellis et al. showed that cold exposure increased ACSF2 expression in the BAT [36]. As such, ACSF2 could possibly affect nonshivering thermogenesis by regulating the substrate availability for this pathway. ACSF3 is essential for mitochondrial fatty acid synthesis and mitochondrial protein malonylation [57,58]. However, its expression in BAT is not regulated by cold exposure suggesting that its effect on thermogenesis may be minor [36]. ACSF1 and ACSF4 do not activate fatty acids and have different substrate specificities. ACSF1 activates the ketone body acetoacetate to acetoacetyl-CoA and is highly expressed in kidney, heart and brain [59]. ACSF4 has been identified as a aminoadipate-semialdehyde dehydrogenase, but its biological function remains unknown [60].

3.2. Type II ACOTs Type II ACOTs (ACOT7–15) have a characteristic structural motif called the ‘Hotdog fold’ domain. This subfamily has been studied more extensively in thermogenesis, and previous studies have identified major regulatory roles for ACOT11 and ACOT13 (Table 1) [19]. ACOT11 is also referred to as thioesterase superfamily member 1 (THEM1) or StARD14, because it contains 2 hotdog fold domains and a lipid-binding START (steroid acute regulatory-related transfer) domain [19]. Notwithstanding its protein domain-based designations, ACOT11 was originally named brown-fat-inducible thioesterase (BFIT) due to its cold-inducibility (2–3 fold upregulation) in mouse BAT [44,68]. Although the factors that promote Acot11 expression remain unknown, it has been suggested that PKCβ-mediated phosphorylation activates Acot11 in vitro [69]. Metabolic monitoring of Acot11−/− mice under thermoneutral (30 °C) and cold (4 °C) temperatures revealed that ACOT11 suppressed thermogenesis in mice by reducing BAT oxygen consumption rates through decreased mitochondrial oxidation of endogenous acyl-CoAs [70]. In these mice, ACOT11 acted as a thioesterase in BAT and deactivated fatty acids that were hydrolyzed from TG by lipolysis, blocking their utilization in mitochondrial oxidation [70]. In contrast to mice, humans express 2 isoforms of ACOT11, known as BFIT1 and BFIT2. Whereas mouse ACOT11 localizes to the cytosol, mitochondria and ER [71], human BFIT2 has only been localized to the mitochondrial matrix of the BAT [72]. Therefore, further studies are needed to elucidate whether different human ACOT11 isoforms act similarly to mouse ACOT11 or instead regulate unique biological functions. ACOT13 (THEM2) is comprised of a single hotdog fold domain and preferentially hydrolyzes LCFA-CoA esters. It is highly expressed in oxidative tissues including thermogenic tissues such as mouse BAT [73,74]. Its expression increases after cold exposure (4 °C) and is regulated by the activity of PPARα [36,74]. The BAT of Acot13−/− mice displays reduced LD size and increased expression of thermogenic genes. Furthermore, cultured primary brown adipocytes from Acot13−/ − mice exhibit increased TG hydrolysis and oxygen consumption together with elevated expression of thermogenic genes in response to adrenergic stimulation [75]. Moreover, the loss of phosphatidylcholine transfer protein (PC-TP) expression, which binds and activates ACOT13 [74,76], similarly increases adaptive thermogenesis in mice [77]. Together these data clearly demonstrate that ACOT13 suppresses coldinduced thermogenesis. ACOT7 contains two hotdog fold domains, accounts for the majority of the thioesterase activity in the brain [78] and the endocrine pancreas [79] and plays a role in the inflammatory function of macrophages [80]. Notwithstanding its extra-thermogenic functions, ACOT7 is one of few thioesterases whose protein expression in the BAT increases in response to cold exposure in mice [37]. Surprisingly, its expression also increases with obesity in human scWAT [53]. Furthermore, the overexpression of an active form of ACOT7 in the adipose tissue of mice results in intolerance to cold exposure and reduced body temperature compared to control mice [36].

3. Fatty acid deactivation: Acyl-CoA thioesterase family The reaction catalyzed by the members of the ACS family is reversed by enzymes that hydrolyze acyl-CoA thioesters, generating NEFA and CoA. This enzymatic activity is performed by members of the ACOT family. Similar to the ACS family, the regulatory role of ACOT in fatty acid metabolism depends on their substrate specificity, tissue expression and subcellular localization. For example, deactivation of fatty acids at the ER may traffic fatty acids away from pathways associated with the ER membrane, such as glycerolipid biosynthesis [61]. Two structurally different ACOT types lead to a similar enzymatic activity in vitro, dividing the family into type I and type II ACOTs (Table 1). 3.1. Type I ACOTs Type I ACOTs (ACOT1–6) contain the α/β-hydrolase domain, which is also present in many lipases and esterases [19]. None of the type I ACOTs have been conclusively linked to the regulation of thermogenesis yet. However, the expression of type I ACOTs is generally enhanced in response to the activation of PPARα. Considering that the pharmacological activation of PPARα and PPARγ promotes UCP1 expression in primary brown adipocytes in vitro and BAT in vivo [62], the expression of type I ACOTs might play a regulatory role in β-oxidation and thermogenesis [63]. In fact, ACOT1 localizes to the cytosol and nucleus and regulates TG hydrolysis and PPARα activation through two independent mechanisms. Adenovirus-mediated knockdown of ACOT1 results in enhanced hepatic TG hydrolysis and β-oxidation. In contrast, the loss of ACOT1 also results in reduced activation of PPARα, a strong activator of βoxidation. Therefore, the catabolic activity of cytosolic ACOT1 appears to dominate over its nuclear function of providing fatty acid ligands for the activation of hepatic PPARα [64]. Furthermore, Ellis et al. showed that cold exposure significantly increased BAT ACOT1 mRNA expression levels in mice [36]. These studies suggest that the effect of ACOT1 on thermogenic factors may represent an interesting topic for future studies. ACOT2 has been reported to be important in brown adipocytes where its mRNA expression levels increase up to 18-fold during brown adipocyte differentiation and 2-fold in mouse BAT following cold exposure [44,52,65]. BAT mitochondrial ACOT2 protein expression increases upon cold exposure in mice [37]. Furthermore, proteomics studies that compared human WAT to scWAT ranked ACOT2 as one of 85

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thermogenic activity), pharmacotherapy (β3-AR and PPARα/γ agonists) and environmental factors including diet and exercise can influence thermogenesis through central and peripheral pathways. The latter has put mining thermogenic capacity forward as a mean of elevating metabolic rate and burning excess carbohydrates and fatty acids for the treatment of metabolic disorders that are associated with excessive nutrition such as type 2 diabetes and obesity [98]. Although BAT mainly utilizes fatty acids to promote thermogenesis [16], its contribution to the clearance of dietary fatty acids is minimal compared to other oxidative and adipose tissues [99]. On the other hand, BAT absorbs large quantities of glucose, especially in the setting of cold-exposure and following meals [13,100]. In fact, human BAT was initially identified by tracing the uptake of a glucose analogue, 18Ffluorodeoxyglucose into thermogenic tissue using positron-emission tomography (PET) [101]. The utilization of carbohydrates as energy source sharply increases postprandially and decreases afterwards in humans, as determined by the respiratory exchange quotient, which indicates substrate utilization [13]. The whole-body increase in postprandial glucose uptake is partially stimulated by increased pancreatic secretion of insulin following food intake [102]. Improvement of overall insulin sensitivity in humans results in increased glucose uptake by the BAT [103]. Obesity is associated with reduced insulin sensitivity, which explains its association with decreased glucose utilization as energy source in the human scWAT [53,103]. Since BAT possesses a great capacity for glucose uptake and metabolism, and an ability to regulate insulin sensitivity, improving glucose clearance by BAT has been proposed as a therapeutic approach to treat diabetes [104,105]. On the other hand, increased glucose uptake into BAT without improving its trafficking towards heat production could have undesirable consequences since the majority of glucose that is taken up by the BAT must be converted into TG before it can be used as a substrate in thermogenesis (Fig. 1) [14]. Its conversion into fatty acids includes the formation of acetyl-CoA, used to generate NEFA in BAT which will be esterified to replenish intracellular TG pools used during thermogenesis [14,15,105]. This is essential because channeling glucose away from lipogenesis [8] or inhibiting TG lipolysis [14,106] impairs cold-induced thermogenesis. A proteomics study, which compared changes in BAT protein expression between high fat diet-induced obese mice and chow diet-fed controls, has concluded that obesity results in reduced expression of lipogenic factors and increased expression of proteins that are involved in β-oxidation [54]. A similar study in human scWAT also identified obesity-induced suppression of proteins that mediate glucose uptake as well as lipogenesis [53]. Therefore, high fat diet-mediated adaptations to both BAT glucose uptake and its conversion into TG must be addressed to fully activate thermogenesis. Although the necessity of lipolysis in BAT thermogenesis is well established, the relative contribution of intracellular versus circulatory lipids may vary with the extent of cold challenge as well as nutritional state [107–111] and thus remains to be further elucidated. Earlier studies directly measuring arteriovenous differences across the BAT found increased glucose uptake and glycerol release but limited NEFA uptake after the induction of thermogenesis, suggesting that triglyceride hydrolysis was sufficient to fuel BAT thermogenesis [107]. This was confirmed in humans where acute cold exposure resulted in the depletion of intracellular BAT TG coupled with reduced plasma concentration of glucose but not TG, supporting the role of BAT intracellular lipolysis in response to acute cold challenge in humans [108]. However, recent transgenic mouse models with impaired intracellular BAT lipolysis demonstrate that thermogenesis depends on circulating lipids originating from WAT lipolysis rather than lipolysis within the BAT [110,111]. This does not appear to be a compensatory mechanism in response to the loss of intracellular BAT lipolysis because the loss of expression for CD36, a fatty acid and lipoprotein transporter, severely impairs BAT thermogenesis due to reduced uptake of both lipoprotein lipase (LPL)-hydrolyzed lipoproteins as well as holoparticles into BAT [109]. Furthermore, the loss of expression of lysosomal acid

ACOT9 contains two hotdog fold domains, localizes to the mitochondria [81] and exhibits ubiquitous tissue expression with higher levels in BAT, WAT and kidneys [82,83]. Acot9 mRNA and protein expression do not change after cold exposure [36,37]. However, unlike other ACOTs, ACOT9 is the only thioesterase that is exclusively phosphorylated in the BAT [70]. This post-translational modification of ACOT9 in the BAT suggests that it may play a role in rapid response to thermogenic stimuli. Nevertheless, the role of ACOT9 activity in BAT thermogenesis warrants further investigation. Due to their subcellular localization, the other type II ACOTs have been implicated in other metabolic pathways: similar to type I ACOTs, ACOT8 is a peroxisomal protein [84] that is not expressed in the BAT [70] and has been implicated in the pathophysiology of hepatocellular carcinoma [85]. Although BAT ACOT8 mRNA increases up to 2.5-fold following chronic cold exposure in mice [44,52], a biological function in thermogenesis has not been established. ACOT12, which is the paralogue of ACOT11, carries a peroxisomal targeting sequence although only a minor fraction of the protein associates with peroxisomes [86]. ACOT12 differs from ACOT11 in specifically targeting acetyl-CoA for hydrolysis. Although ACOT12 could hypothetically reverse the activities of ACSS1–3 (discussed above), ACOT12 mRNA levels are nearly undetectable in the BAT of mice [36]. Nevertheless, ACOT12 mRNA increases 4.5- and 16-fold following 3 [44] and 10 [52] days of chronic cold exposure, respectively. ACOT14 (THEM4) was initially characterized as the C-terminal modulator protein (CTMP) because it binds the Cterminus and regulates the activity of Akt [87,88]. Although ACOT14 localizes to the mitochondria and exhibits thioesterase activity towards MCFA and LCFA-CoA esters [19], its role in fatty acid metabolism remains unclear. ACOT15 (THEM5) has high substrate specificity for linoleyl-CoAs, which are essential for cardiolipin remodeling. Mice lacking ACOT15 exhibit altered cardiolipin remodeling, reduced mitochondrial fatty acid levels and β-oxidation, which consequently promote the development of fatty liver disease [89]. 4. Fatty acid-mediated modulation of thermogenic genes Besides serving as substrates for β-oxidation, fatty acids also promote thermogenesis by directly binding and activating UCP1 [24]. Purine nucleotides inhibit the spontaneous proton leak through UCP1, and this inhibition can be lifted in the presence of fatty acids that act in competition with purine nucleotides in order to increase UCP1 activity [90,91]. However, the role of fatty acid activation in UCP1 activity modulation remains unclear. The upstream enhancer region of Ucp1 includes nuclear receptor binding sites called the peroxisome proliferator response element (PPAR), which regulate the transcription of UCP1 under the control of PPARα and PPARγ [92]. The activation of these transcription factors can be modulated by fatty acids and their CoA esters. Interestingly, fatty acyl-CoAs have been shown to act as antagonists for PPARα [93] and PPARγ [94], while NEFA have been shown to activate PPARα [95] and act as ligands that can bind PPARα, PPARγ and PPARδ [96]. However, the function of PPARδ remains to be determined. The activation of PPAR transcription factors by NEFA may act as a compensatory signal to increase fatty acid activation and β-oxidation: PPARα drives the expression of thermogenic genes, ACS family members [97] and the cytosolic ACOT1 [63], which shuttles fatty acids away from esterification and TG formation in the cytosol, towards β-oxidation. In contrast, the antagonistic effect of fatty acyl-CoA esters may serve as a negative feedback mechanism to balance and limit the thermogenic activity. 5. Harvesting thermogenic activity for the treatment of metabolic disorders In addition to cold exposure, which maintains thermogenesis through β3-AR, a wide variety of factors such as aging (decreases 86

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favor trafficking of glucose away from oxidation and towards lipogenesis and other metabolic pathways. In addition to affecting the BAT, cold exposure leads to selective and dramatic increases in cardiac glucose uptake [100]. It has been hypothesized that the increased use of glucose as an alternative fuel source is rather a consequence of heart's failure to metabolize fatty acids effectively under duress such as cold-induced hypertrophy [135,136]. While this results in increased lipid accumulation and lipotoxicity in the heart, it has been suggested that improving mitochondrial oxidation of fatty acids rather than regulating the use of glucose as cardiac fuel source would prove beneficial in increasing cardiac output. It should be also noted that, the heart size increases in response to exercise as well as cold exposure in order to meet the demands of the increased metabolism [137,138]. In exercise, the elevation in cardiac mass is due to moderate increases in left ventricle wall thickness (LVWT) in a majority of athletes. However, increases over 12 mm in LVWT has been associated with pathophysiological markers of hypertrophic cardiomyopathy (HCM), which may explain the predisposition of small number of individuals with unusually high LVWT to sport-related cardiovascular diseases [138]. Therefore monitoring HCM symptoms or predisposition to heart disease may be necessary while considering the introduction of lifestyle changes such as cryotherapy or high intensity exercise training.

lipase (LAL), which only hydrolyzes circulating lipoprotein following their endocytic uptake into BAT lysosomes, results in the depletion of BAT lipid droplets, increased accumulation of lysosomes and decreased uptake of circulatory lipids, which culminate in hypothermia in mice [112]. The importance of CD36-mediated uptake of lipoproteins into BAT in the maintenance of thermogenesis is further supported by the observation that the expression of CD36 and LPL increase (but not that of FATP1) in the mouse model with impaired intracellular BAT lipolysis [110]. While the differences between rodent and human lipid metabolism will have to be taken into account, future studies should aim to distinguish the reliance of BAT on intracellular versus circulatory lipids in response to acute versus chronic cold exposure. This may shed light into whether BAT switches fuel source to circulatory lipids only when intracellular lipids are depleted or depends on lipoproteins regardless of local lipid stores. In order to mine thermogenesis for the catabolism of obesity-induced excess glucose and lipids, improved fuel uptake and utilization must be followed by enhanced thermogenic capacity. However, the abundance of BAT UCP1 is not correlated with improved glucose homeostasis in the setting of excessive nutrition. In fact, UCP1 expression in the BAT is positively regulated by high-fat diet: A systematic review has identified 62 publications, from which 42 showed that UCP1 expression is positively regulated by high-fat feeding while 11 showed no effect, and 9 reported a decrease [113]. In contrast to BAT, the same review found that high-fat feeding tended to decrease the browning of WAT. This review identified 24 publications among which 11 publications reported significant decreases in UCP1 mRNA levels while 10 displayed either no difference or no detectable amount of UCP1 mRNA levels, and only 3 demonstrated increased UCP1 mRNA levels in WAT after high-fat diet [113]. In addition, cold-induced increases in thermogenic gene expression in the scWAT (browning) is greatly suppressed in obese humans compared to lean controls [11]. This suggests that promoting the browning of WAT as opposed to increasing the thermogenic capacity of existing BAT might be an effective approach to treat obesity-related metabolic disorders. To that effect, physical exercise has been proposed to promote WAT browning. Whereas the effect of exercise on BAT activity is controversial [114–118], several reports in rodents suggest that exercise results in increased browning of WAT [119–123]. Although it is surprising that exercise, an energy-consuming process, would induce the transformation of cells that inherently increase energy expenditure, one possible explanation is that the exerciseinduced decrease in fat mass may necessitate an increased heat production to facilitate the insulation of the body [124]. Although the effect of cold and exercise on browning in rodents is generally accepted, its effect in humans remains controversial [124–126]. Simultaneous activation of anabolic and catabolic pathways is a unique property of BAT [127]. The concerted activation of glycolysis, lipogenesis and fatty acid oxidation in BAT goes against the classical models of energy utilization such as the Randle Cycle, whereby competition for common substrates would blunt concurrent activation of all three pathways. BAT appears to sidestep this inhibitory crosstalk to maintain cold-induced lipogenesis and increase thermogenesis by optimizing glucose and fatty acid utilization, and by doing so, perhaps creating an additional futile cycle to supplement thermogenesis [128]. Compared to UCP1-dependent thermogenesis, futile cycles such as creatine cycling [129] and sarcolipin-mediated inhibition of sarcoplasmic/endoplasmic reticulum calcium ATPase (SERCA) [130] promote thermogenesis by futile ATP hydrolysis. Similarly, substrate cycling between lipogenesis and oxidation has been previously proposed to create a thermogenic futile cycle to prevent lipotoxicity in muscle [131]. In BAT, trafficking of glucose towards lipogenesis rather than immediate oxidation would result in the hydrolysis of 14 ATPs and therefore may represent an additional futile cycle that could supplement thermogenesis. We should also consider that BAT utilizes glucose as substrate for the synthesis of complex lipids and glycerol [132–134]. Therefore, BAT-specific activities of glucose metabolizing enzymes may

6. Concluding remarks Our understanding of the BAT fuel utilization in response to cold exposure and diet has improved vastly in recent years. Cumulative analysis of current research indicates that not only the induction of UCP1 expression but also the differential uptake of glucose into brown/ beige adipocytes and their conversion into TG play major roles in the contribution of thermogenic activity to overall metabolism in the setting of obesity and cold exposure. Whereas the influx of extracellular fatty acids and lipolysis of intracellular TG stores provide the essential fuel source for thermogenesis, these fatty acids must still undergo activation by specific ACS family members prior to their oxidation in the mitochondria. In contrast, the activation of these fatty acids will be counter-regulated by ACOT family members. Therefore, changes in the BAT-specific ACS and ACOT expression in response to cold exposure and diet contribute to the regulation of thermogenesis under duress. While cold exposure increases the expression levels of both ACS and ACOT in the BAT mitochondria of mice, thereby increasing activation of fatty acids by ACS and facilitating thermogenesis, the biological significance of increased ACOT activity in response to cold remains unclear. One possible explanation for the deactivation of BAT fatty acids is the preservation of energy in response to chronic cold stress. Another reason for the deactivation of fatty acids in BAT mitochondria could be the reduction of reactive oxygen species (ROS) stress. However, this is unlikely as BAT ROS promotes thermogenesis [129], and increasing the accumulation of superoxides in the adipose tissues above biological levels does not cause apoptosis but leads to increased thermogenic activity [139]. It is conceivable that ACOTs could have an indirect role in energy conservation by this ROS reduction. More consistent with the model invoking reduced β-oxidation as a contributor to metabolic dysfunction, obesity results in moderate decreases in ACS activity and slight increases in ACOT activity in the scWAT of humans. Although high fat diet also increases ACOT activity in the BAT of mice, the expression of ACS is more prominently upregulated, once again displaying inconsistencies between mice and humans. Furthermore, identifying a role for ACOT in the preservation of energy becomes more convoluted using mouse models as extended cold-exposure leads to torpor whereby the body temperature drops to a lower resting state. Therefore, future studies to investigate the effect of cold exposure on the activities of ACS and ACOT in human brown and beige adipocytes would clarify the relevance to human pathophysiology. 87

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Overall, our analysis indicates that in addition to targeting the induction of glucose uptake and lipogenesis, modulating enzymes that regulate the activation of fatty acids also represent a valuable target to improve thermogenic capacity and treatment of metabolic disorders related to obesity.

(7–8) (2008) 642–650. [25] P.A. Watkins, et al., Evidence for 26 distinct acyl-coenzyme A synthetase genes in the human genome, J. Lipid Res. 48 (12) (2007) 2736–2750. [26] D.G. Mashek, et al., Revised nomenclature for the mammalian long-chain acyl-CoA synthetase gene family, J. Lipid Res. 45 (10) (2004) 1958–1961. [27] C.C. Dirusso, et al., Comparative biochemical studies of the murine fatty acid transport proteins (FATP) expressed in yeast, J. Biol. Chem. 280 (17) (2005) 16829–16837. [28] Q. Wu, et al., Fatty acid transport protein 1 is required for nonshivering thermogenesis in brown adipose tissue, Diabetes 55 (12) (2006) 3229–3237. [29] H. Doege, A. Stahl, Protein-mediated fatty acid uptake: novel insights from in vivo models, Physiology (Bethesda) 21 (2006) 259–268. [30] S. Lobo, et al., Fatty acid metabolism in adipocytes: functional analysis of fatty acid transport proteins 1 and 4, J. Lipid Res. 48 (3) (2007) 609–620. [31] B. Hubbard, et al., Mice deleted for fatty acid transport protein 5 have defective bile acid conjugation and are protected from obesity, Gastroenterology 130 (4) (2006) 1259–1269. [32] A.C. Johnson, A. Stahl, R.A. Zager, Triglyceride accumulation in injured renal tubular cells: alterations in both synthetic and catabolic pathways, Kidney Int. 67 (6) (2005) 2196–2209. [33] Z. Pei, et al., Mouse very long-chain Acyl-CoA synthetase 3/fatty acid transport protein 3 catalyzes fatty acid activation but not fatty acid transport in MA-10 cells, J. Biol. Chem. 279 (52) (2004) 54454–54462. [34] R.E. Gimeno, et al., Characterization of a heart-specific fatty acid transport protein, J. Biol. Chem. 278 (18) (2003) 16039–16044. [35] A. Manichaikul, et al., Genetic association of long-chain acyl-CoA synthetase 1 variants with fasting glucose, diabetes, and subclinical atherosclerosis, J. Lipid Res. 57 (3) (2016) 433–442. [36] J.M. Ellis, C.E. Bowman, M.J. Wolfgang, Metabolic and tissue-specific regulation of acyl-CoA metabolism, PLoS One 10 (3) (2015) e0116587. [37] F. Forner, et al., Proteome differences between brown and white fat mitochondria reveal specialized metabolic functions, Cell Metab. 10 (4) (2009) 324–335. [38] D.J. Murphy, The biogenesis and functions of lipid bodies in animals, plants and microorganisms, Prog. Lipid Res. 40 (5) (2001) 325–438. [39] H. Yao, J. Ye, Long chain acyl-CoA synthetase 3-mediated phosphatidylcholine synthesis is required for assembly of very low density lipoproteins in human hepatoma Huh7 cells, J. Biol. Chem. 283 (2) (2008) 849–854. [40] F. Schroeder, et al., Role of fatty acid binding proteins and long chain fatty acids in modulating nuclear receptors and gene transcription, Lipids 43 (1) (2008) 1–17. [41] D.G. Mashek, L.O. Li, R.A. Coleman, Long-chain acyl-CoA synthetases and fatty acid channeling, Futur. Lipidol. 2 (4) (2007) 465–476. [42] H.A. Parkes, et al., Overexpression of acyl-CoA synthetase-1 increases lipid deposition in hepatic (HepG2) cells and rodent liver in vivo, Am. J. Physiol. Endocrinol. Metab. 291 (4) (2006) E737–E744. [43] D.G. Mashek, et al., Rat long chain acyl-CoA synthetase 5 increases fatty acid uptake and partitioning to cellular triacylglycerol in McArdle-RH7777 cells, J. Biol. Chem. 281 (2) (2006) 945–950. [44] A.B. Marcher, et al., RNA-seq and mass-spectrometry-based lipidomics reveal extensive changes of glycerolipid pathways in brown adipose tissue in response to cold, Cell Rep. 13 (9) (2015) 2000–2013. [45] X. Wu, et al., Long chain fatty Acyl-CoA synthetase 4 is a biomarker for and mediator of hormone resistance in human breast cancer, PLoS One 8 (10) (2013) e77060. [46] D.L. Golej, et al., Long-chain acyl-CoA synthetase 4 modulates prostaglandin E(2) release from human arterial smooth muscle cells, J. Lipid Res. 52 (4) (2011) 782–793. [47] J.R. Marszalek, et al., Long-chain acyl-CoA synthetase 6 preferentially promotes DHA metabolism, J. Biol. Chem. 280 (11) (2005) 10817–10826. [48] P. Schonfeld, L. Wojtczak, Short- and medium-chain fatty acids in energy metabolism: the cellular perspective, J. Lipid Res. 57 (6) (2016) 943–954. [49] N. Baba, E.F. Bracco, S.A. Hashim, Enhanced thermogenesis and diminished deposition of fat in response to overfeeding with diet containing medium chain triglyceride, Am. J. Clin. Nutr. 35 (4) (1982) 678–682. [50] M.P. St-Onge, et al., Medium-chain triglycerides increase energy expenditure and decrease adiposity in overweight men, Obes. Res. 11 (3) (2003) 395–402. [51] S. Muller, et al., Proteomic analysis of human brown adipose tissue reveals utilization of coupled and uncoupled energy expenditure pathways, Sci. Rep. 6 (2016) 30030. [52] M. Rosell, et al., Brown and white adipose tissues: intrinsic differences in gene expression and response to cold exposure in mice, Am. J. Physiol. Endocrinol. Metab. 306 (8) (2014) E945–E964. [53] A. Mardinoglu, et al., Defining the human adipose tissue proteome to reveal metabolic alterations in obesity, J. Proteome Res. 13 (11) (2014) 5106–5119. [54] J. Li, et al., Comparative proteome analysis of brown adipose tissue in obese C57BL/6J mice using iTRAQ-coupled 2D LC-MS/MS, PLoS One 10 (3) (2015) e0119350. [55] I. Sakakibara, et al., Fasting-induced hypothermia and reduced energy production in mice lacking acetyl-CoA synthetase 2, Cell Metab. 9 (2) (2009) 191–202. [56] P.A. Watkins, et al., Evidence for 26 distinct acyl-coenzyme A synthetase genes in the human genome, J. Lipid Res. 48 (12) (2007) 2736–2750. [57] C.E. Bowman, et al., The mammalian malonyl-CoA synthetase ACSF3 is required for mitochondrial protein malonylation and metabolic efficiency, Cell Chem. Biol. 24 (6) (2017) 673–684 (e4). [58] A. Witkowski, J. Thweatt, S. Smith, Mammalian ACSF3 protein is a malonyl-CoA synthetase that supplies the chain extender units for mitochondrial fatty acid synthesis, J. Biol. Chem. 286 (39) (2011) 33729–33736.

Transparency document The Transparency document associated with this article can be found, in online version. Acknowledgements This work was supported by NIH grant to B.A.E. (DK102733) and a 2017 Belgian American Education Foundation (BAEF) Postdoctoral Fellowship to S.S. References [1] E.S. Goetzman, Modeling disorders of fatty acid metabolism in the mouse, Prog. Mol. Biol. Transl. Sci. 100 (2011) 389–417. [2] P. Schrauwen, W.D. van Marken Lichtenbelt, Combatting type 2 diabetes by turning up the heat, Diabetologia 59 (11) (2016) 2269–2279. [3] K. Inokuma, et al., Indispensable role of mitochondrial UCP1 for antiobesity effect of beta3-adrenergic stimulation, Am. J. Physiol. Endocrinol. Metab. 290 (5) (2006) E1014–E1021. [4] C.L. Mattsson, et al., beta(1)-Adrenergic receptors increase UCP1 in human MADS brown adipocytes and rescue cold-acclimated beta(3)-adrenergic receptorknockout mice via nonshivering thermogenesis, Am. J. Physiol. Endocrinol. Metab. 301 (6) (2011) E1108–E1118. [5] J. Zhao, B. Cannon, J. Nedergaard, alpha1-Adrenergic stimulation potentiates the thermogenic action of beta3-adrenoreceptor-generated cAMP in brown fat cells, J. Biol. Chem. 272 (52) (1997) 32847–32856. [6] A. Matthias, et al., Thermogenic responses in brown fat cells are fully UCP1-dependent. UCP2 or UCP3 do not substitute for UCP1 in adrenergically or fatty scidinduced thermogenesis, J. Biol. Chem. 275 (33) (2000) 25073–25081. [7] M.E. Harper, K. Green, M.D. Brand, The efficiency of cellular energy transduction and its implications for obesity, Annu. Rev. Nutr. 28 (2008) 13–33. [8] A. Caron, et al., Loss of UCP2 impairs cold-induced non-shivering thermogenesis by promoting a shift toward glucose utilization in brown adipose tissue, Biochimie 134 (2017) 118–126. [9] P. Seale, et al., PRDM16 controls a brown fat/skeletal muscle switch, Nature 454 (7207) (2008) 961–967. [10] J. Wu, et al., Beige adipocytes are a distinct type of thermogenic fat cell in mouse and human, Cell 150 (2) (2012) 366–376. [11] P.A. Kern, et al., The effects of temperature and seasons on subcutaneous white adipose tissue in humans: evidence for thermogenic gene induction, J. Clin. Endocrinol. Metab. 99 (12) (2014) E2772–E2779. [12] M.A. Zuriaga, et al., Humans and mice display opposing patterns of "browning" gene expression in visceral and subcutaneous white adipose tissue depots, Front Cardiovasc. Med. 4 (2017) 27. [13] M. Hibi, et al., Brown adipose tissue is involved in diet-induced thermogenesis and whole-body fat utilization in healthy humans, Int. J. Obes. 40 (11) (2016) 1655–1661. [14] S.M. Labbe, et al., In vivo measurement of energy substrate contribution to coldinduced brown adipose tissue thermogenesis, FASEB J. 29 (5) (2015) 2046–2058. [15] Q. Hao, et al., Transcriptome profiling of brown adipose tissue during cold exposure reveals extensive regulation of glucose metabolism, Am. J. Physiol. Endocrinol. Metab. 308 (5) (2015) E380–E392. [16] J. Lee, J.M. Ellis, M.J. Wolfgang, Adipose fatty acid oxidation is required for thermogenesis and potentiates oxidative stress-induced inflammation, Cell Rep. 10 (2) (2015) 266–279. [17] D.E. Cooper, et al., Physiological consequences of compartmentalized Acyl-CoA metabolism, J. Biol. Chem. 290 (33) (2015) 20023–20031. [18] D.E. Cohen, New players on the metabolic stage: how do you like them acots? Adipocytes 2 (1) (2013) 3–6. [19] V. Tillander, S.E.H. Alexson, D.E. Cohen, Deactivating fatty acids: acyl-CoA thioesterase-mediated control of lipid metabolism, Trends Endocrinol. Metab. 28 (7) (2017) 473–484. [20] R.E. Gimeno, Fatty acid transport proteins, Curr. Opin. Lipidol. 18 (3) (2007) 271–276. [21] J. Storch, B. Corsico, The emerging functions and mechanisms of mammalian fatty acid-binding proteins, Annu. Rev. Nutr. 28 (2008) 73–95. [22] T. Fujino, et al., Molecular identification and characterization of two mediumchain acyl-CoA synthetases, MACS1 and the Sa gene product, J. Biol. Chem. 276 (38) (2001) 35961–35966. [23] J.M. Ellis, et al., Adipose acyl-CoA synthetase-1 directs fatty acids toward betaoxidation and is required for cold thermogenesis, Cell Metab. 12 (1) (2010) 53–64. [24] I.G. Shabalina, et al., Within brown-fat cells, UCP1-mediated fatty acid-induced uncoupling is independent of fatty acid metabolism, Biochim. Biophys. Acta 1777

88

BBA - Molecular and Cell Biology of Lipids 1864 (2019) 79–90

S. Steensels, B.A. Ersoy

[89] E. Zhuravleva, et al., Acyl coenzyme A thioesterase Them5/Acot15 is involved in cardiolipin remodeling and fatty liver development, Mol. Cell. Biol. 32 (14) (2012) 2685–2697. [90] I.G. Shabalina, et al., Native UCP1 displays simple competitive kinetics between the regulators purine nucleotides and fatty acids, J. Biol. Chem. 279 (37) (2004) 38236–38248. [91] S.G. Huang, Limited proteolysis reveals conformational changes in uncoupling protein-1 from brown adipose tissue mitochondria, Arch. Biochem. Biophys. 420 (1) (2003) 40–45. [92] G. Zhang, Q. Sun, C. Liu, Influencing factors of thermogenic adipose tissue activity, Front. Physiol. 7 (2016) 29. [93] M. Elholm, et al., Acyl-CoA esters antagonize the effects of ligands on peroxisome proliferator-activated receptor alpha conformation, DNA binding, and interaction with co-factors, J. Biol. Chem. 276 (24) (2001) 21410–21416. [94] K. Murakami, et al., Fatty-acyl-CoA thioesters inhibit recruitment of steroid receptor co-activator 1 to alpha and gamma isoforms of peroxisome-proliferatoractivated receptors by competing with agonists, Biochem. J. 353 (Pt 2) (2001) 231–238. [95] M. Gottlicher, et al., Fatty acids activate a chimera of the clofibric acid-activated receptor and the glucocorticoid receptor, Proc. Natl. Acad. Sci. U. S. A. 89 (10) (1992) 4653–4657. [96] B.M. Forman, J. Chen, R.M. Evans, Hypolipidemic drugs, polyunsaturated fatty acids, and eicosanoids are ligands for peroxisome proliferator-activated receptors alpha and delta, Proc. Natl. Acad. Sci. U. S. A. 94 (9) (1997) 4312–4317. [97] G. Martin, et al., Coordinate regulation of the expression of the fatty acid transport protein and acyl-CoA synthetase genes by PPARalpha and PPARgamma activators, J. Biol. Chem. 272 (45) (1997) 28210–28217. [98] N. Slocum, et al., Responses of brown adipose tissue to diet-induced obesity, exercise, dietary restriction and ephedrine treatment, Exp. Toxicol. Pathol. 65 (5) (2013) 549–557. [99] D.P. Blondin, et al., Dietary fatty acid metabolism of brown adipose tissue in coldacclimated men, Nat. Commun. 8 (2017) 14146. [100] A. Bartelt, et al., Brown adipose tissue activity controls triglyceride clearance, Nat. Med. 17 (2) (2011) 200–205. [101] H.W.D. Yeung, et al., Patterns of F-18-FDG uptake in adipose tissue and muscle: a potential source of false-positives for PET, J. Nucl. Med. 44 (11) (2003) 1789–1796. [102] K.L. Townsend, Y.H. Tseng, Brown fat fuel utilization and thermogenesis, Trends Endocrinol. Metab. 25 (4) (2014) 168–177. [103] M. Chondronikola, et al., Brown adipose tissue improves whole-body glucose homeostasis and insulin sensitivity in humans, Diabetes 63 (12) (2014) 4089–4099. [104] P. Lee, et al., Brown adipose tissue exhibits a glucose-responsive thermogenic biorhythm in humans, Cell Metab. 23 (4) (2016) 602–609. [105] M.K. Hankir, M.A. Cowley, W.K. Fenske, A BAT-centric approach to the treatment of diabetes: turn on the brain, Cell Metab. 24 (1) (2016) 31–40. [106] D.P. Blondin, et al., Inhibition of intracellular triglyceride lipolysis suppresses cold-induced Brown adipose tissue metabolism and increases shivering in humans, Cell Metab. 25 (2) (2017) 438–447. [107] S.W. Ma, D.O. Foster, Uptake of glucose and release of fatty acids and glycerol by rat brown adipose tissue in vivo, Can. J. Physiol. Pharmacol. 64 (5) (1986) 609–614. [108] V. Ouellet, et al., Brown adipose tissue oxidative metabolism contributes to energy expenditure during acute cold exposure in humans, J. Clin. Invest. 122 (2) (2012) 545–552. [109] A. Bartelt, et al., Thermogenic adipocytes promote HDL turnover and reverse cholesterol transport, Nat. Commun. 8 (2017) 15010. [110] H. Shin, et al., Lipolysis in brown adipocytes is not essential for cold-induced thermogenesis in mice, Cell Metab. 26 (5) (2017) 764–777 (e5). [111] R. Schreiber, et al., Cold-induced thermogenesis depends on ATGL-mediated lipolysis in cardiac muscle, but not brown adipose tissue, Cell Metab. 26 (5) (2017) 753–763 (e7). [112] M. Duta-Mare, et al., Lysosomal acid lipase regulates fatty acid channeling in brown adipose tissue to maintain thermogenesis, Biochim. Biophys. Acta 1863 (4) (2018) 467–478. [113] T. Fromme, M. Klingenspor, Uncoupling protein 1 expression and high-fat diets, Am. J. Phys. Regul. Integr. Comp. Phys. 300 (1) (2011) R1–R8. [114] X. Xu, et al., Exercise ameliorates high-fat diet-induced metabolic and vascular dysfunction, and increases adipocyte progenitor cell population in brown adipose tissue, Am. J. Phys. Regul. Integr. Comp. Phys. 300 (5) (2011) R1115–R1125. [115] K. Hirata, T. Nagasaka, Enhancement of calorigenic response to cold and to norepinephrine in physically trained rats, Jpn. J. Physiol. 31 (5) (1981) 657–665. [116] S.J. Wickler, et al., Thermogenic capacity and brown fat in rats exercise-trained by running, Metabolism 36 (1) (1987) 76–81. [117] M.V. Wu, et al., Thermogenic capacity is antagonistically regulated in classical brown and white subcutaneous fat depots by high fat diet and endurance training in rats: impact on whole-body energy expenditure, J. Biol. Chem. 289 (49) (2014) 34129–34140. [118] O. Boss, et al., Effect of endurance training on mRNA expression of uncoupling proteins 1, 2, and 3 in the rat, FASEB J. 12 (3) (1998) 335–339. [119] P. Bostrom, et al., A PGC1-alpha-dependent myokine that drives brown-fat-like development of white fat and thermogenesis, Nature 481 (7382) (2012) 463–468. [120] K.I. Stanford, et al., A novel role for subcutaneous adipose tissue in exercise-induced improvements in glucose homeostasis, Diabetes 64 (6) (2015) 2002–2014. [121] L.N. Sutherland, et al., Exercise and adrenaline increase PGC-1{alpha} mRNA expression in rat adipose tissue, J. Physiol. 587 (Pt 7) (2009) 1607–1617.

[59] M. Ohgami, et al., Expression of acetoacetyl-CoA synthetase, a novel cytosolic ketone body-utilizing enzyme, in human brain, Biochem. Pharmacol. 65 (6) (2003) 989–994. [60] L. Wang, et al., Cloning and characterization of a novel human homolog⁎ of mouse U26, a putative PQQ-dependent AAS dehydrogenase, Mol. Biol. Rep. 32 (1) (2005) 47–53. [61] B.A. Ersoy, et al., Thioesterase-mediated control of cellular calcium homeostasis enables hepatic ER stress, J. Clin. Invest. 128 (1) (2018) 141–156. [62] M.J. Barbera, et al., Peroxisome proliferator-activated receptor alpha activates transcription of the brown fat uncoupling protein-1 gene. A link between regulation of the thermogenic and lipid oxidation pathways in the brown fat cell, J. Biol. Chem. 276 (2) (2001) 1486–1493. [63] B. Dongol, et al., The acyl-CoA thioesterase I is regulated by PPARalpha and HNF4alpha via a distal response element in the promoter, J. Lipid Res. 48 (8) (2007) 1781–1791. [64] M.P. Franklin, A. Sathyanarayan, D.G. Mashek, Acyl-CoA thioesterase 1 (ACOT1) regulates PPARalpha to couple fatty acid flux with oxidative capacity during fasting, Diabetes 66 (8) (2017) 2112–2123. [65] A. Momose, et al., Regulated expression of acyl-CoA thioesterases in the differentiation of cultured rat brown adipocytes, Biochem. Biophys. Res. Commun. 404 (1) (2011) 74–78. [66] C. Moffat, et al., Acyl-CoA thioesterase-2 facilitates mitochondrial fatty acid oxidation in the liver, J. Lipid Res. 55 (12) (2014) 2458–2470. [67] I.J. Lodhi, C.F. Semenkovich, Peroxisomes: a nexus for lipid metabolism and cellular signaling, Cell Metab. 19 (3) (2014) 380–392. [68] S.H. Adams, et al., BFIT, a unique acyl-CoA thioesterase induced in thermogenic brown adipose tissue: cloning, organization of the human gene and assessment of a potential link to obesity, Biochem. J. 360 (Pt 1) (2001) 135–142. [69] Y. Li, et al., N-terminal phosphorylation of thioesterase superfamily member 1 (Them1) regulates its subcellular localization in brown adipocytes, FASEB J. 31 (S1) (2017). [70] K. Okada, et al., Thioesterase superfamily member 1 suppresses cold thermogenesis by limiting the oxidation of lipid droplet-derived fatty acids in brown adipose tissue, Mol. Metab. 5 (5) (2016) 340–351. [71] Y.Z. Zhang, et al., Targeted deletion of thioesterase superfamily member 1 promotes energy expenditure and protects against obesity and insulin resistance, Proc. Natl. Acad. Sci. U. S. A. 109 (14) (2012) 5417–5422. [72] D. Chen, et al., Human brown fat inducible thioesterase variant 2 cellular localization and catalytic function, Biochemistry 51 (35) (2012) 6990–6999. [73] J. Cao, et al., The mechanisms of human hotdog-fold thioesterase 2 (hTHEM2) substrate recognition and catalysis illuminated by a structure and function based analysis, Biochemistry 48 (6) (2009) 1293–1304. [74] J. Wei, H.W. Kang, D.E. Cohen, Thioesterase superfamily member 2 (Them2)/acylCoA thioesterase 13 (Acot13): a homotetrameric hotdog fold thioesterase with selectivity for long-chain fatty acyl-CoAs, Biochem. J. 421 (2) (2009) 311–322. [75] H.W. Kang, et al., Thioesterase superfamily member 2/Acyl-CoA thioesterase 13 (Them2/Acot13) regulates adaptive thermogenesis in mice, J. Biol. Chem. 288 (46) (2013) 33376–33386. [76] K. Kanno, et al., Interacting proteins dictate function of the minimal START domain phosphatidylcholine transfer protein/StarD2, J. Biol. Chem. 282 (42) (2007) 30728–30736. [77] H.W. Kang, et al., Mice lacking Pctp/StarD2 exhibit increased adaptive thermogenesis and enlarged mitochondria in brown adipose tissue, J. Lipid Res. 50 (11) (2009) 2212–2221. [78] J. Yamada, Long-chain acyl-CoA hydrolase in the brain, Amino Acids 28 (3) (2005) 273–278. [79] A. Martinez-Sanchez, et al., Disallowance of Acot7 in beta-cells is required for normal glucose tolerance and insulin secretion, Diabetes 65 (5) (2016) 1268–1282. [80] J.K. Forwood, et al., Structural basis for recruitment of tandem hotdog domains in acyl-CoA thioesterase 7 and its role in inflammation, Proc. Natl. Acad. Sci. U. S. A. 104 (25) (2007) 10382–10387. [81] V. Poupon, et al., Molecular cloning and characterization of MT-ACT48, a novel mitochondrial acyl-CoA thioesterase, J. Biol. Chem. 274 (27) (1999) 19188–19194. [82] V. Tillander, et al., Acyl-CoA thioesterase 9 (ACOT9) in mouse may provide a novel link between fatty acid and amino acid metabolism in mitochondria, Cell. Mol. Life Sci. 71 (5) (2014) 933–948. [83] E.L. Huttlin, et al., A tissue-specific atlas of mouse protein phosphorylation and expression, Cell 143 (7) (2010) 1174–1189. [84] M.C. Hunt, V. Tillander, S.E. Alexson, Regulation of peroxisomal lipid metabolism: the role of acyl-CoA and coenzyme a metabolizing enzymes, Biochimie 98 (2014) 45–55. [85] Y.H. Hung, et al., Fatty acid metabolic enzyme acyl-CoA thioesterase 8 promotes the development of hepatocellular carcinoma, Oncol. Rep. 31 (6) (2014) 2797–2803. [86] M.A. Westin, M.C. Hunt, S.E. Alexson, Short- and medium-chain carnitine acyltransferases and acyl-CoA thioesterases in mouse provide complementary systems for transport of beta-oxidation products out of peroxisomes, Cell. Mol. Life Sci. 65 (6) (2008) 982–990. [87] S.M. Maira, et al., Carboxyl-terminal modulator protein (CTMP), a negative regulator of PKB/Akt and v-Akt at the plasma membrane, Science 294 (5541) (2001) 374–380. [88] H. Ono, et al., Carboxy-terminal modulator protein induces Akt phosphorylation and activation, thereby enhancing antiapoptotic, glycogen synthetic, and glucose uptake pathways, Am. J. Phys. Cell Phys. 293 (5) (2007) C1576–C1585.

89

BBA - Molecular and Cell Biology of Lipids 1864 (2019) 79–90

S. Steensels, B.A. Ersoy

[122] E. Trevellin, et al., Exercise training induces mitochondrial biogenesis and glucose uptake in subcutaneous adipose tissue through eNOS-dependent mechanisms, Diabetes 63 (8) (2014) 2800–2811. [123] L. Cao, et al., White to brown fat phenotypic switch induced by genetic and environmental activation of a hypothalamic-adipocyte axis, Cell Metab. 14 (3) (2011) 324–338. [124] K.I. Stanford, L.J. Goodyear, Exercise regulation of adipose tissue, Adipocytes 5 (2) (2016) 153–162. [125] M.J. Vosselman, et al., Low brown adipose tissue activity in endurance-trained compared with lean sedentary men, Int. J. Obes. 39 (12) (2015) 1696–1702. [126] B.P. Leitner, et al., Mapping of human brown adipose tissue in lean and obese young men, Proc. Natl. Acad. Sci. U. S. A. 114 (32) (2017) 8649–8654. [127] X.X. Yu, et al., Cold elicits the simultaneous induction of fatty acid synthesis and beta-oxidation in murine brown adipose tissue: prediction from differential gene expression and confirmation in vivo, FASEB J. 16 (2) (2002) 155–168. [128] J. Sanchez-Gurmaches, et al., Brown fat AKT2 is a cold-induced kinase that stimulates ChREBP-mediated de novo lipogenesis to optimize fuel storage and thermogenesis, Cell Metab. 27 (1) (2018) 195–209 (e6). [129] E.T. Chouchani, L. Kazak, B.M. Spiegelman, Mitochondrial reactive oxygen species and adipose tissue thermogenesis: bridging physiology and mechanisms, J. Biol. Chem. 292 (41) (2017) 16810–16816. [130] N.C. Bal, et al., Sarcolipin is a newly identified regulator of muscle-based thermogenesis in mammals, Nat. Med. 18 (10) (2012) 1575–1579. [131] A.G. Dulloo, et al., Substrate cycling between de novo lipogenesis and lipid oxidation: a thermogenic mechanism against skeletal muscle lipotoxicity and glucolipotoxicity, Int. J. Obes. Relat. Metab. Disord. 28 (Suppl. 4) (2004) S29–S37. [132] H. Cao, et al., Identification of a lipokine, a lipid hormone linking adipose tissue to systemic metabolism, Cell 134 (6) (2008) 933–944. [133] M.M. Yore, et al., Discovery of a class of endogenous mammalian lipids with antidiabetic and anti-inflammatory effects, Cell 159 (2) (2014) 318–332. [134] L. Reshef, et al., Glyceroneogenesis and the triglyceride/fatty acid cycle, J. Biol. Chem. 278 (33) (2003) 30413–30416. [135] S.C. Kolwicz, R. Tian, Glucose metabolism and cardiac hypertrophy, Cardiovasc. Res. 90 (2) (2011) 194–201. [136] Y. Cheng, D. Hauton, Cold acclimation induces physiological cardiac hypertrophy and increases assimilation of triacylglycerol metabolism through lipoprotein lipase, Biochim. Biophys. Acta 1781 (10) (2008) 618–626. [137] G.F. Chen, Z. Sun, Effects of chronic cold exposure on the endothelin system, J. Appl. Physiol. 100 (5) (2006) 1719–1726. [138] J. Rawlins, A. Bhan, S. Sharma, Left ventricular hypertrophy in athletes, Eur. J. Echocardiogr. 10 (3) (2009) 350–356. [139] Y.H. Han, et al., Adipocyte-specific deletion of manganese superoxide dismutase protects from diet-induced obesity through increased mitochondrial uncoupling and biogenesis, Diabetes 65 (9) (2016) 2639–2651. [140] X.R. Peng, et al., Unlock the thermogenic potential of adipose tissue: pharmacological modulation and implications for treatment of diabetes and obesity, Front Endocrinol (Lausanne) 6 (2015) 174. [141] Q. Hao, et al., Transcriptome profiling of brown adipose tissue during cold exposure reveals extensive regulation of glucose metabolism, Am. J. Physiol. Endocrinol. Metab. 308 (5) (2015) E380–E392. [142] Z. Irshad, et al., Diacylglycerol acyltransferase 2 links glucose utilization to fatty acid oxidation in the brown adipocytes, J. Lipid Res. 58 (1) (2017) 15–30. [143] J.M. Olsen, et al., beta(3)-Adrenergically induced glucose uptake in brown adipose tissue is independent of UCP1 presence or activity: Mediation through the mTOR pathway, Mol. Metab. 6 (6) (2017) 611–619. [144] J.M. Olsen, et al., Glucose uptake in brown fat cells is dependent on mTOR complex 2-promoted GLUT1 translocation, J. Cell Biol. 207 (3) (2014) 365–374. [145] A.M. Hall, A.J. Smith, D.A. Bernlohr, Characterization of the Acyl-CoA synthetase

[146] [147]

[148]

[149] [150]

[151]

[152]

[153]

[154]

[155] [156]

[157]

[158]

[159]

[160]

[161]

[162]

[163]

[164] [165]

90

activity of purified murine fatty acid transport protein 1, J. Biol. Chem. 278 (44) (2003) 43008–43013. J.E. Schaffer, H.F. Lodish, Expression cloning and characterization of a novel adipocyte long chain fatty acid transport protein, Cell 79 (3) (1994) 427–436. A. Falcon, et al., FATP2 is a hepatic fatty acid transporter and peroxisomal very long-chain acyl-CoA synthetase, Am. J. Physiol. Endocrinol. Metab. 299 (3) (2010) E384–E393. P.N. Black, et al., Targeting the fatty acid transport proteins (FATP) to understand the mechanisms linking fatty acid transport to metabolism, Immunol., Endocr. Metab. Agents Med. Chem. 9 (1) (2009) 11–17. K. Milger, et al., Cellular uptake of fatty acids driven by the ER-localized acyl-CoA synthetase FATP4, J. Cell Sci. 119 (Pt 22) (2006) 4678–4688. H. Doege, et al., Targeted deletion of FATP5 reveals multiple functions in liver metabolism: alterations in hepatic lipid homeostasis, Gastroenterology 130 (4) (2006) 1245–1258. S. Yan, et al., Long-chain acyl-CoA synthetase in fatty acid metabolism involved in liver and other diseases: an update, World J. Gastroenterol. 21 (12) (2015) 3492–3498. M.J. Kang, et al., A novel arachidonate-preferring acyl-CoA synthetase is present in steroidogenic cells of the rat adrenal, ovary, and testis, Proc. Natl. Acad. Sci. U. S. A. 94 (7) (1997) 2880–2884. S.Y. Bu, D.G. Mashek, Hepatic long-chain acyl-CoA synthetase 5 mediates fatty acid channeling between anabolic and catabolic pathways, J. Lipid Res. 51 (11) (2010) 3270–3280. I. Boomgaarden, et al., Comparative analyses of disease risk genes belonging to the acyl-CoA synthetase medium-chain (ACSM) family in human liver and cell lines, Biochem. Genet. 47 (9–10) (2009) 739–748. C.M. Roche, et al., Physiological role of acyl coenzyme a synthetase homologs in lipid metabolism in Neurospora crassa, Eukaryot. Cell 12 (9) (2013) 1244–1257. Y. Ikeda, et al., Transcriptional regulation of the murine acetyl-CoA synthetase 1 gene through multiple clustered binding sites for sterol regulatory elementbinding proteins and a single neighboring site for Sp1, J. Biol. Chem. 276 (36) (2001) 34259–34269. G. Perez-Chacon, et al., Control of free arachidonic acid levels by phospholipases A2 and lysophospholipid acyltransferases, Biochim. Biophys. Acta 1791 (12) (2009) 1103–1113. M.C. Hunt, et al., Analysis of the mouse and human acyl-CoA thioesterase (ACOT) gene clusters shows that convergent, functional evolution results in a reduced number of human peroxisomal ACOTs, FASEB J. 20 (11) (2006) 1855–1864. K. Huhtinen, et al., The peroxisome proliferator-induced cytosolic type I acyl-CoA thioesterase (CTE-I) is a serine-histidine-aspartic acid alpha/beta hydrolase, J. Biol. Chem. 277 (5) (2002) 3424–3432. Y. Kuramochi, et al., Characterization of mouse homolog of brain acyl-CoA hydrolase: molecular cloning and neuronal localization, Brain Res. Mol. Brain Res. 98 (1–2) (2002) 81–92. J.M. Ellis, G.W. Wong, M.J. Wolfgang, Acyl coenzyme A thioesterase 7 regulates neuronal fatty acid metabolism to prevent neurotoxicity, Mol. Cell. Biol. 33 (9) (2013) 1869–1882. S. Han, D.E. Cohen, Functional characterization of thioesterase superfamily member 1/Acyl-CoA thioesterase 11: implications for metabolic regulation, J. Lipid Res. 53 (12) (2012) 2620–2631. R.L. Prass, F. Isohashi, M.F. Utter, Purification and characterization of an extramitochondrial acetyl coenzyme A hydrolase from rat liver, J. Biol. Chem. 255 (11) (1980) 5215–5223. Z. Cheng, et al., Crystal structure of human thioesterase superfamily member 2, Biochem. Biophys. Res. Commun. 349 (1) (2006) 172–177. H. Zhao, et al., The Akt C-terminal modulator protein is an acyl-CoA thioesterase of the hotdog-fold family, Biochemistry 48 (24) (2009) 5507–5509.