Fat in hearts: Uptake, storage, and turnover

Chad M Trent

Submitted in partial fulfillment of the requirements for the degree of Doctor of Philosophy under the Executive Committee of the Graduate School of Arts and Sciences

COLUMBIA UNIVERSITY 2014

© 2014

Chad M Trent

All rights reserved

Fat in hearts: Uptake, storage, and turnover

Chad M Trent

Abstract

The heart is one of the most energy demanding organs and is metabolically flexible to meet those demands. A major fraction of the hearts energy is derived from the uptake of circulating lipids; this dissertation reviews general pathways of lipid metabolism and then discusses how the heart obtains lipids and how these lipids are processed once inside the heart. Furthermore, derangements in lipid uptake and processing that are associated or causative of cardiac dysfunction are discussed. A new finding describes how the mouse heart requires lipoprotein lipase derived fatty acids for intracellular lipid droplets formation. Finally, the implications of these findings and future studies are described.

Table of Contents

List of Figures ...... iv

List of Tables ...... viii

List of Abbreviations ...... ix

Chapter 1: Introduction to lipid metabolism ...... 1

Overview of lipids ...... 1

Activation of lipids ...... 2

Esterification of lipids ...... 3

Fatty acid oxidation ...... 4

Fatty acid synthesis ...... 6

Dietary lipid absorption and circulation ...... 6

In vivo models of lipoprotein lipase function ...... 9

Characterization of the fatty acid transporter cluster of differentiation 36 ...... 11

Additional proteins involved in FFA transport ...... 13

Lipid droplets and associated proteins ...... 19

Lipid droplet synthesis and degradation ...... 25

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Intracellular lipolysis ...... 26

Transcriptional regulation of lipid metabolism ...... 27

Chapter 2: Lipid metabolism and toxicity in the heart ...... 31

Introduction ...... 31

Lipid uptake and turnover in the heart ...... 33

Creation of cardiac lipotoxicity in mice ...... 37

Lipid stores and the causes of toxicity ...... 38

Correction of lipotoxic heart disease ...... 40

Conclusions ...... 44

Chapter 3: Lipoprotein lipase activity is required for cardiac lipid droplet production ...... 45

Abstract ...... 45

Introduction ...... 45

Materials and Methods ...... 47

Results ...... 56

Fasted Ppara-/- mice do not store lipids in the heart ...... 56

Fasted Cd36-/- mice do not store lipids in the heart ...... 66

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Fasted hLpL0 mice do not store lipids in the heart ...... 75

Plin2 and Plin5 protein expression is not affected by cardiac lipid stores ...... 85

Blocking circulating TG degradation prevents cardiac lipid accumulation ...... 90

Cardiac TG accumulation during fasting does not impair heart function ...... 104

Discussion ...... 109

Chapter 4: Conclusions and future directions ...... 121

References ...... 125

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List of Figures

Figure 1 - Uptake, storage, and oxidation of fatty acids ...... 5

Figure 2 - Adrenergic regulation of intracellular lipolysis ...... 23

Figure 3 - PPARs: transcriptional control of lipid metabolism ...... 28

Figure 4 - Cartoon of cardiac lipid metabolism ...... 36

Figure 5 - Correcting cardiac lipotoxicity ...... 43

Figure 6 - Plasma biochemistry of fed and fasted Ppara+/+ and Ppara-/- mice ...... 56

Figure 7 - Lipid measurements from hearts of fed and fasted Ppara+/+ and Ppara-/- mice ...... 57

Figure 8 - ORO staining of hearts from fed and fasted Ppara+/+ and Ppara-/- mice ...... 58

Figure 9 - PAS staining of hearts from fed and fasted Ppara+/+ and Ppara-/- mice ...... 59

Figure 10 - Glycogen content of hearts from fed and fasted Ppara+/+ and Ppara-/- mice ...... 60

Figure 11 - Gene expression in hearts from fed and fasted Ppara+/+ and Ppara-/- mice ...... 62

Figure 12 - Supplemental gene expression in hearts from fed and fasted Ppara+/+ and Ppara-/- mice ...... 64

Figure 13 - TG lipase activity in hearts from fed and fasted Ppara+/+ and Ppara-/- mice ...... 65

Figure 14 - Plasma biochemistry of fed and fasted Cd36+/+ and Cd36-/- mice ...... 67

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Figure 15 - Lipid measurements from hearts of fed and fasted Cd36+/+ and Cd36-/- mice ...... 68

Figure 16 - ORO staining of hearts from fed and fasted Cd36+/+ and Cd36-/- mice ...... 69

Figure 17 - PAS staining of hearts from fed and fasted Cd36+/+ and Cd36-/- mice ...... 70

Figure 18 - Glycogen content of hearts from fed and fasted Cd36+/+ and Cd36-/- mice ...... 71

Figure 19 - Gene expression in hearts from fed and fasted Cd36+/+ and Cd36-/- mice ...... 72

Figure 20 - Supplemental gene expression in hearts from fed and fasted Cd36+/+ and Cd36-/- mice

...... 73

Figure 21 - TG lipase activity in hearts from fed and fasted Cd36+/+ and Cd36-/- mice ...... 74

Figure 22 - Plasma biochemistry of fed and fasted LpLflox/flox and hLpL0 mice ...... 76

Figure 23 - Lipid measurements from hearts of fed and fasted LpLflox/flox and hLpL0 mice ...... 77

Figure 24 - ORO staining of hearts from fed and fasted LpLflox/flox and hLpL0 mice ...... 78

Figure 25 - PAS staining of hearts from fed and fasted LpLflox/flox and hLpL0 mice ...... 79

Figure 26 - Glycogen content of hearts from fed and fasted LpLflox/flox and hLpL0 mice ...... 80

Figure 27 - Gene expression in hearts from fed and fasted LpLflox/flox and hLpL0 mice ...... 82

Figure 28 - Supplemental gene expression in hearts from fed and fasted LpLflox/flox and hLpL0 mice ...... 83

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Figure 29 - TG lipase activity in hearts from fed and fasted LpLflox/flox and hLpL0 mice ...... 84

Figure 30 - Western blot of PLIN2 and PLIN5 in hearts of fed and fasted Cd36+/+ and ...... 86

Figure 31 - Band density analysis of PLIN2 and PLIN5 western blots in hearts of fed and fasted

Cd36+/+ and Cd36-/- mice ...... 87

Figure 32 - Western blot of PLIN2 and PLIN5 in hearts of fed and fasted LpLflox/flox and hLpL0 mice ...... 88

Figure 33 - Band density analysis of PLIN2 and PLIN5 western blots in hearts of fed and fasted

LpLflox/flox and hLpL0 mice ...... 89

Figure 34 - Plasma biochemistry of fasted PBS and P407-treated mice ...... 91

Figure 35 - Lipid measurements from hearts of fasted PBS and P407-treated mice ...... 92

Figure 36 - ORO staining of hearts from fasted PBS and P407-treated mice ...... 93

Figure 37 - Plasma disappearance of [3H]-oleate in fasted PBS and P407-treated mice ...... 95

Figure 38 - Heart and liver uptake of FFA in fasted PBS and P407-treated mice ...... 96

Figure 39 - Plasma disappearance of [14C]-2-deoxyglucose in fasted PBS and P407-treated mice

...... 97

Figure 40 - Heart and liver uptake of glucose in fasted PBS and P407-treated mice ...... 98

Figure 41 - Glycogen content of hearts from fasted PBS and P407-treated mice ...... 99

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Figure 42 - PAS staining of hearts from fasted PBS and P407-treated mice ...... 99

Figure 43 - Gene expression in hearts from fasted PBS and P407-treated mice ...... 101

Figure 44 - Supplemental gene expression in hearts from fasted PBS and P407-treated mice .. 102

Figure 45 - TG lipase activity in hearts from fasted PBS and P407-treated mice ...... 103

Figure 46 - Basal and isoproterenol-stimulated fractional shortening in fasted Cd36+/+ and Cd36-/- mice ...... 105

Figure 47 - M-mode images of basal and isoproterenol-stimulated fractional shortening in fasted

Cd36+/+ and Cd36-/- mice ...... 106

Figure 48 - Basal and isoproterenol-stimulated fractional shortening in fasted LpLflox/flox and hLpL0 mice ...... 107

Figure 49 - M-mode images of basal and isoproterenol-stimulated fractional shortening in fasted

LpLflox/flox and hLpL0 mice ...... 108

Figure 50 - Distribution of plasma FFA in lipoprotein fractions ...... 113

Figure 51 - Pathways of Lipid Uptake Leading to Lipid Droplet Formation ...... 117

Figure 52 - Ceramide measurement in fed and fasted wild-type mice ...... 118

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List of Tables

Table 1 - Fatty acid transporters 13

Table 2 - Lipid droplet proteins 19

Table 3 - Correction of cardiac lipotoxicity 42

Table 4 - Primer sequences for "LpL activity is required for cardiac lipid droplet production" 55

Table 5 - Models of altered cardiac TG accumulation during starvation 124

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List of Abbreviations Abhydrolase domain containing 5, ABHD5 Fatty acid synthase, FASNFatty acid transport protein, FATP Acyl-CoA synthetase long-chain, ACSL Free fatty acid, FFA Acyl-CoA oxidase, ACOX Glycosylphosphatidylinositol anchored Adipose triglyceride lipase, ATGL high-density lipoprotein binding protein 1, Angiopoietin-like 4, ANGPTL4 GPI-HBP1

Apolipoprotein B, ApoB Hormone sensitive lipase, HSL

Apolipoprotein C, ApoC Lipoprotein lipase, LpL

Brown adipose tissue, BAT Medium chain acyl-CoA dehydrogenase, MCAD Carnitine palmitoyl transferase, CPT Microsomal triglyceride transfer protein, Chylomicron, CM MTTP

Cell death activator, CIDE Monoacylglycerol acyltransferase, MGAT

Cluster of differentiation 36, CD36 Perilipin, PLIN

Diacylglycerol acyltransferase, DGAT Peroxisome-proliferator activated receptor, PPAR Diacylglycerol, DAG

Triglyceride, TG Fatty acid, FA

Very low-density lipoprotein, VLDL Fatty acid binding protein, FABP

White adipose tissue, WAT

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Chapter 1: Introduction to lipid metabolism Overview of lipids

Lipids are a class of organic molecules that are defined as soluble in organic solvents and weakly soluble in water. These molecules can organize in large, supramolecular structures through noncovalent interactions to form membranes and droplets. Lipids perform three major roles in living cells. First, lipids are a major component of membranes, facilitating compartmentalization and separation. Lipids containing hydrocarbon chains can function as an efficient long-term energy storage form. Finally, lipids are important molecules for cellular signal transduction.

One of the major lipid classes is the fatty acid (FA). FAs are weak acids that are usually comprised of an even number of carbon atoms with one carboxyl group. Due to this carboxyl group, FAs are sequestered into lipids that are more complex by their sequential esterification

(1). The major energy storage molecule for eukaryotic organisms is triglyceride (TG), which is comprised of three FAs esterified to a glycerol backbone. In general, non-esterified FA (or free fatty acid, FFA) concentrations are kept at very low concentrations inside of cells by near constant incorporation into TGs or via stepwise oxidation to produce energy. Since FAs contain high numbers of single bonds, they are particularly suited to stepwise oxidation to produce an enormous amount of reducing equivalents for mitochondrial oxidative production of adenosine triphosphate (ATP), the energy currency of the cell.

Phospholipids and sphingolipids are formed from the esterification of FAs to other organic compounds. Phospholipids are comprised of two FAs esterified to glycerol, typically in the sn-1 and sn-2 positions, and a phosphate group esterified to the sn-3 position (1). There are a

1 number of hydrophobic head groups, including but not limited to choline, serine, and ethanolamine, which are esterified to the phosphate group. Phospholipids are amphipathic molecules and are the major lipid constituent of cell membranes. Furthermore, membrane phospholipids are hydrolyzed to release FAs, diacylglyceride (DAG), or their head group; these molecules can activate intracellular signaling pathways. Sphingolipids are similar molecules, comprised of except that they are consist of a FA amide-bonded to a serine (1). Ceramides are formed from the addition of an additional FA in an amide bond to a sphingolipid. These molecules are major cell membrane lipids and function as signaling molecules.

Another class of lipid molecules are the sterols, which are important as cell membrane constituents, for formation of bile salts, and in the generation of steroid hormones. Free cholesterol is esterified with a FA to form a cholesteryl ester; this is the major storage and circulating form of cholesterol.

Activation of lipids

After FFA enters a cell, it is thioesterified to a CoASH molecule by a class of enzymes called long-chain acyl-CoA synthetases (Acsl). The fatty acyl-CoA molecule can enter a number of pathways, including mitochondrial β-oxidation or esterification into TGs (2). There are 5 canonical Acsl enzymes, as well as 6 FA transport proteins that have Acsl activity, postulated to have differing roles in directing FAs to distinct cellular fates. The best characterized of these is

ACSL1, which is responsible for synthesis of acyl-CoA at the cell membrane and directing FFA to β-oxidation (3,4).

2

Esterification of lipids

Due to the relatively nonpolar hydrocarbon chain in FAs, lipids droplets are the most entropically stable form in an aqueous environment. Therefore, the majority of TGs in cells exist as lipid droplets. These droplets cannot easily traverse cell membranes and must be partially degraded to release FFAs that can enter or exit a cell. Almost all cell types possess the ability to esterify acyl-CoA into TG (5). The two tissues with the most TG esterification are the liver and the white adipose tissue (WAT). In the liver, glycerol kinase can phosphorylate free glycerol, which forms the backbone for sequential esterification of two fatty acyl-CoA substrates in the sn-

1 and sn-2 positions via the action of glycerol-3-phosphate acyltransferase and then acyl-glycerol phosphate acyltransferase to produce phosphatidic acid. Phosphatidic acid is converted to DAG via dephosphorylation of the phosphate group esterified to the sn-3 carbon by phosphatidic acid phosphatase. This process is much the same in white adipose tissue, except that glycerol-3- phosphate is produced from the NADH-dependent reduction of the glycolytic intermediate dihydroxyacetone phosphate.

Once DAG is synthesized, the final (and rate-limiting) step of TG synthesis is catalyzed by the enzyme diacylglycerol acyltransferase (DGAT). This enzyme esterifies a fatty acyl-CoA on the sn-3 position of diacylglycerol. There are two known DGAT isoforms – Dgat1 and Dgat2

– that are expressed in most tissues (5). Dgat1 was the first gene to be cloned and then knocked out (6,7); these knockout mice were resistant to weight gain and had reduced adipose tissue mass, but DGAT activity was still detected in some tissues. Shortly thereafter, a second Dgat was identified in a fungal species and then detected via sequence homology in mammalian tissues (8). When Dgat2 was knocked out in mice, these mice died shortly after birth due to skin barrier defects because of inability to synthesize TGs in the skin (9). These studies (and others)

3 illustrate both the redundant roles of DGAT1 and DGAT2, as well as distinct roles for each enzyme (10). Interestingly, Dgat1 also catalyzes a number of other reactions – it has the ability to make retinyl esters from retinol and acyl-CoAs (11).

Fatty acid oxidation

FAs are the most energy-rich substrates for ATP production in cells. As mentioned, FAs are activated by ACSLs to acyl-CoA molecules that can then enter multiple pathways. Short and medium chain acyl-CoA molecules can enter directly into the mitochondrial matrix for β- oxidation, but long chain (more than 10 carbons) must undergo a multi-step process to enter the mitochondrial matrix (12). The first, and rate-limiting, step is the exchange of the CoA cofactor for a carnitine molecule, by the enzyme carnitine-palmitoyltransferase 1 (CPT1). There are multiple CPT1 isozymes; CPT1A is liver specific, CPT1B is primarily expressed in heart and skeletal muscle, and CTP1C is expressed in the brain (13). The critical importance of these enzymes are emphasized by the phenotype of human loss-of-function mutations; impaired energy production from fat means these patients are prone to hypoglycemia and fatigue, but diets enriched in short- and medium chain FAs can ameliorate the deficiency (13).

Acylcarnitines enter the mitochondrial matrix via the carnitine-acylcarnitine translocase, where a second enzyme, CPT2, re-exchanges the carnitine for another CoA (13). The regeneration of the acyl-CoA inside the mitochondrial matrix is thus able to begin β-oxidation. In brief, acyl-CoA is oxidized by acyl-CoA dehydrogenases that catalyze the stepwise removal of acetyl-CoA and production of NADH and FADH2 (14). These reducing equivalents enter the electron transport chain to drive oxidative phosphorylation, and the acetyl-CoA enters the tricarboxylic acid cycle to generate more reducing equivalents.

4

Figure 1 - Uptake, storage, and oxidation of fatty acids

FFAs may enter a cell through a number of transporters or via flip-flop. Inside of the cell, FFAs are activated by the thioesterase ACSL, at which point they are directed to mitochondrial β- oxidation or sequential esterification. CPT1 directs fatty acyl-CoAs to β-oxidation, whereas monoacylglycerol acyltransferase (MGAT) and DGAT direct fatty acyl-CoAs towards esterification into TG.

5

Fatty acid synthesis

Just as FAs are oxidized to yield reducing equivalents and acetyl-CoA, the reverse process can also occur. Acetyl-CoA and malonyl-CoA can be ligated and reduced to form long- chain FAs via a multistep process (15). First, acetyl-CoA carboxylase (ACC) adds a CO2 to the acetyl-CoA to yield malonyl-CoA. This molecule is the substrate for FA synthase (FASN); through a series of NADPH-dependent reductions, two carbons derived from acetyl-CoA are added and the CO2 is removed, leading to an extension of two carbons at every step (16). FASN is a homodimeric protein that has two acyl carrier protein regions; at one site, the nascent FA is bound, and the other site binds the additional malonyl-CoA that added to the growing FA (17).

The other regions of FASN are responsible for enzymatic reductions and dehydrations that catalyze formation of the FA. Malonyl-CoA is also a potent allosteric inhibitor of CPT1, and it is thought that tissues that have low de novo lipogenesis use a second enzyme, ACC2, to produce malonyl-CoA in order to decrease the rate of β-oxidation (18). The major product of de novo lipogenesis is palmitate (C16); longer FAs may be produced by the elongation-of-very-long- chain-FAs (ELOVL) enzymes (19). Unsaturated and polyunsaturated FAs can be formed by a class of enzymes called the stearoyl-CoA desaturases (SCDs) (19).

Dietary lipid absorption and circulation

Dietary lipids primarily exist as TGs. The major site of dietary lipid absorption is in the small intestine. TGs are first emulsified in combination with bile salts secreted from the gall bladder and then degraded, primarily by pancreatic lipase, to form FFAs and monoglycerides

(20). These lipids form a mixed micelle, which adsorbs onto the surface of the enterocytes and then are taken up into the enterocyte – long and very-long chain FAs are transported primarily via the FA transporter cluster of differentiation 36 (CD36) (21). These FFAs and monoglycerides

6 are repackaged in a stepwise process to enter circulation. First, the FFAs and monoglycerides are re-esterified into TGs, primarily via the two enzymes diacylglycerol acyltransferase (DGAT) 1 and 2 that catalyze the final step in TG synthesis (10). Monoglycerides and FFAs are esterified to form DAG by the enzyme monoacylgycerol acyltransferase (MGAT) (22). Cholesterol, as well as retinol (vitamin A), are similarly hydrolyzed, absorbed, and re-esterified with FAs (23). These

TGs are loaded into a nascent lipoprotein via the protein microsomal TG transfer protein

(MTTP). Apolipoprotein B48 (ApoB48), a truncated splice-isoform of the full ApoB mRNA transcript, is the constituent apolipoprotein. These newly formed particles are termed chylomicrons (CM) and transport dietary lipids from the enterocyte into the lymph. Via the lymphatic circulation, CMs are transported into the vena cava to the heart and then pumped throughout the body.

Once the CMs enter the blood circulation, many tissues use them as a source of energy.

The major destinations for chylomicron lipids are the heart, skeletal muscle, and adipose tissue; however, other organs also take up some CM lipids. Approximately 50% of chylomicron TGs along with most of the ApoB48, retinyl esters, and cholesteryl esters are cleared from the circulation by the liver (24). The TG core of the chylomicron must be hydrolyzed into FFAs so that these FFAs can enter cells. Lipoprotein lipase (LpL) is the major enzyme responsible for the hydrolysis of circulating TGs; however, there are other identified lipases in the vascular endothelium such as hepatic lipase and endothelial lipase, but their substrates are not large TG- rich lipoproteins (25). LpL is secreted as a catalytically active dimer, primarily from muscle cells and adipocytes, and is bound to the endothelial cell surface via the glycosylphosphatidylinositol anchored high-density lipoprotein binding protein 1 (GPI-HBP1) anchored in the vascular endothelium and heparan-sulfate proteoglycans (26,27). Once FAs are liberated from the core

7

TG of chylomicrons, they can enter cells and be oxidized for ATP production or re-esterified and stored as TG. The liver can take up the TG-depleted CMs via receptor-mediated endocytosis; this is the major pathway for delivery of dietary cholesterol and fat-soluble vitamins to the liver (24).

LpL activity is regulated at multiple levels. At the most basic level, high FFA concentrations inhibit LpL activity through product inhibition in both in vivo and in vitro experiments (28-30). LpL activity is increased by the binding of the apolipoprotein C (ApoC)II, and inhibited by the binding of ApoCI and ApoCIII (25). Transgenic mice that overexpress human ApoCI or ApoCIII have hyperchylomicronemia due to attenuated LpL activity (31,32).

Although ApoCII deficiency also caused hypertriglyceridemia (33), overexpression of ApoCII also inhibited LpL activity by interfering with CM binding to heparin sulfate proteoglycans (34).

Fasting decreases white adipose tissue LpL activity via the local secretion of angiopoietin like 4

(AngPtl4), which inhibits LPL lipoprotein lipase by breaking apart the catalytic dimer (35). LpL transcription and translation are positively regulated by insulin stimulation in adipose tissue

(36,37); transcriptional regulation of LpL is discussed later.

Alternate pathways for TG uptake also exist; however, these seem to mostly function either in concert with or complementary to the activity of LpL. Two additional extracellular TG lipases, hepatic lipase and endothelial lipase, belong to the same gene family as LpL (38,39).

Hepatic lipase is secreted from hepatocytes and binds to the vascular epithelial cells of the liver; similar to LpL, it plays a role in both hydrolyzing circulating TG as well as anchoring lipoproteins and increasing endocytosis via the LDL receptor (40). This anchoring activity is most important for the uptake of CM remnants in the liver (41). Endothelial lipase is synthesized

8 and secreted from endothelial cells; although it has some TG lipase activity, there is much higher activity for the hydrolysis of lipoprotein phospholipids (39).

The VLDL receptor can facilitate TG uptake via endocytosis of -rich lipoproteins; in particular, it recognizes ApoE containing particles – excluding uptake of LDL (42). VLDLR is expressed in non-hepatic tissue and is most abundant in the brain. VLDLR deficient mice have normal circulating TG concentrations (43); later studies suggest VLDLR may be most important for anchoring circulating TG-rich lipoproteins and increasing LpL activity (42,44,45).

In vivo models of lipoprotein lipase function

Since the advent of knockout and transgenic mouse models, there have been a number of knockout and transgenic mice that have characterized LpL in vivo. The first LpL knockout mouse was described in 1995 by Levak-Frank et al. (46) and exhibited severe hypertriglyceridemia and died shortly after birth. Concurrently, a transgenic mouse that had muscle-specific overexpression of human LpL (MCK-LpL) was developed (46). The MCK-LpL mouse exhibited dramatically lower plasma TG and extensive mitochondrial proliferation in skeletal muscle, and this transgene was able to rescue the post-natal lethality in the LpL knockout mice.

Later, LpL was floxed and a cardiac specific knockout of LpL (hLpL0) was developed

(47). This mouse had reduced heart LpL activity and increased plasma TG. Furthermore, these mice exhibited increased heart glucose and FFA uptake, but a decreased expression of PPARα target genes involved in FA oxidation. These hLpL0 mice were further characterized to have increased glucose oxidation and decreased FA oxidation, and eventually developed cardiac dysfunction (47). Similar results were observed using an inducible, cardiac-specific knockout of

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LpL (48). These studies implicate the critical role of circulating TG in providing energy substrates for the heart, and is discussed in more detail in Chapters 2 and 3.

More recently, LpL has been knocked out in several other tissues. Skeletal muscle- specific LpL deletion led to increased muscle glucose uptake and increased insulin sensitivity, at the expense of decreased glucose uptake and insulin resistance in other tissues (49). LpL was thought to be a major factor in adipose tissue accretion of TG mass; surprisingly, despite 70-85% reductions in LpL activity the white-adipocyte specific LpL knockout mouse has relatively normal WAT mass, but decreased brown adipose tissue (BAT) size and function (50). Mass spectrometry analysis indicated an altered WAT FA profile with less abundance of dietary lipids

(linoleic acid), probably being compensated for by de novo lipogenesis; this is similar to observations in the adipose tissue of LpL-deficient humans (51).

Neuron-specific LpL knockout mice become obese on chow diets at 4 months of age, which was attributed to increased food intake and decreased energy expenditure (52). TG uptake in the hypothalamus was significantly reduced, which suggests that circulating lipids play a role in modulating central regulation of metabolism. Finally, macrophage-specific deletion of LpL had less severe development of atherosclerotic lesions, indicating a role for LpL in development of atherosclerosis (53). Interestingly, there was no difference in adipose tissue mass or adipocyte size in these mice, despite the abundance of adipose tissue macrophages. It is unclear exactly why there was less atherosclerosis in these mice; however, it was suggested that a secondary decrease in FA transporter CD36 expression or impaired macrophage energy availability might decrease macrophage infiltration.

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Similar to models of LpL deficiency, the GPI-HBP1 knockout mouse closely resembles the phenotype of the LpL-knockout mouse model (27). These mice have hyperchylomicronemia and impaired postprandial clearance of lipids; it was suggested that GPI-HBP1 is important for anchoring of chylomicrons. It was later demonstrated that this protein helps LpL to migrate from its origin in myocytes or adipocytes to the capillary endothelium (26).

Characterization of the fatty acid transporter cluster of differentiation 36

After the lipolysis of circulating TG and release of FFA, these FFA are taken up into tissues. Although at least some FFA can pass directly through the membrane via flip-flop and diffusion (54), a majority of FFA uptake is facilitated by transport proteins. The most essential and best-characterized FFA transporter is the scavenger receptor cluster of differentiation 36

(CD36) (55). Generation and study of Cd36-/- mice elucidated the primary roles of CD36 as a fatty acid transporter for the adipocyte, as well as confirmed earlier evidence that CD36 mediates macrophage uptake of oxidized LDL particles (56). Cd36-/- mice had elevated plasma FFA and

TG, as well as a decrease in circulating glucose. This decrease in circulating glucose is probably due to a metabolic compensation due to decreased availability of lipid substrates for energy production, as these mice were later shown to be more insulin and glucose sensitive when fed a high-glucose diet (57).

Later studies demonstrated the critical role of CD36 in uptake of FFA into heart and skeletal muscle; uptake of a synthetic FFA analog (15-(p-iodophenyl)-3-(R, S)-methyl pentadecanoic acid) was reduced about 50% (58). Notably, Cd36 deletion did not impair FFA uptake into the liver, suggesting that other FA transporters may be more important. However, chronic high-fat diet feeding increased CD36 in the liver, and adenoviral hepatic overexpression

11 of Cd36 exacerbated fatty liver. Furthermore, Cd36-/- mice were protected from high- carbohydrate and alcohol diet feeding induced hepatic steatosis (59). Diet-induced obese mice had increased liver CD36 protein expression, and adenoviral-mediated liver Cd36 overexpression further increased liver FFA uptake and TG content, suggesting that CD36 will transport FFA into hepatocytes during states of caloric excess (60). These data suggest that

CD36 is an important pathologic liver FA transporter in the context of chronic caloric excess and hepatic dysfunction.

A number of studies have elucidated the intimate relationship between LpL and Cd36 in the absorption of circulating TGs – both chylomicron and VLDL TG. Cd36 deficient humans have increased postprandial hypertriglyceridemia (61); knockout mice have a similar phenotype attributed to decreased LpL activity secondary to Cd36 deficiency (29). An in vitro LpL activity assay incubated with plasma from Cd36-/- mice demonstrated that product inhibition was probably responsible for the decreased LpL activity in vivo. It was later demonstrated that, when crossed onto a heart specific LpL knockout, the heart had decreased uptake of VLDL TG, but not

CM TGs (30). These data suggest that CD36-mediated FFA uptake is saturable and very high local FFA concentrations rely on other transporters, diffusion, or flip/flop.

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Additional proteins involved in FFA transport

Table 1 - Fatty acid transporters

Gene Aliases name CD36 Fatty acid translocase (FAT) Solute carrier family 27, member 1 (SLC27A1), Acyl-CoA synthetase very long chain family FATP1 member 4 (ASCVL4) Solute carrier family 27, member 2 (SLC27A2), Acyl-CoA synthetase very long chain family FATP2 member 1 (ASCVL1) Solute carrier family 27, member 3 (SLC27A3), Acyl-CoA synthetase very long chain family FATP3 member 3 (ASCVL3) Solute carrier family 27, member 4 (SLC27A4), Acyl-CoA synthetase very long chain family FATP4 member 5 (ASCVL5) Solute carrier family 27, member 5 (SLC27A5), Acyl-CoA synthetase very long chain family FATP5 member 6 (ASCVL6) Solute carrier family 27, member 6 (SLC27A6), Acyl-CoA synthetase very long chain family FATP6 member 6 (ASCVL2) FABP1 Liver fatty acid-binding protein (LFABP) FABP2 Intestinal fatty acid-binding protein (IFABP) FABP3 Muscle and heart fatty acid binding protein FABP4 Adipocyte fatty acid-binding protein (AFABP), Adipocyte protein 2 (AP2) FABP5 Epidermal fatty acid-binding protein (EFABP) FABP6 Ileal fatty acid-binding protein (ILBP) FABP7 Brain fatty acid-binding protein (FABPB) GOT2 Fatty acid binding protein, plasma membrane (FABPpm)

Other than CD36, there are additional proteins that facilitate uptake of circulating FFA

(Table 1). The FA transport protein (Fatp) proteins belong to a gene family that can complement or replace CD36 in certain tissues. Fatp1 was first identified in a library screening strategy and proposed as a FA transporter (62); expression profiling revealed the highest expression in muscle and adipose tissue (63). It was later shown that FATP1-mediated FA transport was insulin sensitive, and that insulin stimulation promoted the translocation of FATP1 from intracellular

13 membranes to the cell surface (64). This effect was abolished in FATP1 knockout mice (65).

FATP1 has both FA transport abilities as well as ACSL activity; cell culture studies indicate a particular role for directing FAs to TG synthesis (66). Fatp1 deficient mice were protected from diet-induced obesity induced skeletal muscle insulin resistance, which suggests a causative role for lipid uptake in skeletal muscle in the development of insulin resistance (65). However, it is also likely that skeletal muscle that has defective FFA uptake may compensate by increasing uptake of glucose. Fatp1 is also essential for BAT thermogenesis; it was suggested that this was a result of decreased FFA uptake in BAT, but it could also be due to decreased ACSL activity

(67).

Five additional Fatp family member, Fatp2 through Fatp6, were later identified using genomic sequence analysis based on sequence homology (68). Fatp2 was identified as being primarily expressed in the liver and localized to the peroxisome; although this enzyme was particularly important for peroxisomal transport of very long (greater than C22) chain FA, as well as having ACSL activity, knockout mice that had reduced peroxisomal β-oxidation did not have any accumulation of VLCFAs (69). Later studies indicated the FATP2 is primarily localized to the plasma membrane and critical for liver uptake of LCFAs – knockout mice had

40% reduced liver FFA uptake and 50% reduced peroxisomal oxidation (70).

Fatp3 is primarily expressed in testis, ovary, and adrenal glands and exhibits both FA transport activity as well as ACSL activity (71); it is primarily localized to the mitochondria.

RNA interference experiments in cultured cells implicated FATP3 primarily as an ACSL enzyme rather than a FA transporter. Fatp4 was identified as being a major intestinal FA transporter (72); however later investigation suggested that the FATP4 was primarily an ACSL enzyme;

14 overexpression of a mitochondrial-specific ACSL was also able to increase FA uptake (73).

Human genetics studies implicated defective FATP4 activity in development of ichthyosis due to the reduced ability to esterify VLCFA into phospholipids, cholesteryl esters, and TGs (74).

Fatp3 and Fatp4 may be particularly important for transcellular FFA uptake, as a very high level of mRNA expression of both genes was found in endothelial cells of the heart, skeletal muscle, and BAT (75).

The fifth family member Fatp5 is most abundantly expressed in the liver. FATP5 exhibits

ACSL activity for VLCFAs (C18, C20, C24, C26) (76), and also has thioesterification activity for cholate, suggesting that it may play a role in hepatic bile synthesis (77). Fatp5-/- mice were generated and demonstrated lower FFA uptake in isolated hepatocytes, as well as lower liver TG and FFA concentrations (78). In concordance with the importance for bile acid production,

Fatp5-/- mice have altered bile acid composition due to decreased choline conjugation to taurine and glycine (79). Since CD36 seems to have a negligible role in physiologic liver FFA uptake

(58,80) – and actually have increased hepatic TG deposition (81) —, it is likely that FATP2 and

FATP5 are the primary physiologic FFA transporters in the liver.

Finally, Fatp6 is most abundantly expressed in the heart and is localized to the cardiomyocyte sarcolemmal membrane, colocalizing with CD36 (82). In vitro experiments demonstrated that Fatp6 could transport FFA. Sequence analysis suggests that Fatp6 also has

ACSL activity, but there is no direct evidence (83). There have been no reports of Fatp6 knockout or overexpression models that would further elucidate the role of Fatp6 in cardiac lipid metabolism.

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Another class of proteins involved in FFA transport are the intracellular FA binding proteins (FABPs). These proteins bind hydrophobic molecules inside of cells, such as FFA and acyl-CoA. The first FABP to be cloned was most highly expressed in the liver, and was initially called the liver FABP (FABP1) (84). Crystallization reveals that FABP1 forms a β-barrel structure with a hydrophobic core that can reversibly bind palmitate (85). Cell culture studies as well as human loss-of-function mutants implicate FABP1 in modulating the uptake of circulating lipids (86,87). Fabp1-/- mice were somewhat protected from diet-induced obesity and nearly completely protected from hepatic lipid accumulation, but increased systemic glucose catabolism

(88). Forty-eight hour fasted Fabp1-/- mice had much lower hepatic FFA uptake, but relatively normal fasting-induced gene expression (89). FABP1 can localize to cell nuclei as well as bind to the transcription factors PPARα and PPARγ, suggesting that it may play a role in ligand binding to hormone receptors (90). Taken together, these studies strongly implicate FABP1 as an intracellular protein that sense hepatic lipid contents to modulate uptake.

Sequence analysis of the Fabp1 cDNA was used to discover homologous mRNAs and to identify an intestinal FABP (FABP2) (91). Human genetics studies implicated gain-of-function mutations in FABP2 as a predictor for increased plasma TG as well as cardiovascular and metabolic diseases (92,93). Intestinal FABP was thought to facilitate dietary lipid absorption and

CM-TG secretion. Female Fabp2-/- mice weighed less on both chow and high-fat diet feeding, whereas chow-fed male Fabp2-/- mice weighed more and were only slightly protected from diet- induced obesity (94). It is unclear why this occurred, but FABP1 and FABP2 probably have distinct roles in transport of their ligands towards oxidation versus storage (95). It is clear that

FABP2 is not essential for dietary lipid absorption; this mainly demonstrates the multiple redundancies in pathways of lipid absorption.

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Fabp3 is most highly expressed in the heart, particularly after birth reflecting the increased reliance on the postnatal heart on lipid metabolism (96). Fabp3 is also expressed in mammary tissue, but Fabp3-/- mice have no overt mammary phenotype (97). The most striking phenotype of Fabp3-/- mice was a 90% reduction of cardiac FA uptake and decreased FA oxidation, compensated for by increased glucose uptake and oxidation. These mice later developed cardiomyopathy (98,99). FABP3 is also expressed in the skeletal muscle, but knockout mice had a minimal reduction in muscle FFA uptake (100). Interestingly, brain lipids were also reduced – with long chain FFAs and total phospholipids reduced around 20% (101).

The fourth family member was initially characterized as adipocyte protein 2 (aP2, now known as Fabp4) due to high expression in white adipocytes and macrophages (102,103).

Interestingly, Fabp4-/- mice gained more weight than wild-type mice on a high fat diet, but were protected from insulin resistance and adipose tissue inflammation (104). FABP4 overexpression in macrophages alleviated lipid induced ER stress (103). Human genetic studies also implicates loss-of-function mutations in FABP4 with protection from type 2 diabetes and cardiovascular disease (105). These studies suggest that FABP4 reduces lipid toxicity, perhaps by directing excess FAs to neutral lipid synthesis.

The epithelial FABP (Fabp5) is highly expressed in differentiated keratinocytes, with lower levels of expression in heart and white adipocytes (106,107). Since many other lipid abnormalities result in dramatic skin phenotypes, it was surprising when FABP5 deficient mice were superficially normal and had only minor disruption of skin barrier function (108). Fabp6 is expressed mainly in the ileum of the small intestine (109). Knockout mice suggest that FABP6 is primarily important for the ileal absorption of bile acids from the lumen of the small intestine

17 into the portal circulation (110). Fabp7 is most highly expressed in neurons (111). Newborn

Fabp7-/- mice had decreased docosahexaenoic acid in brains; older mice had increased arachidonic acid and palmitic acid in adult brains associated with anxious behavior (112). QTL analysis found an association with anxiety and schizophrenic behaviors in mice (113). Knockout of both Fabp7 and Fabp5 has implicated these genes in neural differentiation and brain development; it is unclear if this is related to their role in directing lipids to oxidation, phospholipid synthesis, or gene transcription (114,115).

The first FA binding protein and transporter identified, known as the plasma membrane

FABP, is better known as glutamate oxaloacetate transferase (GOT2) due to having aminotransferase activity (116,117). This protein localizes to the plasma membrane, particularly in hepatocytes that have long-term alcohol exposure (118). Studies in cultured skeletal myocytes have implicated GOT2 as directing FFA to TG esterification (119,120). GOT2 can localize to both the plasma membrane and the mitochondria; however, studies in skeletal muscle of exercised rats suggest that it primarily transports FFA into the cell rather than into mitochondria

(121).

The identification of FA transporters and intracellular FA binding proteins has linked uptake of circulating lipids with specific intracellular fates, whether that is β-oxidation, esterification into TG, or transcriptional activation. Existing literature primarily focuses on the distinct roles of these proteins, and further investigation will likely elucidate how these proteins work in concert.

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Lipid droplets and associated proteins

The classical lipid droplet protein categorization has been according to the so-called PAT protein

– perilipin, adipophilin, and TIP47. Recently, a uniform naming convention has been adopted

(122). The most recent nomenclature and common aliases are listed in Table 2.

Table 2 - Lipid droplet proteins

Gene Aliases

PLIN1 Perilipin, Lipid droplet associated protein, Perilipin A

PLIN2 Adipophilin (ADFP), Adipose differentiation-related protein (ADRP)

PLIN3 Cargo Selection protein TIP47, Mannose-6-phosphate receptor-binding protein 1

PLIN4 Adipocyte protein S3-12

PLIN5 Lipid droplet-associated protein PAT-1, Lipid storage droplet protein 5 (Lsdp5), Myocardial Lipid Droplet Protein (MLDP), Oxidative PAT protein (OXPAT)

ABHD5 1-acylglycerol-3-phosphate O-acyltransferase, Abhydrolase domain-containing protein 5, Lipid droplet binding protein CGI-58

CIDEA Cell death-inducing DFFA-like effector A

CIDEB Cell death-inducing DFFA-like effector B

CIDEC Cell death activator CIDE-C, fat-specific protein 27 (FSP27)

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Adipocytes in WAT usually have a single large lipid droplet. As described previously, PLIN1 is the primary lipid droplet coat protein in WAT (123). When Plin1 was knocked out in mice, these animals consumed more food but were protected from weight gain; furthermore, knockout protected against obesity when crossed with the leptin-receptor deficient obese mice (124,125).

These studies also implicated Plin1 as a regulator of TG degradation via hormone-sensitive lipase (HSL) – both as protecting the lipid droplet from lipolysis, but also in the activation of lipase activity. At the time of these studies, it was believed that HSL was the most important gene for TG lipolysis; later studies demonstrated that knockout of Hsl only partially reduced TG lipase activity and did not result in TG accretion in WAT. Therefore, Plin1 is implicated in both regulating HSL and the other major TG lipase – the subsequently described adipose triglyceride lipase (ATGL) (126,127).

The second perilipin family member, Plin2, has been most extensively characterized in the liver. The most dramatic phenotype of Plin2-deficient mice was reduced hepatic TG content as well as protection from diet-induced fatty liver (128). These observations were attributed to a defect in lipid droplet biogenesis and a possible microsomal accumulation of TG. When crossed with obese leptin-deficient mice, these mice were somewhat protected from hepatic TG accumulation, about a 20% decrease (129). More dramatically, these obese mice lacking Plin2 had improved insulin and glucose sensitivity. This was coupled with increased VLDL secretion.

Later it was shown that Plin2 knockout mice on wild-type background were protected from diet- induced obesity, probably due to redistribution and increased mitochondrial proliferation and thermogenesis in white adipocytes (130). These data suggest that Plin2 may play a role in extrahepatic tissues; these functions have yet to be described.

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There have been no published Plin3 knockout mice; however, RNA interference targeting of Plin3 led to a reduction of hepatic TG contents and VLDL secretion (131). It is unclear why this occurred, and further studies are required to elucidate the molecular mechanism of Plin3 regulation of lipid droplet formation, structure, and turnover.

Plin4 is involved in lipid droplet biogenesis in white adipocytes. This protein is localized on the surface of lipid droplets, along with PLIN1 and ABHD5 (132). Although the highest levels of Plin4 expression are in white adipocytes, knockout animals displayed no overt adipose tissue phenotype, normal fat mass accretion, and no change in WAT lipolytic activity (133).

However, these mice had an interesting cardiac phenotype – failure to accumulate TG lipid droplets after an overnight fast – probably due to decreased mRNA and protein of Plin5. It is unclear why Plin5 expression decreased in the hearts of these Plin4-/- mice.

Plin5 is referred to as the oxidative lipid droplet protein, due to its high expression level in oxidative tissues like the heart, skeletal muscle, and BAT (134). PLIN5 is similar to PLIN1 in that it coats lipid droplets and translocates into the cytosol upon adrenergic stimulation (135).

Unlike PLIN1, it also binds directly to both ATGL and the co-activator, ABHD5. As expected,

Plin5 knockout mice had decreased TG in many oxidative tissues, and most notably failed to store lipid droplets during fasting (136). These hearts had increased lipid oxidation and eventually developed cardiac dysfunction as measured by fractional shortening. Two recent studies demonstrated cardiac steatosis in mice that specifically overexpress Plin5 in cardiac myocytes, most likely due to decreased intracellular TG lipolysis (137,138).

Concurrent with the description of the Atgl knockout mouse, the co-activator abhydrolase domain containing 5 (Abhd5) was characterized (139). This protein is responsible for increasing

21 activity of ATGL (140). Loss of function mutations in this gene are associated with the neutral lipid storage disorder Chanarin-Dorfman syndrome, typified by TG accretion in many organs. In adipocytes, ABHD5 bound to PLIN1 moves off the lipid droplet upon adrenergic stimulation, which facilitates interaction with ATGL to stimulate increased lipolysis (Figure 2) (141).

Knockout of this gene in mice resulted in early postnatal lethality and exhibited ichthyosis due to failure to produce skin wax lipids (142). More recently, a muscle-specific knockout of Abhd5 resulted in cardiac lipid accumulation, probably due to a decrease in lipolytic activity of ATGL

(143).

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Figure 2 - Adrenergic regulation of intracellular lipolysis

Upon binding of adrenaline to the β–adrenergic receptor, adenylate cyclase (AC) is activated.

This leads to the production of cyclic AMP (cAMP), which activates protein kinase A (PKA).

PKA phosphorylates PLIN1, which leads to the release of the ATGL activator ABHD5. PKA also phosphorylates HSL. In concert, this leads to the net release of FFA.

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In summary, lipid droplet proteins serve two primary functions – lipid droplet structure and regulation of lipase activity. Plin1 and Plin5 seem important for both of these roles, whereas

Abhd5 is primarily important for regulation of lipase activity. Plin2 seems primarily important for lipid droplet structure and probably has some role in VLDL assembly and secretion. The exact functional roles of Plin3 and Plin4 are unclear.

There are a number of other lipid droplet proteins, the cell death activator (CIDE) proteins – CIDEA, CIDEB, and CIDEC. Cidea is most highly expressed in the BAT, but there is detectable expression in the heart. Cidea knockout mice are lean and protected from diet-induced obesity; the authors stated that it was a negative regulator of BAT thermogenesis by suppressing

UCP1 activity (144). Cideb deficiency resulted in ubiquitously increased lipid oxidation resulting in lower plasma TG and FFA, as well as low TG accumulation in other tissues and protection from diet induced obesity (145). There is no clear mechanism why Cideb deletion causes this phenotype; however, VLDL secretion was impaired during fasting in Cideb knockout mice

(146,147). Cidec, better known as fat-specific protein 27 (Fsp27), is involved in promoting the accumulation of unilocular lipid droplets in white adipocytes; knockout mice had impaired white adipose tissue growth and exhibited multilocular lipid droplets and mitochondrial proliferation in white adipocytes (148). Furthermore, these mice were protected from diet-induced obesity and had normal glucose disposal and insulin tolerance. Later it was shown that CIDEC mediates the transfer of lipids from smaller to larger lipid droplets, which most likely explains the phenotype in the knockout mice (149,150).

Most of the understanding of lipid droplet proteins comes from cultured cells or knockout animals. Future studies are critical to evaluate the role of these proteins and their interactions.

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Most in vivo characterizations of these proteins rely on the overexpression and knockouts; thus it is critical to use innovative techniques to characterize their complex interplay.

Lipid droplet synthesis and degradation

Understanding intracellular lipid droplet accumulation and turnover in both liver and

WAT is essential to understand how these organs regulate circulating lipids. The liver synthesizes FA and secretes TG and cholesteryl esters in TG-rich lipoproteins. The WAT has some ability to synthesize FAs, but primarily functions as a TG storage and FFA secretion organ.

The liver is central to metabolism of all macronutrient classes. Excess dietary carbohydrates can be converted to acetyl-CoA and serve as a building block for the synthesis of new FAs. Furthermore, the liver has a tremendous capacity to take up circulating albumin-bound

FFAs (151). The hepatic pool of FFAs is rapidly esterified into TGs. The liver can store some of these TGs, but the main role of the liver is to secrete apolipoprotein B-100 very-low density lipoproteins (VLDL). The assembly process of VLDL particles is similar to the assembly process of chylomicrons, where MTTP mediates the loading of TG molecules into a nascent apoB-100 containing lipoprotein. These VLDL particles contain TGs as well as cholesteryl esters; in the post-prandial state these particles contain about 80% of circulating FA. Circulating VLDL-TG is hydrolyzed by LpL to release FAs for tissue uptake.

The primary role of the WAT is to store excess calories as TG. The WAT is a major depot of CM lipolysis via LpL, leading to storage of dietary lipids. WAT LpL activity is increased in the fed state to facilitate the storage of excess dietary lipids (152,153). On the other hand, fasting decreases WAT lipoprotein lipase activity via the local secretion of AngPtl4, which inhibits LpL by breaking apart the catalytic dimer (35). Decreasing WAT lipoprotein lipase

25 activity during fasting is probably to facilitate a net flux of circulating TGs to tissues like the heart and skeletal muscle that need to maintain constant energy production, and to avoid futile cycling of lipids in and out of WAT. The critical role for LpL in the clearance of plasma TG is evident in humans who have genetic deficiencies that lead to familial hyperchylomicronemia

(154,155).

Intracellular lipolysis

The CM- and VLDL-derived FFAs that are taken up in white adipocytes are then re- esterified and are stored in a single large lipid droplet. Lipid droplet proteins and intracellular lipases control turnover of the lipid droplet. The primary lipid droplet protein in white adipocytes is perilipin (Plin) 1. This protein is responsible for both maintaining the structure of the lipid droplet and for regulating the activity of intracellular lipases (156). The recently described rate- limiting intracellular lipase is ATGL, which primarily catalyzes the release of a single FFA from a TG molecule (127,139,140). The second enzyme in the lipolytic cascade is hormone-sensitive lipase (HSL), which yields a second FFA and a monoglyceride molecule. It should be noted that the enzyme activity termed “hormone-sensitive lipase” has been used in classic literature and in textbooks to represent the total intracellular lipolytic activity, and not just the activity of the enzyme HSL.

For decades, intracellular lipolytic activity was attributed to the activity of HSL. HSL was cloned and identified to be stimulated by catecholamines and repressed by insulin, primarily through phosphorylation and dephosphorylation (157,158). However, when the HSL gene was knocked out in mice, there was only a partial decrease in adipose tissue lipolysis (126). It had been predicted that a failure to lipolyze intracellular TG would result in obesity; it did not. This

26 suggested the existence of another rate-limiting lipolytic enzyme. This other enzyme, adipose TG lipase (ATGL) was first identified from adipose tissue extracts as having TG lipase activity

(159). This enzyme was knocked out in mice that then displayed the expected obesity phenotype, as well as ectopic lipid accumulation (127). In particular, these mice had increased cardiac TG accumulation associated with hypertrophy and heart dysfunction, leading to early mortality.

The activity of ATGL and HSL enzymes in WAT are increased via β-adrenergic stimulation. Upon binding of catecholamines to the β-2 and β-3 adrenergic receptors in white adipocytes, a G-protein coupled receptor signaling cascade leads to the phosphorylation of

PLIN1 by protein kinase A. The phosphorylation of PLIN1 leads to its release from the lipid droplet, allowing the lipolytic enzymes to access the TG (160,161). The release of PLIN1 also causes the release of a second protein, ABHD5, which can then co-activate ATGL (139). HSL is also phosphorylated by protein kinase A which activates its enzymatic activity (162). In concert, this leads to an approximate two-three fold increase in intracellular lipolysis, which has the net effect of increasing plasma FFAs (139). Conversely, insulin stimulation inhibits WAT lipolysis via the activation of phosphodiesterases, which inhibit the adrenergic signaling cascade

(163,164).

Transcriptional regulation of lipid metabolism

The peroxisome proliferator-activated receptor (PPAR) family of proteins is of particular importance for transcriptional regulation of lipid metabolism. The PPARs have overlapping and distinct roles in transcriptional regulation of lipid metabolism (Figure 3). These are nuclear steroid hormone receptors that heterodimerize with the retinoid X receptor (RXR) to activate or repress transcription of genes, primarily those involved in lipid metabolism.

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Figure 3 - PPARs: transcriptional control of lipid metabolism

The first family member, PPARα, is primarily thought to drive transcription of genes involved in FA oxidation, such as Cpt1, acyl-CoA oxidase (Acox), and medium chain acyl-CoA dehydrogenase (Mcad). Not surprisingly, Ppara-/- mice have decreased FA oxidation (165).

PPARα is particularly important in the transition from the postprandial to the fasted state, as it can increase the extrahepatic utilization of FAs and spare glucose utilization (166,167). Fibrate drugs activate PPARα and have the net effect of lowering plasma lipids; this is probably due to both increasing lipid oxidation as well as increasing lipid uptake by increasing LpL and Cd36

(168). Furthermore, PPARα increased transcription of Plin2 and Plin5 in the heart and liver; this effect was abolished in PPARα knockout mice (169). The Atgl knockout mice, which have myocardial lipid accumulation associated with cardiac dysfunction and death, were treated and rescued by treatment with the PPARα agonist WY14,643 (170) suggesting that lipid droplet mobilization may be important for activation of PPARα.

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Next, PPARδ activates transcription of a similar gene program to stimulate FA uptake and oxidation. While PPARα is thought to act primarily in the heart and liver, PPARδ is most important in the skeletal muscle (171) – knockout mice had reduced skeletal muscle β-oxidation, and stimulation with a PPARδ agonist increased skeletal muscle β-oxidation in two independent studies (172,173). Furthermore, treatment with a PPARδ agonist protected against weight gain in leptin-receptor deficient obese mice. It was later shown that overexpression of constitutively active PPARδ specifically in skeletal muscle dramatically boosted not only muscle oxidation, but dramatically improved exercise performance due to increased mitochondrial biogenesis and fiber type switching (174).

PPARγ is perhaps best known as a transcription factor for stimulating adipogenesis (175) and is the target of thiazolidinediones (168). This member is most highly expressed in white adipocytes and drives expression of genes involved in the uptake, storage, and turnover of lipids, such as LpL, Cd36, Plin1, and Atgl (176,177). PPARγ knockout mice are embryonic lethal due to failure to correctly develop placenta; however, having a single null allele for PPARγ prevented diet induced obesity, at least partially due to impaired adipocyte proliferation and growth (178).

In summary, the PPAR family of transcription factors seem to have distinct and overlapping roles in determining the expression of lipid metabolism genes

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Figure 3). Each seems to be particularly important in certain organs, but may have important roles in particular conditions of stress. Many of the cardiac specific roles of PPARs are discussed in Chapter 2; in particular, how they may contribute to the development or amelioration of toxic lipid accumulation and heart dysfunction.

The sterol regulatory element binding proteins (SREBPs) are another family of transcription factors that regulate lipid metabolism. SREBPs are endoplasmic reticulum membrane bound proteins that sense cellular lipid balance; when lipids are low, SREBP is cleaved by the SREBP cleavage activating protein (SCAP), at which point it can translocate to the nucleus to bind DNA and activate transcription of lipogenic genes (179). There are two distinct Srebp genes; splice-isoforms SREBP1a and SREBP1c primarily regulate transcription of genes involved in de novo lipogenesis, such as Acc1¸ Fasn, and Dgat1. SREBP2 drives transcription of genes to increase cellular cholesterol concentrations by activating transcription of genes such as HMG-CoA reductase (the rate-limiting enzyme for cholesterol synthesis) and the LDL receptor. Insulin signaling is believed to activate de novo lipogenesis in the liver primarily by activating SREBP1 (180,181). Although SREBP1 and 2 have been best characterized in how they regulate lipid metabolism in the liver, future studies may uncover important roles in other tissues.

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Chapter 2: Lipid metabolism and toxicity in the heart

This chapter has been published as a review in Cell Metabolism (182).

Goldberg, I.J., Trent, C.M., Schulze, P.C.

Introduction

Although the heart is by far the most energy-requiring organ of the body, studies of cardiac lipid metabolism, especially in vivo, are relatively scarce compared with investigations in adipose tissue or liver. In adult fasting mammals, 60-80% of cardiac energy metabolism relies on the oxidation of FAs with glucose, lactate, and ketones providing substrates for the remainder

(183). The adult heart, however, has the ability to switch to different substrates for ATP generation depending on feeding, hormonal status and overall nutritional supply as characterized by the Randle cycle (14). Of note, there are major species differences with mice relying more on glucose, lactate and ketone bodies, and much less on FAs (30-40% from fat) (184,185). The fetal heart operates under low oxygen pressure and primarily depends on glucose and lactate for ATP generation, whereas the adult heart utilizes FAs but conserves the ability to switch other substrates. Older animals and humans use relatively less FAs and more glucose.

The heart avidly acquires lipids both from circulating FFAs and esterified FAs bound to lipoproteins (Figure 4). Observations made studying arterial venous differences in substrate concentrations showed that esterified FAs were a major source of lipids for the human heart

(186). More recent methods to study heart lipid metabolism have relied on tracers of FFAs in isolated perfused hearts. These studies quantify conversion of FFAs to CO2 and TCA cycle intermediates under a variety of experimental conditions. In vivo studies can assess the uptake and loss of tracers from the heart. Although the heart can synthesize lipoproteins as it expresses

31 both ApoB and microsomal TG transfer protein (187,188), under most circumstances, the heart probably does not re-secrete appreciable amounts of glucose or lipids, and the uptake should indicate oxidation plus a relatively small amount of substrate that is stored and a small amount of substrate used for structural requirements of the cell.

In some situations, the heart adjusts to maintain lipid homeostasis. Increases in work load

(189) and myocardial ischemia (190) cause a rapid switch from fat to glucose utilization for ATP generation. This finding has led to several animal studies showing that administration of compounds that reduce FA oxidation protect the heart from the consequences of ischemia and ischemia-reperfusion injury (189,190). This is presumed to be due to reduced oxygen requirements for non-FA substrates. Deleterious effects of cardiac ischemia could be due in part to excess cardiac lipid accumulation via the VLDL receptor (45). Similarly, in another mouse model of cardiomyocyte death adiponectin-induced activation of a ceramidase and reduction of ceramide was beneficial (191). Therefore, abnormal regulation of lipid uptake or its intracellular metabolism might play an important role in heart diseases other than metabolic dilated cardiomyopathy.

An imbalance between FA uptake and oxidation leads to accumulation of long chain FAs that are incorporated into TG and phospholipids, as well as a multitude of other lipid subspecies.

Although TG is the most easily detected, other lipids are more likely to be toxic. DAGs and ceramides are signaling lipids that are thought to be toxic when their intracellular concentrations are increased. Defective mitochondrial FA oxidation could lead to accumulation of medium chain acyl carnitines (192), another possible toxin. Finally, saturated long chain FAs, most

32 notably palmitate, are associated with toxicity in cells either because of their direct actions or because of their incorporation into phospholipids (193).

Lipid uptake and turnover in the heart

All tissues obtain lipids from FFAs associated with albumin, lipoproteins, and de novo synthesis (Figure 4A). Although de novo synthesis is thought to play a minor role in heart lipid metabolism, a recent study of deletion of FA synthase in heart showed that de novo synthesis is important to maintain cardiac function during aortic constriction and aging (194). Loss of lipoprotein lipase (LpL)-derived lipids leads to increased glucose uptake in mouse hearts (195).

In humans, deficiency in CD36 is associated with increased glucose uptake (196). CD36 appears most important in the setting of lower concentrations of FFAs (58). Therefore, it is not surprising that when large amounts of FFA are generated during hydrolysis of large TG-rich lipoproteins like chylomicrons, heart uptake of lipids appears to be exclusive of this receptor (30). Lipolysis of lipoproteins is also a pathway for delivery of esterified core lipids such as cholesteryl esters and retinyl esters into the heart (30).

Excess lipid, especially TG, beyond that needed for cellular structures and ATP generation is stored in lipid droplets (Figure 4B). Within the heart, there normally is little lipid droplet accumulation, suggesting that uptake and oxidation are finely regulated. Lipid droplets are found in hearts of patients with diabetes and metabolic syndrome (197-199) and in those of high-fat diet fed rodents and genetically altered mice (see below and Table 3). In addition, after an overnight fast, lipid droplets appear in the hearts of wild type mice (200).

Lipid droplet protein makeup in the heart is different from that of adipocytes. In the heart, there is minimal expression of PLIN1. However, the other major lipid droplet proteins, PLIN2,

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PLIN3, PLIN4, and PLIN5 are all expressed in the heart (201). Plin2 expression might be most upregulated in some forms of lipotoxicity and be important for non-toxic lipid storage (202).

PLIN5 appears to regulate TG oxidation by approximating lipid droplets and mitochondria

(203,204). Of these droplet proteins, only Plin2 has been deleted and chow-fed Plin2-/- mice do not have an obvious cardiac phenotype (128). Thus, knowledge of how and whether these proteins, and probably others, modulate heart lipid accumulation and TG oxidation is likely to be forthcoming.

Lipid droplet turnover is regulated by lipid droplet associated proteins, intracellular lipases, and acyltransferases (205). Cardiac myocytes and skeletal myocytes have similar regulatory pathways that govern lipid metabolism. Lipid droplet TG can be hydrolyzed by ATGL and HSL, both of which are expressed in the heart. In the adipose tissue, insulin inhibits lipolysis, whereas catecholamines, thyroid hormone, and glucagon stimulate lipolysis. Whether similar regulation occurs in the heart is not known at present.

The roles of lipolytic enzymes in the heart have been studied using genetically modified mice (206). Lipid droplet accumulation in overnight fasting mice is prevented by overexpression of HSL (207). In the total HSL knockout mouse, cardiac TG lipase activity was decreased, but cardiac TG was not dramatically changed, and there was no overt cardiac phenotype (126). In contrast, Atgl-/- mice have markedly reduced cardiac TG lipase activity, massive lipid accumulation, and severe cardiomyopathy (127). In part, this is likely due to a defect in the hydrolysis of TG that the heart stores for potential energy (208). Treatment of Atgl-/- mice with a

PPARα agonist corrected the cardiac phenotype (170). Therefore, the excess accumulation of TG in the Atgl-/- hearts was at least partially due to increased lipid storage secondary to defective FA

34 oxidation. A less dramatic phenotype but one also associated with decreased FA oxidation activation occurred with cardiac deletion of Acsl1 (4). This study and another from this group (3) suggest that PPAR activation is via a product of the ACSL reaction.

Recent studies have elucidated the role of autophagy in hepatic TG lipolysis. In the rodent heart, autophagy has been investigated as a stress response mechanism in myocardial infarction and pressure induced cardiac hypertrophy. Fasting induces autophagy in the heart

(209), yet characterization of autophagy in the heart has not focused on lipid metabolic derangements. Inducible heart-specific autophagy knockouts develop cardiomyopathy (210); whether this is associated with increased lipid accumulation, as has been found in other organs

(211), is unknown.

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Figure 4 - Cartoon of cardiac lipid metabolism

A. FAs esterified as triacylglycerol (TG) within lipoproteins require hydrolysis by lipoprotein lipase

(LpL) associated with proteoglycans and GPIHBP1 on the luminal surface of endothelial cells.

ANGPTL4 is an LpL inhibitor. FFAs associated with albumin likely are internalized by membrane transporters such as CD36. These lipids must cross the endothelial barrier; how this occurs is unclear. B.

Within the cardiomyocytes the FAs are esterified to CoA and either stored in the LD or used for energy.

At least four lipid droplet proteins (PLINs) are expressed in the heart. The lipid droplet supplies some oxidized FAs via the actions of adipose TG lipase ATGL and HSL. CGI58 is the ATGL co-activator.

ATGL and LpL actions both provide ligands for PPAR activation.

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Creation of cardiac lipotoxicity in mice

A number of reasons for the association between diabetes and heart dysfunction in the absence of underlying vascular disease have been proposed; one of these is excess accumulation of lipids in cardiomyocytes (212). This possible cause of cardiomyopathy has been modeled by creating genetically modified animals in which lipid accumulation without generalized metabolic derangements leads to contractile impairment (Figure 5). These animals have an imbalance between lipid uptake and oxidation due to either increased lipid uptake or decreased oxidation.

Increased uptake of circulating FFAs or lipoprotein-derived lipids as occurs with transgenic expression of LpL (213,214) leads to reduced heart function. Transgenic mice with cardiomyocyte specific expression of FATP1 (215) and ACSL1 (216) are thought to have increased FFA uptake or trapping in the heart leading to heart failure. PPAR transcription factors drive FA oxidation; however, the increased lipoprotein-lipid uptake in PPARα transgenic mice

(217) must exceed the increased FA oxidation found in this model because the hearts have excess stored lipids. Cardiomyocyte PPARγ overexpression leads to a similar phenotype (202).

Surprisingly, PPARδ expression does not lead to cardiac dysfunction or toxicity (218), presumably because upregulation of the LpL inhibitor ANGPTL4 prevents excess lipid uptake

(219).

LpL is the key enzyme for distribution of circulating lipids between organs. Perhaps the most clinically relevant model of cardiac lipotoxicity is one created by accident. Wang et al. deleted LpL using a skeletal muscle specific promoter (49). The mice were meant to model physically inactive humans who also have reduced muscle LpL and FA oxidation. These mice have increased insulin sensitivity in skeletal muscle, as would be expected with reduced FA

37 uptake, but develop insulin resistance in the heart, often a precursor of eventual heart dysfunction.

Reduced lipid oxidation can also lead to lipid accumulation and cardiomyopathy. This occurs with cardiac specific knockout of PPARδ (220) and a cardiac LpL transgene crossed onto the Ppara-/- background (214). Similarly, heart dysfunction with excess lipid accumulation is found when heart-specific GLUT1 overexpressing mice are placed on a high fat diet (221). In contrast, when these animals eat chow their function is improved in the presence of hypertension

(222). Therefore, in some situations, dietary driven lipid uptake – a likely accompaniment of our western diet – needs to be added to create a lipotoxic environment.

Perhaps of most interest for understanding heart lipid metabolism and toxicity are the situations where reduced FA oxidation does not lead to lipid accumulation. In many cases, the reason for this has not been investigated, but we would presume that there is a compensation, such as a reduction in lipid uptake. This occurs with genetic or pharmacologic deficiency of

DGAT1, which markedly reduces CD36 expression (223). Pharmacologic FA oxidation inhibitors proposed for reduction of ischemia (190) and genetically engineered defects in FA oxidation defects (224) would be expected to also create cardiac lipid accumulation and toxicity.

That this does not occur indicates the existence of some processes that balance reduced oxidation or that lead to non-toxic lipid storage.

Lipid stores and the causes of toxicity

Although the most obvious and easiest to measure accumulated lipid is TG, TG itself might not be toxic. Consistent with cellular studies (225), several experimental situations have dissociated TG accumulation from toxicity. Total body knockout of HSL was associated with

38 more re-feeding TG accumulation, but no toxicity (226). A cross of the PPARγ transgene onto the

Ppara-/- background corrected toxicity without reducing heart TG, ceramide or DAGs, but redistributed the lipids into larger droplets (227).

Although the preferred energy substrate for the heart is FA, a case has been made that excess reliance on FA oxidation is harmful, even under non-ischemic conditions. Some experimental data suggest the opposite. High fat diets, which usually increase reliance of FA oxidation, may be beneficial in the setting of non-ischemic heart failure (228). Increased FA oxidation, has been found with a Pparγ transgene crossed onto the Ppara-/- background (227),

Dgat1 transgenic expression (229), and most recently with PPARα agonist treatment of Atgl-/- mice (170) all of which improve heart failure.

Heart lipid content can also be increased by non-genetic means. While in some ways these models may more closely reflect human pathology, they suffer from the many other systemic effects of over-nutrition or diabetes. In addition, the heart phenotypes are relatively mild compared to those found with genetic modifications. Mice fed a high fat diet rapidly develop cardiac insulin resistance, suggesting that lipid accumulation rapidly causes changes in heart metabolism allowing it to rely more on FAs as its fuel (230). However, the effects of high fat on cardiac function are mixed. Hypertensive rats fed a 60% fat diet had less left ventricular hypertrophy and systolic dysfunction than animals fed 10% fat (228). Similarly, rats fed a high- fat diet appeared to compensate by increasing FA oxidation, whereas those eating a lower fat but higher carbohydrate western diet developed dysfunction (231). Not all fats are equal. Saturated

FA-rich diets alone increased cardiomyocyte apoptosis, perhaps due to accumulation of ceramide

39

(232). In contrast, medium chain FAs are protective against cardiac dysfunction in some situations (233,234).

Other pharmacologic and genetic alterations change heart TG content. Using oxfenicine to block CPT-1, mice fed a diet enriched in long-chain saturated FA accumulated TG, but had no changes in left ventricular dimensions or systolic function, while PPAR-regulated genes were upregulated (235). Ob/ob and db/db mice have increased heart FA oxidation that develops prior to hyperglycemia (236). Eventually these mice develop decreased contractile function (237).

Correction of lipotoxic heart disease

A role of animal models is to test interventions that might be beneficial in human disease.

Genetic approaches define targets, which might be amenable to pharmacologic or dietary interventions. Although a common underlying theme of lipotoxic cardiomyopathy is that it is created via an alteration in lipid metabolism, a single underlying toxic lipid species might not be causative in all case. Thus, interventions in one model might not prove to be beneficial in another. Similarly, the genetic and dietary variation amongst humans might also lead to multiple causes of cardiomyopathy associated with lipid accumulation.

Altering the amount or type of FAs acquired by the heart will prevent toxicity both in genetic and dietary models of lipid toxicity. Deletion of either CD36 or LpL corrected the cardiac toxicity associated with cardiomyocyte overexpression of PPARα (217,238). CD36 deletion was also reported to improve heart function in aged mice that were eating a diet enriched in medium chain FAs (239). Medium chain FA-rich diets were also beneficial in

PPARα transgenic mice (240), presumably because of a reduction in saturated long chain fat- enriched lipid accumulation. Other methods to reduce heart lipid content may also correct or

40 prevent toxicity (Table 3). The approaches to do that have included transgenic expression of

HSL to increase lipolysis of stored lipids (241), reduced expression glycerol-3-phosphate acyltransferase-1 to decrease TG accumulation (242), and overexpression of ApoB to increase cardiac lipid secretion (243) .

A targeted approach to modify one lipid species would seem to be an ideal way to both treat the disease and define the toxic lipid species. Such an approach was taken by reducing ceramide concentrations in LpL-overexpressing mice using the serine palmitoyl transferase inhibitor myriocin and SPT deficient mice (244). A similar attempt to modify a specific toxic lipid, DAG was attempted by overexpressing DGAT1 in cardiomyocytes. Although this intervention reduced DAG concentrations and heart dysfunction, ceramide was also reduced

(229). This study suggests that alteration of a single lipid species in the heart might be difficult due to the inter-relationship of lipid metabolism pathways.

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Table 3 - Correction of cardiac lipotoxicity

Reference Cardiac Lipotoxicity Reference Corrections

Leptin treatment (245) MHC-ACS1 (216) (229) X MHC-DGAT1 (246) α-Lipoic acid MHC-FATP1 (215)

Heart specific PPAR δ (220) knockout

X heart specific LpL knockout (217) MHC-PPARα (247) (238) X CD36-/- (240)

Medium chain TG diet

MHC-PPARγ (202) X PPARα knockout (227)

(248) Leptin deficiency (ob/ob) Leptin Infusion (249) (249)

ATGL knockout (140) PPARα agonists (170)

Ceramide synthesis MHC-LpLGPI (213) (244) inhibition MHC-LPL X (214) Ppara-/-

MHC-GLUT1 on HFD (221)

42

Figure 5 - Correcting cardiac lipotoxicity

Genetically modified mice have been created that have either increased lipid uptake or decreased oxidation. Uptake occurs via the cell surface molecules lipoprotein lipase (LpL) and perhaps the

FA transporter CD36 and/or FATPs. More FAs are “trapped” by complexing to CoA. Stored TG accumulates with defective hydrolysis due to deletion of ATGL. Two interventions, loss of

PPARδ and high fat feeding in mice overexpressing Glut1, lead to reduced lipid oxidation.

Transgenic expression of PPARα and PPARγ, which induce lipid oxidation genes, also cause lipid accumulation, presumably because uptake exceeds oxidation.

43

Conclusions

Although lipid-induced cardiac toxicity clearly can be created, a number of clinical and experimental issues await clarification. We do not know about the exact clinical implications of this pathophysiologic phenomenon. Is it really a primary cause of cardiac dysfunction in patients with type 2 diabetes and metabolic syndrome? If so, does this occur because of unregulated cardiac uptake of FAs or reduced FA oxidation? If the latter, one could then postulate that in the setting of ischemia and afterload-induced heart failure, reduced FA oxidation should also promote accumulation of toxic lipids. Alternatively, as has been postulated in ischemia, does a disproportionate use of FAs create toxicity? Which lipids are toxic and which pathways are induced that lead to cellular dysfunction? A marriage of human and animal experimentation should sort out the answers to many of these questions in the next decade.

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Chapter 3: Lipoprotein lipase activity is required for cardiac lipid droplet production

This chapter has been published in the Journal of Lipid Research (250).

Chad M. Trent; Shuiqing Yu; Yunying Hu; Nathan Skoller; Lesley A. Huggins; Shunichi

Homma; Ira J. Goldberg

Abstract

The rodent heart accumulates TG and lipid droplets during fasting. The sources of heart lipids could be either FFAs liberated from adipose tissue or FAs from lipoprotein-associated TG via the action of lipoprotein lipase (LpL). Because circulating concentrations of FFAs increase during fasting, it has been assumed that albumin transported FFAs are the source of lipid within heart lipid droplets. We studied mice with three genetic mutations: peroxisomal proliferator-activated receptor alpha deficiency, cluster of differentiation (CD) 36 deficiency, and heart specific LpL deletion. All three genetically altered mice had defective accumulation of lipid droplet TG.

Moreover, hearts from mice treated with poloxamer 407, an inhibitor of lipoprotein TG lipolysis, also failed to accumulate TG, despite increased uptake of FFAs. TG storage did not impair maximal cardiac function as measured by stress echocardiography. Thus, LpL hydrolysis of circulating lipoproteins is required for the accumulation of lipids in the heart of fasting mice.

Introduction

The human heart will accumulate TG in lipid droplets in disease states such as obesity and diabetes. Whether TG storage directly leads to reduced heart function, i.e. lipotoxicity (182,251), or is a marker for accumulation of other toxic lipids is unclear. Evidence suggesting that stored

TG in cardiomyocytes is not always toxic has come from experiments in genetically modified

45 mice. For instance, overexpression of the final enzyme in TG synthesis, DGAT1 in cardiomyocytes increased TG stores but reduced accumulation of toxic lipids and did not reduce heart function (229). Overexpression of DGAT1 in skeletal muscle also increased TG storage in mice with diet-induced obesity and mimicked the “athlete’s paradox” observed in endurance- trained humans; skeletal muscle DGAT1 transgenic mice had increased FA oxidation and improved insulin sensitivity (252). In contrast, increased TG accumulation in the human heart correlates with reduced heart function (197,199). Moreover, greater TG stores are often associated with greater FA oxidation and greater injury during ischemia/reperfusion in isolated perfused hearts (190). Thus, the role of TG stores in the heart is unclear.

Even the substrate used for heart TG production has not been established. One physiologic stimulus that causes lipid accumulation in mouse hearts is prolonged fasting (200).

Since starvation is a threat to survival, lipid accumulation in the heart may be an adaptation to accommodate future energetic demands, to protect the heart from lipotoxicity, or to do both.

Understanding the features of this adaptation may provide insight into mechanisms that drive lipid accumulation under physiologic and pathological conditions. During fasting, animals rely exclusively on stored energy. While the liver produces and releases glucose under fasting conditions, this is insufficient to meet the energetic demands of the body (253). Adipose tissue is the major storage depot for energy in the form of TG. During the fed state, dietary glucose stimulates insulin secretion, which simultaneously promotes glucose utilization and lipid storage.

During fasting, circulating insulin concentrations fall while glucagon and catecholamines increase. This shift in the hormonal milieu leads to an activation of glycogenolysis in the skeletal muscle and liver and lipolysis in the adipose tissue. However, prolonged fasting will deplete glycogen stores and thus the energy demands of peripheral tissues must rely on both the liver –

46 to secrete glucose, ketone bodies, and TG – and the adipose tissue – to secrete FFAs and glycerol. Adipose tissue secreted glycerol, as well as lactate secreted from both adipose tissue and muscle, are taken up by the liver and used as substrates for gluconeogenesis.

We tested whether reduced FA oxidation increased fasting-induced TG accumulation in the heart. To do this we studied peroxisomal proliferator-activated receptor (PPAR) α knockout mice. Surprisingly, we found that fasted Ppara-/- mice had no lipid droplet accumulation in hearts and had a marked reduction in mRNA levels of the FA transporter cluster of differentiation (CD) 36 as well as lipoprotein lipase (LpL) (254,255); LpL is required for heart uptake of FFAs from lipoprotein TG. We then assessed the specific roles of CD36 and LpL in heart TG accumulation. Our data show that LpL activity is required for the accumulation of heart lipid droplets. In addition, we demonstrated that lipid droplet accumulation does not affect maximal systolic function of the heart.

Materials and Methods

Animals and Fasting We used 3-4 month old male C57BL/6 mice, Ppara-/- mice (166), Cd36-/- mice (55), floxed LpL mice (LpLflox/flox) and heart specific LpL knockout (hLpL0) mice (48).

Mice were fed a normal chow diet. C57BL/6 mice were used as controls for both Ppara-/- and

Cd36-/- mice and LpLflox/flox littermates served as controls for the hLpL0 studies. Mice of each genotype were divided into two groups. One group was subjected to a 16-hour overnight fast and the other group was fed ad libitum over the same time period. These mice were then sacrificed with a lethal injection of 100 mg/kg ketamine and 10 mg/kg xylazine. All procedures were approved by the Columbia University Institutional Animal Care and Use Committee.

47

Tissue Collection A ventral incision was made after administration of 10 mg/kg ketamine- xylazine. The left ventricle of the heart was perfused with 10 ml of PBS or until the liver appeared blanched. Tissues were rapidly removed and frozen in liquid nitrogen. Heart pieces were embedded into Tissue-Tek O.C.T. compound (Sakura, USA) for histology.

Measurement of Plasma Lipids and Glucose 200 µL of blood was drawn from each animal after administration of ketamine/xylazine and then centrifuged at 2,000 rpm for 10 minutes to obtain plasma. Plasma was utilized for measurement of TG, FFA, and glucose by colorimetric assays. TG measurements were made using the Thermo Scientific Infinity assay (Thermo

Scientific, USA), FFA was measured using the Wako NEFA kit, and plasma glucose was measured using the Wako Autokit Glucose kit (Wako Life Sciences, USA).

Lipid Extraction from Tissues The lipid extraction protocol was adapted from the Folch method (256). Approximately 100 mg of ventricular tissue in 1 mL of PBS were homogenized using stainless steel beads for 1 minute in a bead beater homogenizer. From each sample, 50 µL were removed for protein analysis and 3 ml of 2:1 chloroform: methanol was added to the rest and vortexed. Samples were then centrifuged for 10 minutes at 3000 rpm at 4°C. The lower, organic phase was then collected and dried under nitrogen gas. The dried lipid was then dissolved in 500 µL of 1% Triton-X 100 in chloroform, further dried and then dissolved in 100µl of double distilled water.

Lipid and Protein Measurements of Tissues The sample of tissue lysate retained from the lipid extraction protocol was assayed for protein content using Bradford reagent (Bio-Rad, USA) following the instructions of the manufacturer. Using the tissue lipid extract, assays for TG and

48

FFA were performed as previously described for plasma lipids. Lipid measurements were normalized to protein content or tissue weight.

Microscopy for Cardiac Lipid Visualization Heart pieces were embedded in Tissue-Tek

O.C.T. compound (Sakura, USA) and then air-dried and fixed with formalin. Sections were washed with distilled water and isopropanol. Lipids were then stained with Oil Red O for 18 minutes, washed with isopropanol and distilled water, and then counterstained with hematoxylin.

Slides were once again washed with distilled water and covered with clear nail polish. Images were taken using a Leica DMLB microscope and digital camera. Shown are representative images obtained from 5 animals of each genotype.

Glycogen Staining and Quantification Periodic acid – Schiff reagent (PAS) staining was used to demonstrate heart glycogen. Sections of OCT embedded hearts were placed in 10% neutral buffered formalin. Ventricular tissue sections were fixed in methanol for 10 minutes and stained with PAS (Poly Scientific), hematoxylin, and eosin. Images were taken using a Leica DMLB microscope and digital camera. 4-5 mouse hearts were used for each genotypes and for each feeding condition, and several representative images were captured for each mouse.

Glycogen was also measured by extracting total insoluble carbohydrates and digesting with amyloglucosidase, and free glucose was then measured and reported as ratio to tissue weight used for measurement as previously described (257). Ventricular tissue was hydrolyzed in 300

o µL of 5.4 M KOH in a 100 C water bath for 30 minutes. Then, 100 µL 1 M Na2SO4 and 800 µL of 100% ethanol were added to each sample. Samples were boiled for 5 minutes and then centrifuged at 10,000 G for 5 minutes. The glycogen pellet was dissolved in 200 µL water and ethanol precipitation was performed twice with addition of 800 µL of 100% ethanol. Finally, the

49 glycogen pellet was dissolved in 200 µL of 60 units/mL amyloglucosidase (Sigma, USA) in 0.2

M sodium acetate (pH 4.8) and incubated for 3 hours at 40oC. Each sample was diluted 5X and glucose concentration was measured using the Wako Autokit Glucose kit (Wako Life Sciences,

USA).

Cardiac Gene Expression Total RNA was purified from a 30-50 mg piece of heart ventricular tissue using the TRIzol reagent (Invitrogen, USA) according to the instructions of the manufacturer. cDNA was synthesized using the SuperScript III First-Strand Synthesis SuperMix

(Invitrogen, USA) and quantitative real-time PCR was performed with SYBR Green PCR Core

Reagents (Agilent Technologies, USA) using an Mx3000 sequence detection system (Stratagene,

La Jolla, CA). Genes of interest were normalized against 18s rRNA. Primer sequences are listed in Table 4.

Western Blotting Hearts were excised as previously described. Approximately 20 mg of ventricular tissue was homogenized in RIPA buffer containing protease inhibitor cocktail

(Sigma-Aldrich, USA). 25 µg of protein extract was applied to SDS-PAGE and transferred onto

PVDF membranes. Antibodies for PLIN2 and PLIN5 were obtained from Santa Cruz

Biotechnology (PLIN2, β-actin) and American Research Products (PLIN5). Band density measurements were made using ImageJ software. PLIN2 and PLIN5 band densities were normalized to β-actin band density.

Stress Echocardiography Echocardiography was performed on 3-4 month old male Cd36+/+

(wild type), Cd36-/-, LpLflox/flox, and hLpL0 mice fasted for 16 hours. Two-dimensional echocardiography was performed using a high-resolution imaging system with a 30-MHz imaging transducer (Vevo 770; VisualSonics, USA) in unconscious mice. The mice were

50 anesthetized with 1.5-2% isoflurane and thereafter maintained on 0.5% isoflurane throughout the procedure. Care was taken to minimize sedation by monitoring the heart rate of the mice; measurements were only made after the heart rate was 400-500 beats per minute. Two- dimensional echocardiographic images were obtained using short-axis views at the level of papillary muscles, and each parameter was measured using M-mode view. Images were analyzed offline by a researcher blinded to the murine genotype. Left ventricular end-diastolic dimension

(LVEDd) and left ventricular end-systolic dimension (LVEDs) were measured. Percentage fractional shortening (FS), which quantifies contraction of the ventricular wall and is an indication of muscle function, was calculated as FS = ([LVEDd − LVEDs] / LVEDd) × 100. To assess stress response 0.3-mg/kg isoproterenol (Sigma-Aldrich, USA) was administered intraperitoneally. Successful administration of drug was confirmed by observation of increased heart rate.

Pharmacologic Inhibition of LpL Poloxamer 407 (P407) was prepared in PBS as previously described (258). Mice were injected intraperitoneally with 1 mg/g body weight of P407 and then fasted for 16 hours. Control mice were injected with an equivalent volume of PBS. Mice were killed and analyzed as previously described.

In vivo Assessment of Cardiac Glucose and FFA Uptake FFA and glucose uptake was assessed in mice that were injected with PBS or P407 and then fasted for 16 hours. [9,10-3H(N)] oleate (PerkinElmer Life Sciences) was complexed to 6% FA-free bovine serum albumin

(Sigma). Mice were injected intravenously with 1 µCi [9,10-3H(N)] oleate-bovine serum albumin and blood was collected at 0.5, 1, 3, and 5 minutes after injection. Five minutes after injection, the left ventricle of the heart was perfused with 10 ml of PBS by cardiac puncture and tissues

51 were excised. Tissue was homogenized in PBS and radioactive counts were measured. Basal glucose uptake was measured in hearts following an intravenous administration of 2.5 µCi of 2- deoxy-D-[1-14C] glucose (PerkinElmer Life Sciences). Blood was collected 2, 30, and 60 minutes following injection. At 60 minutes, hearts were perfused with PBS, tissues excised, and radioactive counts measured. For all turnover studies, radioactivity per gram of tissue was normalized to the respective 30 second or 2 minute plasma counts (injected dose).

Density Ultracentrifugation of Plasma Approximately 60 µL of mouse plasma was separated by density ultracentrifugation as previously described (259). Briefly, 60 µL of plasma was underlayed to 60 µL of 1.006 g/mL saline solution in an ultracentrifuge tube. Each tube was centrifuged at 70,000 RPM at 12oC for 3 hours. The supernatant was removed and contains

VLDL. Next, 60 µL of 1.12 g/mL KBr solution was added to the remaining infranatant. This was centrifuged at 70,000 RPM at 12oC for 12 hours. The supernatant was removed and contains

LDL. The infranatant was removed and 60 µL of 1.34 g/mL mixture of KBr and NaCl added.

This was centrifuged at 70,000 RPM at 12oC for 12 hours. The supernatant was removed and contains HDL. The remaining infranatant contains all other plasma proteins, including FFA complexed to albumin. Counts of [3H]-oleate was measured in each fraction and reported as percentage of total counts measured.

Ceramide Quantification

Ceramides were measured as previously described (260). All solvents for sample extraction and

LC/MS were LC/MS grade (or LC grade when LC/MS grade was not available) and were purchased from Fisher Scientific (Pittsburgh, PA, USA). Ceramide standards were purchased from Avanti Polar Lipid, Inc. (Alabaster, AL, USA). Samples were processed as described

52 previously with modification (2). Briefly, 3 ml of the methanol containing 20 µl of a 2 µM internal standard mixture (Avanti LM-6002, containing C12 and C25 ceramides) were added to

100 µl aqueous of heart homogenate containing 10 mg heart tissue in a clean glass tube, vortex- mixed and allowed equilibrate for 10 minutes on ice. Then the mixture was vortexed again and centrifuged at 3,000g for 10 minutes at 4 C degree. The organic upper phase was transferred to a second clean glass tube and evaporated under nitrogen. The extracted lipids were reconstituted in 300 µl of methanol: acetonitrile (v:v=1:1) and transferred to LC/MS autosampler vials

(Waters, P/N 600000670CV) for evaporation under nitrogen. The lipid extract was finally reconstituted in 50 µl methanol for injection. All LC/MS/MS running were carried out on a

Waters Xevo TQ MS ACQUITY UPLC system (Waters, Milford, MA) controlled by Mass Lynx

Software version 4. 1. The sample was maintained at 4°C in the autosampler and 5 µl was loaded onto a Waters ACQUITY UPLC BEH Phenyl column (3 mm inner diameter × 100 mm with 1.7

µm particles), preceded by a 2.1 × 5 mm guard column containing the same packing. The UPLC flow rate was continuously 300 µl/min in a binary gradient mode with methanol and water both containing 0.2% formic acid and 1 mM ammonium formate. Positive ESI-MS/MS mass spectrometry was performed to identify ceramide species of interest. Different species were confirmed by comparing the retention times of experimental compounds with those of authentic standards. Concentrations of ceramides in the samples were quantified by comparing integrated peak areas for those of each ceramide against those of known amounts of purified standards.

Loss during extraction was accounted for by adjusting for the recovery of the internal standard added before extraction.

Triglyceride lipase activity assay TG lipase activity was performed as described (137,143).

Tissue lysates were prepared in solution A (0.25 M sucrose, 1 mM EDTA, 1 mM DTT, pH 7.0, 1

53

µg/mL pepstatin, 2 µg/mL antipain, and 20 µg/mL leupeptin). Protein concentration was measured with Bradford reagent, and 50 µg of protein was used for each assay. Micelles were prepared by sonication in 0.1 M potassium phosphate buffer (pH 7.0) with 1.67 mM triolein, 10

µCi [9,10-3H(N)] triolein/mL (Perkin-Elmer), 142.5 mM phosphatidylcholine, and 47.5 mM phosphatidylinositol. FA free BSA was added to the micelles at 2% concentration

(weight/volume). 100 µL of radiolabeled substrate was incubated with diluted protein lysates for

1 hour at 37oC in a 125 RPM shaking incubator. The reaction was stopped by the addition of

3.25 mL mixture of methanol, chloroform, and n-heptanes (10:9:7, volume). FAs were extracted by addition of 1 mL of 0.1 M potassium carbonate, 0.1 M boric acid (pH 10.5) and centrifugation at 3000 RPM for 10 minutes. 200 µL of the aqueous phase were counted for radioactivity.

Statistical Analysis Data are expressed as mean + SE. Data were analyzed by the use of unpaired Student’s t-test or 2-way ANOVA.

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Table 4 - Primer sequences for "LpL activity is required for cardiac lipid droplet production"

Gene Forward primer Reverse primer 18s 5'-CCATCCAATCGGTAGTAGCG-3' 5'-GTAACCCGTTGAACCCCATT-3' Abhd5 5'-TGACAGTGATGCGGAAGAAG-3' 5'-AGATCTGGTCGCTCAGGAAA-3' Acc2 5'-TGGAGTCCATCTTCCTGTCC-3' 5'-GGACGCCATACAGACAACCT-3' Acox1 5'-GGATGGTAGTCCGGAGAACA-3' 5'-AGTCTGGATCGTTCAGAATCAAG-3' Atgl 5'-CGCCTTGCTGAGAATCACCAT-3' 5'-AGTGAGTGGCTGGTGAAAGGT-3' Cd36 5'-TGTGTTTGGAGGCATTCTCA-3' 5'-TGGGTTTTGCACATCAAAGA-3' Cpt1b 5'-TCTAGGCAATGCCGTTCAC-3' 5'-GAGCACATGGGCACCATAC-3' Dgat1 5'-GTGCACAAGTGGTGCATCAG-3' 5'-CAGTGGGATCTGAGCCATCA-3' Dgat2 5'-CTGTCACCTGGCTCAACAGA-3' 5'-TATCAGCCAGCAGTCTGTGC-3' Fabp3 5'-GACGAGGTGACAGCAGATGA-3' 5'-TGCCATGAGTGAGAGTCAGG-3' Fasn 5'-TTGCTGGCACTACAGAATGC-3' 5'-AACAGCCTCAGAGCGACAAT-3' Fatp6 5'-GGTCACGGTGCTGGATAAGT-3' 5'-CGAGGAGTGGTTCAGGAGAG-3' Glut1 5'-GCTGTGCTTATGGGCTTCTC-3' 5'-CACATACATGGGCACAAAGC-3' Glut4 5'-ACTCTTGCCACACAGGCTCT-3' 5'-CCTTGCCCTGTCAGGTATGT-3' Got2 5'-GTTGAAATGGGACCTCCAGA-3' 5'-GGGCAGGTATTCTTTGTCCA-3' Lpl 5'-GCTGGTGGGAAATGATGTG-3' 5'-TGGACGTTGTCTAGGGGGTA-3' Pdk4 5'-TTCTCGGAGTCTGGAATGCT-3' 5'-GCTCTAGCCGAACACGAATC-3' Plin2 5'-CTACGACGACACCGAT-3' 5'-CATTGCGGAATACGGAG-3' Plin5 5'-GTGATCAGACAGCTCAGGACCCT-3' 5'-CGATTCACCACATTCTGCTGG-3' Scd1 5'-TGCGATACACTCTGGTGCTC-3' 5'-TAGTCGAAGGGGAAGGTGTG-3' Slc27a1 5'-TTCTCGGAGTCTGGAATGCT-3' 5'-GCTCTAGCCGAACACGAATC-3'

55

Results Fasted Ppara-/- mice do not store lipids in the heart

We first assessed heart lipid storage in Ppara-/- mice. We fasted Ppara+/+ (wild type) and Ppara-/- mice overnight for 16 hours. Fasting increased plasma FFA 2-fold in Ppara+/+ mice and 3-fold in

Ppara-/- mice, but had no significant effect on plasma TG (Figure 6). Plasma glucose decreased approximately 30% in fasted Ppara+/+ mice and approximately 60% in fasted Ppara-/- mice.

Fasting increased heart TG 5-fold in Ppara+/+ mice, but there was no significant TG accumulation in Ppara-/- mice (Figure 7). Heart FAs increased approximately 30% in Ppara+/+ mice, but were not increased in Ppara-/- mice (Figure 7). Fasted Ppara+/+ mice had increased Oil

Red O staining, but Ppara-/- mice had minimal staining (Figure 8).

Figure 6 - Plasma biochemistry of fed and fasted Ppara+/+ and Ppara-/- mice

Plasma FFA, TG, and glucose were measured in 3-4 month old male Ppara+/+ (n=9) and Ppara-/-

(n=5) mice that were fed or fasted for 16 hours. * indicates p < 0.05, data were compared by

Student’s t-test.

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Figure 7 - Lipid measurements from hearts of fed and fasted Ppara+/+ and Ppara-/- mice

Total lipids were extracted from hearts, and TG and FFA were measured. Lipid content was normalized to protein content. * indicates p < 0.05, data were compared by Student’s t-test.

57

Figure 8 - ORO staining of hearts from fed and fasted Ppara+/+ and Ppara-/- mice

Fed and fasted Ppara+/+ and Ppara-/- mice heart sections (n=5) were stained with Oil Red O, indicating neutral lipid content (1000X).

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Figure 9 - PAS staining of hearts from fed and fasted Ppara+/+ and Ppara-/- mice

Fed and fasted Ppara+/+ and Ppara-/- mice heart sections (n=5) were stained with PAS reagent, indicating glycogen content (400X)

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Figure 10 - Glycogen content of hearts from fed and fasted Ppara+/+ and Ppara-/- mice

Total glycogen was extracted from hearts of fed and fasted Ppara+/+ and Ppara-/- mice and digested with amyloglucosidase, and free glucose was measured. Glucose content was normalized to tissue weight.

60

We then assessed heart glycogen storage in Ppara-/- mice to determine if these hearts depleted their stored carbohydrate. There was no difference in PAS staining of glycogen (Figure

9) or extracted glycogen content (Figure 10) after fasting. There tended to be increased glycogen in Ppara-/- hearts both before and after fasting.

Changes in genes required for lipid and glucose metabolism were determined in hearts of fed and fasted mice. Adipose TG lipase (Atgl), the rate limiting enzyme for intracellular TG lipolysis (127), was decreased by 50% in both fed and fasted Ppara-/- mice compared to Ppara+/+

(Figure 11). Expression of carnitine palmitoyl transferase (Cpt)1b, the rate limiting enzyme for mitochondrial lipid oxidation, was minimal in both fed and fasted Ppara-/- hearts. Surprisingly, mRNA of acyl CoA oxidase (Acox)1, the first enzyme in the lipid oxidation pathway, was increased in fasted Ppara-/- hearts. However, decreased FA oxidation has been previously observed in these hearts (261,262). Therefore, absence of TG stores was not likely due to increased FA oxidation. As expected, mRNA expression of lipid droplet protein genes Plin2 and

Plin5 were minimal in both fed and fasted Ppara-/- mice compared to Ppara+/+ mice (169).

Fasting dramatically increased expression of pyruvate dehydrogenase kinase (Pdk) 4, the negative regulator of glucose oxidation, in hearts from Ppara+/+ mice. Fasted Ppara-/- mice also had increased Pdk4 mRNA levels compared to fed Ppara-/- but these levels were still reduced compared to the Ppara+/+ counterparts. Fasted Ppara-/- mice had increased glucose transporter

(Glut)1, the insulin insensitive glucose transporter, mRNA expression compared to fasted

Ppara+/+ hearts, but there was no difference in Glut4, the insulin sensitive glucose transporter, expression. Most remarkable was that fed and fasted Ppara-/- hearts had an ~80% reduction in lipid uptake genes Cd36 and Lpl.

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Figure 11 - Gene expression in hearts from fed and fasted Ppara+/+ and Ppara-/- mice

Gene expression of Atgl, Cpt1b, Acox1, Plin2, Plin5, Cd36, Lpl, Pdk4, Glut1, and Glut4 was assessed using quantitative real-time PCR. Gene expression is expressed relative to fed Ppara+/+ mice. * indicates p < 0.05 compared within genotype, # indicates p < 0.05 compared to Ppara+/+ mice of same feeding status; data were compared by 2-way ANOVA.

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We assessed heart mRNA levels of several genes involved in both de novo lipogenesis and TG formation. Expression of acetyl-CoA carboxylase (Acc) 2, which is rate-limiting for de novo lipogenesis, was increased in the hearts of fed Ppara-/- mice (FIGURE). FA synthase

(Fasn); the second rate-limiting enzyme for de novo lipogenesis; mRNA expression was increased in the hearts of both fed and fasted Ppara-/- mice. Dgat1, the rate limiting enzyme for

TG synthesis, mRNA levels were decreased in hearts from both fed and fasted Ppara-/- mice. We also measured gene expression of FA transporters other than Cd36. Hearts from both fed and fasted Ppara-/- mice had decreased expression of Slc27a1 (Fatp1), but increased expression of

FA binding protein-plasma membrane (Got2, or FABPpm). Finally, we measured TG lipase activity in hearts of fed and fasted Ppara+/+ and Ppara-/- mice. Both fed and fasted Ppara-/- mice had 80-90% increased heart TG lipase activity compared to Ppara+/+ of the same nutritional status.

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Figure 12 - Supplemental gene expression in hearts from fed and fasted Ppara+/+ and Ppara- /- mice

Gene expression of Slc27a1, Fatp6, Fabp3, Got2, Acc2, Fasn, Scd1, Dgat1, Dgat2, and Abhd5 was assessed using quantitative real-time PCR. Gene expression is expressed relative to fed

Ppara+/+ mice. * indicates p < 0.05 compared within genotype, # indicates p < 0.05 compared to

Ppara+/+ mice of same feeding status; data were compared by 2-way ANOVA.

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Figure 13 - TG lipase activity in hearts from fed and fasted Ppara+/+ and Ppara-/- mice

Heart tissue lysates from fed and fasted Ppara+/+ and Ppara-/- mice (n=5) were incubated with

[3H]-labelled triolein micelles to assess TG lipase activity. # indicates p < 0.05 compared to

Ppara+/+ mice of same feeding status; data were compared by 2-way ANOVA.

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Fasted Cd36-/- mice do not store lipids in the heart

Next, we determined whether CD36 deficiency would be sufficient to prevent heart TG accumulation during the fasted state. Fasting increased plasma FFA 3-fold in Cd36-/- mice

(Figure 14). Plasma TG tended to be higher in fasted Cd36-/- mice compared to Cd36+/+ mice.

Fasted Cd36-/- had a ~50% decrease in plasma glucose. There was no significant TG accumulation in Cd36-/- hearts (Figure 15). Heart FA content also did not increase in hearts from

Cd36-/- mice. Glycogen content was similar in all hearts (Figure 16). Finally, Cd36-/- mouse hearts tended to have decreased expression of lipid metabolism genes (Atgl, Cpt1b, Acox1, Atgl,

Plin2, Plin5) in the fed state, but fasted Cd36-/- mice hearts had similar gene expression to

Cd36+/+ mice (Figure 19). LpL mRNA levels were comparable to control mice in both fasting and fed hearts. Glucose oxidation and uptake genes (Pdk4, Glut1, and Glut4) were comparable between genotypes and feeding conditions. Fed and fasted Cd36-/- mice had decreased expression of Dgat2 (Figure 20). Hearts of fasted Cd36-/- mice had increased expression of Slc27a1, and both fed and fasted hearts had decreased expression of FA binding protein 3 (Fabp3). There were no differences in TG lipase activity between feeding conditions or genotype (Figure 21).

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Figure 14 - Plasma biochemistry of fed and fasted Cd36+/+ and Cd36-/- mice

Plasma FFA, TG, and glucose were measured in 3-4 month old male Cd36+/+ (n=9) and Cd36-/-

(n=9) mice that were fed or fasted for 16 hours. * indicates p < 0.05 compared within genotype,

# indicates p < 0.05 compared to Cd36+/+ mice of same feeding status; data were compared by 2- way ANOVA.

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Figure 15 - Lipid measurements from hearts of fed and fasted Cd36+/+ and Cd36-/- mice

Total lipids were extracted from hearts of Cd36+/+ and Cd36-/- mice, and TG and FFA were measured. Lipid content was normalized to protein content. *indicates p < 0.05, data were compared by Student’s t-test.

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Figure 16 - ORO staining of hearts from fed and fasted Cd36+/+ and Cd36-/- mice

Fed and fasted Cd36+/+ and Cd36-/- mice heart sections (n=5) were stained with Oil Red O

(1000X).

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Figure 17 - PAS staining of hearts from fed and fasted Cd36+/+ and Cd36-/- mice

Fed and fasted Cd36+/+ and Cd36-/- mice heart sections (n=5) were stained with PAS reagent

(400X).

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Figure 18 - Glycogen content of hearts from fed and fasted Cd36+/+ and Cd36-/- mice

Total glycogen was extracted from hearts of fed and fasted Cd36+/+ and Cd36-/- mice and digested with amyloglucosidase, and free glucose was measured. Glucose content was normalized to tissue weight.

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Figure 19 - Gene expression in hearts from fed and fasted Cd36+/+ and Cd36-/- mice

Gene expression of Atgl, Cpt1b, Acox1, Plin2, Plin5, Cd36, Lpl, Pdk4, Glut1, and Glut4 was assessed using quantitative real-time PCR. Gene expression is expressed relative to fed Cd36+/+ mice. * indicates p < 0.05 compared within genotype, # indicates p < 0.05 compared to Cd36+/+ mice of same feeding status; data were compared by 2-way ANOVA.

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Figure 20 - Supplemental gene expression in hearts from fed and fasted Cd36+/+ and Cd36-/- mice

Gene expression of Slc27a1, Fatp6, Fabp3, Got2, Acc2, Fasn, Scd1, Dgat1, Dgat2, and Abhd5 was assessed using quantitative real-time PCR. Gene expression is expressed relative to fed

Cd36+/+ mice. * indicates p < 0.05 compared within genotype, # indicates p < 0.05 compared to

Cd36+/+ mice of same feeding status; data were compared by 2-way ANOVA.

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Figure 21 - TG lipase activity in hearts from fed and fasted Cd36+/+ and Cd36-/- mice

Heart TG lipase activity was measured in fed and fasted Cd36+/+ and Cd36-/- mice (n=5).

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Fasted hLpL0 mice do not store lipids in the heart

If circulating FFAs are the source of heart TG stores during fasting, we would expect that loss of lipoprotein TG hydrolysis in the heart would not affect lipid droplet accumulation during fasting.

To test this, we fasted hLpL0 mice and compared them to LpLflox/flox littermates. After fasting hLpL0 mice had normal increases in plasma FFA concentration, an approximately 2-fold increase (Figure 22). hLpL0 mice tended to have slightly elevated TG in the fasted state, as has been reported (195). Plasma glucose fell 20% in both LpLflox/flox and hLpL0 mice. Surprisingly, hLpL0 mice did not accumulate cardiomyocyte TG during fasting (Figure 23, Figure 24).

Glycogen content was similar in all hearts (Figure 25, Figure 26).

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Figure 22 - Plasma biochemistry of fed and fasted LpLflox/flox and hLpL0 mice

Plasma FFA, TG, and glucose were measured 3-4 month old male LpLflox/flox (n=5) and hLpL0 mice (n=4) that were fed or fasted for 16 hours. * indicates p < 0.05, data were compared by

Student’s t-test.

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Figure 23 - Lipid measurements from hearts of fed and fasted LpLflox/flox and hLpL0 mice

Total lipids were extracted from hearts of LpLflox/flox and hLpL0 mice, and TG and FFA were measured. Lipid content was normalized to protein content. * indicates p < 0.05 compared within genotype, # indicates p < 0.05 compared to LpLflox/flox mice of same feeding status; data were compared by 2-way ANOVA.

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Figure 24 - ORO staining of hearts from fed and fasted LpLflox/flox and hLpL0 mice

Fed and fasted LpLflox/flox and hLpL0 mice heart sections (n=5) were stained with Oil Red O

(1000X).

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Figure 25 - PAS staining of hearts from fed and fasted LpLflox/flox and hLpL0 mice

Fed and fasted LpLflox/flox and hLpL0 mice heart sections (n=5) were stained with PAS reagent

(400X).

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Figure 26 - Glycogen content of hearts from fed and fasted LpLflox/flox and hLpL0 mice

Total glycogen was extracted from hearts of fed and fasted LpLflox/flox and hLpL0 mice and digested with amyloglucosidase, and free glucose was measured. Glucose content was normalized to tissue weight.

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We predicted that changes in gene expression in the fasting hLpL0 hearts would explain the lack of TG stores. mRNA levels of Atgl, Cpt1b, and Acox1 were reduced in both fed and fasted hLpL0 mice hearts (Figure 27), consistent with the reduction in FA oxidation in these hearts

(47). hLpL0 mice also had decreased expression of Plin2 and Plin5 compared to fasted

LpLflox/flox mice. Although these hearts do not have reduced FFA uptake, Cd36 mRNA was reduced in both fed and fasted hLpL0 hearts. Pdk4 expression was reduced in both fed and fasted hLpL0 hearts compared to LpLflox/flox hearts. Although glucose uptake is increased in hLpL0 hearts (47), fasted Glut1 mRNA levels were lower than controls and did not differ between fed and fasted hLpL0 mouse hearts. Glut4 was also reduced in both fed and fasted hLpL0 hearts.

Hearts from both fed and fasted hLpL0 mice had decreased diacylglycerol acyltransferase

(Dgat) 1 mRNA expression (Figure 28). Fasted hLpL0 mice had less Dgat2 mRNA expression compared to fasted LpLflox/flox mice. The rate-limiting enzyme for production of monounsaturated

FAs, stearoyl-CoA desaturase (Scd) 1, mRNA levels were decreased in hearts of fasted hLpL0 mice. Expression of Slc27a1 was increased in fasted floxed LpL mouse hearts, but not as much in fasted hLpL0 mouse hearts. FA transport protein 6 (Fatp6), Fabp3, and Got2 were all decreased in hearts of fasted hLpL0 mice compared to fasted floxed LpL mice. There were no differences in TG lipase activity between feeding conditions or genotype (Figure 29).

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Figure 27 - Gene expression in hearts from fed and fasted LpLflox/flox and hLpL0 mice

Gene expression of Atgl, Cpt1b, Acox1, Plin2, Plin5, Cd36, Lpl, Pdk4, Glut1, and Glut4 was assessed using quantitative real-time PCR. Gene expression is expressed relative to fed

LpLflox/flox mice. * indicates p < 0.05 compared within genotype, # indicates p < 0.05 compared to LpLflox/flox mice of same feeding status; data were compared by 2-way ANOVA.

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Figure 28 - Supplemental gene expression in hearts from fed and fasted LpLflox/flox and hLpL0 mice

Gene expression of Slc27a1, Fatp6, Fabp3, Got2, Acc2, Fasn, Scd1, Dgat1, Dgat2, and Abhd5 was assessed using quantitative real-time PCR. Gene expression is expressed relative to fed

LpLflox/flox mice. * indicates p < 0.05 compared within genotype, # indicates p < 0.05 compared to LpLflox/flox mice of same feeding status; data were compared by 2-way ANOVA.

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Figure 29 - TG lipase activity in hearts from fed and fasted LpLflox/flox and hLpL0 mice

Heart TG lipase activity was measured in fed and fasted LpLflox/flox and hLpL0 mice (n=5)

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Plin2 and Plin5 protein expression is not affected by cardiac lipid stores

A reduction in PLIN2 prevents lipid accumulation in the liver (128,130) and we had noticed a reduction in Plin2 mRNA. For this reason we measured PLIN2 protein in the hearts of fed and fasted mice. Fasting increased PLIN2 protein in Cd36+/+ mice, and unexpectedly also in Cd36-/- mouse hearts (Figure 30). Band density measurement indicated about a 40-50% increase in

PLIN2 after fasting in both Cd36+/+ and Cd36-/- mice (Figure 31). Because PLIN5 blocks TG lipolysis (136-138), a lack of change in mRNA but reduced protein could allow more rapid degradation of stored TG. Therefore, we measured Plin5 protein in the fed and fasted state.

PLIN5 protein levels were highly variable (Figure 30), but not significantly different between the fed and fasted states (Figure 31). Similar changes in both PLIN2 and PLIN5 protein were observed in hLpL0 mice (Figure 32, Figure 33).

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Figure 30 - Western blot of PLIN2 and PLIN5 in hearts of fed and fasted Cd36+/+ and

Cd36-/- mice

PLIN2 and PLIN5 content was assessed with western blotting in hearts of Cd36+/+ and Cd36-/- mice.

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Figure 31 - Band density analysis of PLIN2 and PLIN5 western blots in hearts of fed and fasted Cd36+/+ and Cd36-/- mice

PLIN2 and PLIN5 protein content in hearts of Cd36+/+ and Cd36-/- mice. Protein was quantified by band density measurements of the western blot. Band densities were normalized to β-actin content within each sample. Data are expressed as relative amount compared to fed Cd36+/+ mice. * indicates p < 0.05, data were compared by Student’s t-test (n=4).

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Figure 32 - Western blot of PLIN2 and PLIN5 in hearts of fed and fasted LpLflox/flox and hLpL0 mice

PLIN2 and PLIN5 content was assessed with western blotting in hearts of LpLflox/flox and hLpL0 mice.

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Figure 33 - Band density analysis of PLIN2 and PLIN5 western blots in hearts of fed and fasted LpLflox/flox and hLpL0 mice

PLIN2 and PLIN5 protein content in hearts of LpLflox/flox and hLpL0 mice was quantified by band density measurements of the western blot. Data were normalized to β-actin content within each sample and expressed as relative amount compared to fed LpLflox/flox mice. * indicates p <

0.05, data were compared by Student’s t-test (n=4).

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Blocking circulating TG degradation prevents cardiac lipid accumulation

Because of the surprising observation that hLpL0 mouse hearts did not accumulate TG during fasting - a result suggesting that circulating TG are the primary substrate for cardiac lipid accumulation - we used a drug that blocks lipolysis of circulating TG. Mice were administered

P407 and then fasted for 16 hours. Plasma TG increased 10-fold in the P407 treated mice, indicating a complete block of TG uptake (Figure 34). Surprisingly, plasma FFA and glucose concentrations were also higher in the P407 treated mice. Next, we looked at the heart lipids. As we found in the hLpL0 hearts, cardiac FFA were not affected by P407 treatment, but P407- treated mice had 60% less TG than PBS treated mice (Figure 35). There was decreased Oil Red

O staining of neutral lipids in P407-treated mouse hearts compared to PBS-treated mouse hearts

(Figure 36).

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Figure 34 - Plasma biochemistry of fasted PBS and P407-treated mice

Plasma FFA, TG, and glucose in 3-4 month old C57/BL6 male mice (n=5) that were injected intraperitoneally with 1 mg/g P407 or an equivalent volume of PBS. * indicates p < 0.05, data were compared by Student’s t-test.

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Figure 35 - Lipid measurements from hearts of fasted PBS and P407-treated mice

Total lipids were extracted from hearts of fasted PBS and P407 treated mice (n=5), and TG and

FFA were measured. Lipid content was normalized to heart weight. * indicates p < 0.05, data were compared by Student’s t-test.

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Figure 36 - ORO staining of hearts from fasted PBS and P407-treated mice

Fasted PBS and P407- treated mice heart sections (n=5) were stained with Oil Red O (1000X).

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We then measured uptake of circulating FFA in control and P407-treated mice. Plasma turnover of the label was identical in the control and treated mice (Figure 37). Heart uptake of the FFA was greater in the lipolysis-inhibited mice (Figure 38), consistent with a greater reliance of the heart on FFA than TG. Liver uptake of the label did not differ between groups. Next, we measured uptake of circulating glucose in control and P407-treated mice. Plasma turnover of the labeled glucose in plasma was the same in both groups of mice (Figure 39). Uptake of plasma glucose tended to be increased in P407-treated mice hearts (p=0.08) and livers (Figure 40). Total glycogen content was not changed between groups (Figure 41, Figure 42).

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Figure 37 - Plasma disappearance of [3H]-oleate in fasted PBS and P407-treated mice

Plasma disappearance of the [3H]-oleate was measured at 30 seconds, 1 minute, 3 minutes, and 5 minutes after injection (n=4-5).

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Figure 38 - Heart and liver uptake of FFA in fasted PBS and P407-treated mice

Cardiac and hepatic FFA uptake were assessed using [3H]-oleate (n=4-5). * indicates p < 0.05, data were compared by Student’s t-test.

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Figure 39 - Plasma disappearance of [14C]-2-deoxyglucose in fasted PBS and P407-treated mice

Plasma disappearance of the [14C]-2-deoxyglucose was measured at 2 minutes, 30 minutes, and

60 minutes after injection (n=4-5).

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Figure 40 - Heart and liver uptake of glucose in fasted PBS and P407-treated mice

Cardiac and hepatic glucose uptake were assessed using [14C]-2-deoxyglucose (n=4-5).

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Figure 41 - Glycogen content of hearts from fasted PBS and P407-treated mice

Total glycogen was extracted from hearts of fasted PBS and P407-treated mice and digested with amyloglucosidase, and free glucose was measured. Glucose content was normalized to tissue weight. (n=4-5).

Figure 42 - PAS staining of hearts from fasted PBS and P407-treated mice

Fasted PBS and P407-treated mice heart sections (n=5) were stained with PAS reagent, indicating glycogen content (400X).

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Lipid metabolism genes tended to be increased with P407 but not all increases reached statistical significance. Cpt1b, Acox1, and Lpl were increased in P407-treated mice while increases in Atgl,

Plin2, Plin5, and Cd36 were less robust (Figure 43). Pdk4 expression was not affected by P407- treatment, but Glut1 and Glut4 were both increased in hearts of P407-treated mice. P407 treated fasted mice had increased heart expression of Fabp3 and tended to have increased expression of

Fatp6 (Figure 44). There was a 30% increase in TG lipase activity in fasted mice treated with

P407 (Figure 45).

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Figure 43 - Gene expression in hearts from fasted PBS and P407-treated mice

Gene expression of Atgl, Cpt1b, Acox1, Plin2, Plin5, Cd36, Lpl, Pdk4, Glut1, and Glut4 was assessed using quantitative real-time PCR (n=4-5). * indicates p < 0.05, data were compared by

Student’s t-test.

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Figure 44 - Supplemental gene expression in hearts from fasted PBS and P407-treated mice

Gene expression of Slc27a1, Fatp6, Fabp3, Got2, Acc2, Fasn, Scd1, Dgat1, Dgat2, and Abhd5 was assessed using quantitative real-time PCR (n=4-5). * indicates p < 0.05, data were compared by Student’s t-test.

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Figure 45 - TG lipase activity in hearts from fasted PBS and P407-treated mice

Heart TG lipase activity was measured in fasted mice treated with either PBS or P407 (n=4-5). * indicates p < 0.05, data were compared by Student’s t-test.

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Cardiac TG accumulation during fasting does not impair heart function

Cardiac TG accumulation has been postulated to cause toxicity and to reduce heart function

(182,263). To determine if TG accumulation during fasting affects cardiac function, we fasted mice overnight and measured FS before and after administration of isoproterenol. We used young mice – 3-4 months – prior to the development of severe heart dysfunction that occurs in the hLpL0 mice (47). Basal FS was similar (~40%) in fasted Cd36+/+ and Cd36-/- mice.

Isoproterenol injection resulted in a maximum FS of 70% in fasted Cd36+/+ hearts and 60% in

Cd36-/- hearts (Figure 45, Figure 46). Basal FS tended to be greater in LpLflox/flox mice (56%) compared to hLpL0 mice (47%), but LpLflox/flox and hLpL0 mice had a similar maximal increase to 70% FS upon isoproterenol stimulation (Figure 46, Figure 47).

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Figure 46 - Basal and isoproterenol-stimulated fractional shortening in fasted Cd36+/+ and Cd36-/- mice

3-4 month old male Cd36+/+ (n=10) and Cd36-/- (n=9) mice were fasted for 16 hours and then injected intraperitoneally with 1.5 mg/kg isoproterenol. Cardiac function was assessed with M- mode echocardiography and fractional shortening was measured before and after injection of isoproterenol. * indicates p < 0.05, data were compared by Student’s t-test.

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Figure 47 - M-mode images of basal and isoproterenol-stimulated fractional shortening in fasted Cd36+/+ and Cd36-/- mice

Representative M-mode echocardiography images are pictured.

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Figure 48 - Basal and isoproterenol-stimulated fractional shortening in fasted LpLflox/flox and hLpL0 mice

3-4 month old male LpLflox/flox (n=7) and hLpL0 (n=8) mice were fasted for 16 hours and then injected intraperitoneally with 0.3 mg/kg isoproterenol. Cardiac function was assessed with M- mode echocardiography and fractional shortening was measured before and after injection of isoproterenol. * indicates p < 0.05, data were compared by Student’s t-test.

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Figure 49 - M-mode images of basal and isoproterenol-stimulated fractional shortening in fasted LpLflox/flox and hLpL0 mice

Representative M-mode echocardiography images are pictured.

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Discussion

In this series of experiments, we studied the source of FAs required for heart lipid droplet formation. We first found that hearts from Ppara-/- mice did not accumulate lipid droplets, despite their increase in plasma FFA and reduced FA oxidation. FFA uptake into the heart may be mediated by several transporters and non-receptor-mediated movement of FFA across the membrane (254,255). The FA transporter CD36 facilitates a major fraction of the albumin-bound

FFA uptake into cardiac cells (264) and also FFA derived from LpL hydrolysis of VLDL (30).

Loss of either CD36 or LpL reduced lipid droplet stores. Since CD36 is downstream of LpL (30), we then used a chemical inhibitor of lipolysis to confirm that defective lipolysis in the presence of continued FFA uptake could prevent heart TG accumulation. In agreement with our conclusion, albumin-bound FFA were shown to be more likely to be oxidized and chylomicron

TG-derived FFA more likely to be esterified (265). Our data contrast with studies in isolated hearts, in which supply of excess FFAs can drive lipid droplet formation (266,267). However, in vivo the majority of circulating FAs are esterified as TG and phospholipid (85-90%). Thus it would not be surprising that they also are a major supplier of heart FFA for TG storage. Finally, we showed that TG accumulation during overnight fasting did not impair or improve stimulated cardiac function.

The PPAR family of transcription factors control lipid uptake, oxidation, and storage.

The family members - PPARα, PPARβ/δ, and PPARγ - have overlapping transcriptional control of lipid metabolism gene expression. Two mouse models of heart specific PPAR overexpression, the MHC-PPARα and MHC-PPARγ transgenic mice, have increased lipid uptake, , increased neutral lipid storage, and increased lipid oxidation associated with cardiac dysfunction (202,247).

Although Ppara-/- mice have increased fasting concentrations of plasma FFA (268), the

109 accumulation of heart lipids during fasting had not been assessed. We hypothesized that Ppara-/- mice, which have reduced FA oxidation in the heart (166), would accumulate more lipids during a prolonged fast. We found just the opposite; TG accumulation was drastically reduced. Was this due to a defect in lipid droplet production, greater lipolysis of the stored TG, or reduced lipid uptake into these hearts? Since lipid uptake is upstream of lipid storage and turnover, it seems most likely that a deficiency of lipid uptake would precede any intracellular metabolic abnormalities.

Several mouse models do not accumulate cardiac lipids during fasting. Mice deficient in

HSL, an intracellular TG and diacylglycerol lipase, do not accumulate cardiac lipids during fasting, presumably due to the reduced ability of the adipose tissue to lipolyze stored TG and release FFA (226). HSL knockout mice have lower fasting plasma FFA and TG concentrations and decreased heart FFA uptake (226). Conversely, cardiomyocyte-specific HSL overexpressing mice also do not accumulate lipids after prolonged fasting because of rapid lipid droplet turnover

(207). Ppara-/- mice did not have an increase in ATGL expression; in fact, mRNA levels of

ATGL were dramatically reduced.

Another modulator of intracellular TG lipolysis is the lipid droplet protein PLIN5.

Recently, Plin5-/- mice were described to have defective storage of lipids in the heart (136);

PLIN5 is responsible for regulating ATGL activity, thus with PLIN5 deficiency intracellular lipase activity is constantly turned on. Plin4 deletion resulted in a similar phenotype due to a consequent decrease in Plin5 expression (133). Overexpression of PLIN5 increased heart TG content (137,138). Plin2-/- mice had decreased lipid accumulation in the liver and may play a similar role in promoting lipid accumulation (128). As others have reported (169), we observed

110 that Plin2 and Plin5 mRNA were dramatically reduced in Ppara-/- mice hearts; these genes are canonical PPARα target genes. Therefore, defects in lipid droplet formation in Ppara-/- mice could be due to lack of PLIN2 or PLIN5 or, more likely, the reductions in PLIN2 and PLIN5 are secondary to reduced PPARα activation. Surprisingly, intracellular TG lipase activities were dramatically increased in Ppara-/- mice – perhaps due to decreased Plin2 and Plin5 expression.

Others have demonstrated a fasting-induced increase of TG lipase activity in the heart of wild- type animals (143); we did not find this to be the case.

mRNA levels for Cd36 and Lpl were both markedly reduced in the Ppara-/- mice.

Therefore, we fasted mice that were deficient in CD36 or heart LpL. Cd36-/- mice have decreased cardiac FFA uptake and decreased VLDL-TG uptake, whereas hLpL0 mice have normal to increased cardiac FFA uptake and decreased VLDL-TG uptake (30,80). Since the concentration of circulating FFA increases during fasting, and mice that do not mobilize adipose lipid stores during fasting do not accumulate cardiac lipids (226), we hypothesized that FFA are the primary substrate for lipid accumulation in the hearts of fasted mice. We expected that Cd36-/- mice would not form lipid droplets after an overnight fast, but hLpL0 mice would form lipid droplets.

As expected, fasted Cd36-/- mice failed to accumulate lipids in the heart. Surprisingly, the hLpL0 mice also failed to accumulate cardiac lipids.

We still believed that FFA were driving this process, so we hypothesized that the lipid droplet was not stabilized and was turned over more rapidly in hLpL0 mice. hLpL0 mice have decreased PPARα target gene expression (195,217), so we suspected that lipid droplet proteins

PLIN2 and PLIN5 were also reduced. We found that Plin2 mRNA levels were reduced in both the fed and fasted state. However, Plin5 mRNA was normal. As there is often a mismatch

111 between Plin2 and Plin5 message and protein content, we assayed protein with Western blotting.

Protein expression in the heart were comparable for each genotype, and fasting increased heart

PLIN2 modestly in all genotypes. PLIN5 was comparable between control and hLpL0 mice with both feeding conditions. Finally, we measured heart intracellular TG lipase activity and there was no difference between LpLflox/flox mice and hLpL0 mice. Thus, we concluded that despite decreased PPARα target gene expression, it was unlikely that a defect in storage was preventing lipid accumulation in hLpL0 mice.

We then posited that VLDL-TG, and not FFA, were the source of the cardiac lipid for droplet accumulation observed in fasting. Indeed, cardiac LpL activity is increased after an overnight fast, suggesting an important role for circulating TG in supplying the heart with lipids

(269,270). To test this hypothesis, we treated mice with P407, a detergent that interferes with lipolysis of circulating lipoproteins. P407 treatment dramatically reduced cardiac lipid accumulation, but did not reduce PPARα target gene expression. Furthermore, P407-treated mice had increased FFA uptake, yet still failed to form lipid droplets. However, the decrease in lipid droplet formation could be the result increased intracellular TG lipase activity; it is unclear why intracellular TG lipase activity was increased in the P407 treated mice. We observed that P407 treatment and fasting together dramatically increased plasma FFA; plasma FFA concentrations in

P407 treated mice were 5-fold greater than those in plasma of non-treated fasting mice. This increase may be partially due to the partitioning of plasma FFA onto VLDL particles (Figure 50).

We concluded that TG, and not FFA, are the primary substrate for cardiac lipid accumulation observed in fasting, and that PPARα is necessary, but not sufficient, for cardiac lipid accumulation.

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Figure 50 - Distribution of plasma FFA in lipoprotein fractions

Plasma from PBS and P407-treated 16-hour fasted mice (n=4-5) was fractionated with density ultracentrifugation. 1.006 g/mL corresponds to the VLDL fraction; 1.12 g/mL corresponds to the

LDL fraction; 1.34 g/mL corresponds to the HDL fractions; >1.34 g/mL refers to the albumin/FFA fraction. Counts of [3H]-oleate were measured in each lipoprotein fraction and reported as percentage of total counts in plasma. * indicates p < 0.05. Data were compared by

Student’s t-test.

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In these studies, we focused on lipid uptake pathways that modulate TG accumulation in the hearts of fasted animals. As the heart is a dynamic organ that possesses the ability to use multiple substrates, we asked whether hearts in fasted animals that have defective lipid uptake might have changes in glucose utilization. Mice that have decreased lipid uptake in cardiomyocytes tend to increase glucose uptake and catabolism (47,271). We suspected that this might be happening in mice that failed to store lipid droplets in the cardiomyocytes. Heart gene expression of Pdk4, a negative regulator of glucose oxidation, was increased in all fasted mice, but to a lesser degree in mice that had deficiencies in lipid uptake. This likely indicates an increased reliance on glucose oxidation, i.e. pyruvate conversion to acetyl-CoA. Although gene expression of the glucose transporters GLUT1 and GLUT4 was variable, it has been previously demonstrated that Ppara-/- mice (214), Cd36-/- mice (57), and hLpL0 mice (47) all have increased cardiac glucose uptake. P407-treated fasting mice, which did not accumulate TG in cardiomyocytes, tended to have increased glucose uptake. However, glycogen content of hearts was similar for all genotypes, regardless of feeding status. We should note that others have reported increased heart glycogen with fasting (272).

We would have liked to assess where the albumin-bound FFA from adipose lipolysis and the FFA from LpL’s action on circulating TG were going once they entered the cardiomyocyte.

Different pools of circulating lipids, either FFA or TG, may enter the cardiomyocyte and be immediately oxidized, or they may be esterified as TG and later oxidized, or some combination of the two. FA turnover in the heart is very rapid with hydrolysis of much the pool of FAs within

10 minutes (47). Thus, assessing the residual FAs from labeled FFA or VLDL-TG is challenging. There are a number of elegant studies that have used isolated perfused hearts to track uptake, oxidation, and esterification of labeled TG in chylomicrons and VLDL

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(185,263,273,274). However, methods to study tracer uptake and oxidation in vivo and not in perfused hearts are not available. It is likely that LpL-mediated accumulation of heart TG during fasting is reflective of the much greater amount of circulating lipids that reside in TG particles.

The total amount of albumin-bound FFA might be insufficient to alone promote heart lipid accumulation during fasting. In support of the importance of TG as a source of heart lipids, lipotoxic mice that have excessive cardiac lipid accumulation are cured with cardiac-specific

LpL knockout (217). Furthermore, it might also be that FAs from different sources diverge in their intracellular fate. In support of this, one study of isolated perfused hearts found that FFAs were primarily oxidized, while FAs from chylomicrons divided equally into oxidation and cellular storage (265).

We assessed intracellular TG lipase activity to determine whether increased turnover of lipid droplets might explain why we did not see fasting-induced cardiac lipid accumulation in

Ppara-/- mice, Cd36-/-, hLpL0, and P407-treated mice. Surprisingly, we found there was dramatically increased TG lipase activity in Ppara-/- mice, despite the reduction in Atgl mRNA.

The deficiency of Plin2 and Plin5 that others and we have observed (169) in these mice could also indicate greater TG turnover. We also observed an increase in heart TG lipase activity in fasted mice treated with P407. However, there were no differences in heart TG lipase activity between Cd36-/- or hLpL0 mice and wild-type or floxed controls.

Although our data and others implicate LpL and CD36 as the primary players in cardiac lipid accumulation, it is possible that mice deficient in these proteins will have a reduction in other lipid transporters or de novo lipogenesis. We assessed mRNA expression of a number of

115 other genes and found that some of these genes had increased expression, but this was not sufficient to restore TG lipid droplet accumulation.

Lipid droplet accumulation is thought to occur due to an imbalance between lipid uptake and oxidation. This involves PPARα driven expression of LpL and CD36, allowing increased uptake of circulating lipoprotein TG (Figure 51). Whether TG accumulation is toxic is unresolved (229). Lipid droplets are found in hearts of patients with diabetes and metabolic syndrome (197-199) and in hearts of high fat diet-fed rodents and genetically altered mice (182).

Lipotoxicity can occur when ceramides, diacylglycerol, or other lipid species can alter cardiac cell signaling, disrupt membrane function, or cause apoptosis (275). We measured ceramides in hearts of fasted wild-type mice and did not observe an increase in ceramides coinciding with increased TG content (Figure 52). Excessive FFA oxidation has been proposed to lead to mitochondrial dysfunction, apoptosis and heart failure (190). For this reason, we tested whether increased stored lipid would reduce stimulated heart function. It did not. We found that short- term starvation and consequent TG accumulation did not decrease cardiac output.

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Figure 51 - Pathways of Lipid Uptake Leading to Lipid Droplet Formation

PPARα drives lipid accumulation by regulating transcription of lipid uptake proteins LpL and

CD36. Although both LpL and CD36 are required for cardiac lipid accumulation, LpL mediated lipolysis of lipoprotein TG is required for lipid droplet accumulation.

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Figure 52 - Ceramide measurement in fed and fasted wild-type mice

Individual ceramide species and total ceramides were measured from hearts of fed and 16-hour fasted wild-type mice (n=9). * indicates p < 0.05 compared to fed mice. Data were compared by

Student’s t-test.

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Although several models of lipid-induced heart dysfunction have massively increased amounts of heart TG, others have increases similar to that we found with fasting. ATGL knockout mice have 20 times the amount of cardiac lipids at 12 weeks of age and dramatically decreased heart function leading to 50% mortality by 18 weeks of age(127). However, two groups recently reported that PLIN5 overexpression and a 3-10-fold increase in cardiac TG led to only mild heart dysfunction (137,138). Less dramatic increases in TG are occasionally associated with heart dysfunction, however, in those hearts DAGs and ceramides also are increased (244).

We have previously reported a model of 50% increased cardiac TG in the fed state that is not associated with decreased cardiac function – the MHC-DGAT1 mouse model (229). In fact, this transgene reduces toxicity in other models without changing TG concentrations, but reducing heart ceramide 20% (229). In another model, transgenic MHC-PPARγ mice, a similar 2-3 fold increase in myocardial TG but coupled to increased DAG and ceramide was associated with more than a 50% reduction in FS (200,202,276). The effects of diets and diabetes on heart TG and other lipids and cardiac function are less clear. Three weeks of high fat diet feeding increased heart TG by 2-3 fold, but a 20% decrease of FS was only observed after 20 weeks high fat diet feeding (230,277). Streptozotocin-diabetic mice had a 50% increase in cardiac TG after

12 weeks associated with a 20% reduction of FS (278). Ob/ob and db/db mice all have varying degrees of increased cardiac TGs (from 2-fold to 4-fold) associated with heart dysfunction depending on the duration of the study (248,279). Concentrations of ceramides and DAGs were often not measured in these models. Increased heart TG content is sometimes, but not always, a hallmark of increased accumulation of others lipids, many of which are harmful to the heart.

Storage of TGs in the heart may be an adaptive response to decreased energy intake during starvation. However, it also occurs in the setting of pathological conditions including

119 obesity, diabetes, and non-ischemic heart failure (280,281). Moreover, one report has suggested that heart lipid droplet accumulation occurs post-ischemia and is associated with more tissue damage (45). As we note above, lipid droplet accumulation is sometimes associated with heart dysfunction and also increased concentrations of potentially toxic lipids such as ceramides and

DAGs, but in other situations TG storage alone does not lead to heart dysfunction (182).

Whether the association between heart TG and function in humans is due to TG or other accumulated lipids is not obvious. It should be noted that severe heart failure in humans leads to reduced heart TG, but an increase in potentially toxic ceramides and DAGs (282). Our data show that lipid droplets, at least those that occur during fasting, do not affect heart function.

In summary, our studies show that LpL-lipolysis of TG is required to store lipid in the heart of a fasting mouse. This study further confirms previous work showing that hearts are active organs in TG metabolism. We now show that the very high level of cardiac LpL expression is involved not only in acquisition of circulating FAs for energy, but also for allowing storage.

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Chapter 4: Conclusions and future directions

These studies are a small contribution to the tremendous body of work that demonstrates the critical role of lipid metabolism for both the normal function of the heart and in the setting of cardiac dysfunction. In particular, they highlight the importance of the uptake of circulating TG for cardiac metabolism during starvation (Table 5). Several, but not all, mouse models of impaired cardiac lipid uptake develop cardiac dysfunction (47,98,99,239). However, prevention of excess lipid uptake and oxidation has been demonstrated to have beneficial effects in a number of in vitro studies (190). Since this work suggests that uptake of circulating TG may be more important for driving lipid accumulation, this implies that decreasing TG uptake may be beneficial in some settings of cardiac dysfunction. Indeed, previous work has shown that Vldlr-/- mice have decreased TG accumulation after myocardial infarction as well as improvements in markers of cardiac function (45).

In order to transition these findings into a clinically relevant model, future studies will assess the effects of the inhibition of cardiac TG accumulation after myocardial infarction. The animal models described – Cd36-/- mice, hLpL0 mice, and P407-treated mice - will be used to evaluate changes in heart function and toxic lipid accumulation in both acute and chronic models of myocardial infarction. In brief, animals would be surgically infarcted in the left anterior descending artery and allowed to recover; heart function would be evaluated by measuring fractional shortening. Mice that have reduced cardiac TG accumulation could be protected from development of heart dysfunction. Indeed, excessive FFA uptake and oxidation has been linked to dysfunction in several acute injury models (283). It is likely that the Cd36-/- mice, which have defects in both FFA and TG uptake, may have improved heart function after infarction. Mice that

121 only have a defect in TG uptake – the hLpL0 mice and P407 treated mice – still avidly obtain circulating FFA. This would point specifically to a role for circulating FFA, and not TG, in promoting cardiac lipotoxicity in acute settings. Moreover, a number of in vitro studies using isolated perfused hearts suggest that FFA are acutely toxic (263,283).

To the same end, these findings will be used to ameliorate cardiac lipotoxicity in a model of diet-induced obesity. The knockout mice – Cd36-/- and hLpL0 mice - would be made obese by

8-12 weeks of high-fat diet feeding, which has been shown to induce cardiac dysfunction (230).

Both mouse models should have improved cardiac function associated with reduced cardiac lipid accumulation. Indeed, deletion of either Cd36 or Lpl has been demonstrated to ameliorate cardiac lipotoxicity in a transgenic model (217,238). Similar studies may be conducted to assess whether inhibition of TG accumulation may also improve cardiac function in streptozotocin- diabetic mice.

Of most interest to these studies is developing a potential human therapy – for both the infarcted heart and for cardiac dysfunction associated with obesity. The most promising option is to inhibit VLDL-TG secretion via the use of ApoB or MTTP inhibitors, which reduce circulating

TG concentration (284,285). This is an attractive option as an ApoB antisense drug, mipomersen, is currently undergoing clinical trials (285). Since either ApoB or MTTP inhibition almost completely ablates VLDL-TG secretion, there should be no cardiac lipid accumulation in fasting mice. The next step is to pre-treat mice before surgical infarction to determine whether cardiac lipotoxicity is ameliorated and whether there are any changes in heart function. Finally, the drug will be administered in a rodent model of chronic infarction to determine whether reduction of

TG might ameliorate the deterioration of heart failure.

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Finally, future developments in nuclear imaging and magnetic resonance spectroscopy should elucidate the differential fates of FFA that come from albumin-FFA versus LpL-lipolysis of circulating TG in vivo. These studies may prove useful in preventing excessive lipid accumulation associated with decreased cardiac function. Current state of the art imaging and spectroscopy is better developed in skeletal muscle or liver than in heart (286-288). Future developments may allow the investigation of the intracellular fates of circulating FFA versus TG, as well as carbohydrate and amino acid metabolites.

Finally, it is unclear why the P407-treatment increased circulating glucose and FFA concentrations. It is likely that inhibiting LpL-hydrolysis of circulating TG triggers a stress response; this plasma concentration of stress hormones epinephrine, norepinephrine, and glucagon will be measured. These hormones stimulate increases in plasma FFA and glucose concentrations. This raises the interesting question of how the adrenal glands and pancreas sense energy balance, and suggests that LpL is critical for the regulation of stress hormone levels.

Tissue specific knockouts of LpL in the adrenal glands and pancreatic α-cells (which secrete glucagon) may result in increased catecholamines and glucagon. This would directly link plasma

TG concentration to the regulation of gluconeogenesis and WAT lipolysis and provide novel insight into the dysregulations in metabolic disease.

In conclusion, this thesis suggests future studies and potential clinical interventions targeting cardiac lipid metabolism that may play a critical role in the treatment of heart dysfunction. Addressing these additional questions will provide further insight into the specific role of circulating metabolites in cardiac physiology and pathology.

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Table 5 - Models of altered cardiac TG accumulation during starvation

Model Reference Heart TG Explanation

MHC-HSL (207) Decreased Increased IMTG lipolysis

Hsl-/- (226) Decreased Very low FFA/TG in circulation

Plin4-/- (133) Decreased Decreased Plin5 expression

Plin5-/- (136) Decreased Removal of lipolytic barrier

MHC-ATGL (289) Decreased Increased IMTG lipolysis

MHC-Plin5 (138) Increased Decreased ATGL activity

MHC-Plin5 (137) Increased Lipid droplet protected from ATGL

Ppara-/- (250) Decreased Decreased expression of Cd36, LpL; possibly decreased expression of Plin2, Plin5

Cd36-/- (250) Decreased Decreased LpL activity

hLpL0 (250) Decreased Decreased LpL activity

P407 treatment (250) Decreased Decreased LpL activity

124

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