THE REGULATION OF EICOSANOID BIOSYNTHESIS THROUGH PHOSPHOLIPID

REMODELING

by

SARAH ASHLEY MARTIN

B.S. Biochemistry, North Central College, 2010

A thesis submitted to the

Faculty of the Graduate School of the

University of Colorado in partial fulfillment

of the requirements for the degree of

Doctor of Philosophy

Pharmacology Program

2015

This thesis for the Doctor of Philosophy degree by

Sarah Ashley Martin

has been approved for the

Pharmacology Program

by

Tatiana Kutateldze, Chair

Donna L. Bratton

Kim A. Heidenreich

Raphael A. Nemenoff

Dennis R. Voelker

Robert C. Murphy, Advisor

Date 12/18/2015

ii Sarah Ashley Martin (Ph.D., Pharmacology)

The Regulation of Eicosanoid Biosynthesis Through Phospholipid Remodeling

Thesis directed by Professor Robert C. Murphy

ABSTRACT

Since the discovery of eicosanoids and their biological effect, prostaglandins and leukotrienes have been the targets of pharmacologic inhibition. Yet completely blocking bioactive lipid production has been plagued with complexities. In large part this is due to

the complex series of biochemical steps involved in regulating lipid mediators. Previous

work had shown that eicosanoid production could be regulated through the availability of

the precursor arachidonic acid (AA), specifically by inhibiting the reacylation of free AA back into an esterified form.

The work presented in this dissertation focused on two enzyme classes, the acyl‐

coenzyme A synthetase long‐chains (ACSL) and the lysophospholipid acyltransferases

[LPATs]. Two approaches were taken to assess the role of these enzymes in the esterification of AA into phospholipids. The first was a collaborative project that used an

inducible deletion of ACSL1 in mice, and the second used short‐hairpin RNAs to selectively

decrease membrane‐bound O‐acyltransferase 5 or lysophosphatidylcholine acyltransferase

3 (MBOAT5/LPCAT3), membrane‐bound O‐acyltransferase 7 or lysophosphatidylinositol

acyltransferase 1 (MBOAT7/LPIAT1), and ACSL4 in a murine monocyte cell line, RAW

264.7.

The inducible deletion of ACSL1 in mice led to a significant decrease of enzymatic

activity in cardiac tissue, mitochondria metabolism switching, left ventricle enlargement,

and significant acyl chain remodeling of the phospholipid pool. Analysis by liquid

chromatography‐ tandem mass spectrometry (LC‐MS/MS) revealed a specific change in a

glycerophospholipid (cardiolipin), which could be restored with diet. However, the diet was

iii insufficient to repair the functional changes in the mitochondria/left ventricle and the other phospholipid acyl compositions that were lost by the decrease in ACSL activity.

To further analyze lysophospholipid acyltransferase activity, a competitive enzyme assay was developed which used microsomal extracts incubated with multiple lysophospholipids and fatty acyl CoA esters in one reaction. Analysis by LC‐MS/MS revealed activity changes specific to the selective decrease of MBOAT5/LPCAT3and

MBOAT7/LPIAT1in the RAW 264.7 cell. The RAW 264.7 cells that contained shRNAs against

LPCAT3, LPIAT1, and ACSL4 were stimulated to produce eicosanoids. The targeted decrease of MBOAT5/LPCAT3revealed no effect on the levels of free AA or eicosanoid production.

ACSL4 depletion led to increases in the levels of free AA but no change in the production of eicosanoids. MBOAT7/LPIAT1reduction led to decreases in the levels of free AA and the production of eicosanoids after cells were stimulated. Taken together, the data presented in this dissertation suggests that phospholipid remodeling is critical in overall eicosanoid production by a cell, as well as a process operating to increase phospholipid molecular species diversity.

The form and content of this abstract are approved. I recommend its publication.

Approved: Robert C. Murphy

iv

To the strongest women I will ever know,

My great‐grandma, Elizabeth Bileddo

My grandmother, Prudence Martin

And my mother, Diane Martin

v ACKNOWLEDGEMENTS

I would to like to extend my gratitude to all of the individuals who have help me through the good and bad to find the light at the end of the tunnel. First and foremost to my family members, specifically my mother for all of the phone calls she had to sit through, and all of the support she has provided me throughout my entire life. In addition, my

grandmother, the strongest woman I have ever known. Without the contributions of these

two self‐less women, I would not have become the person that I am today.

I wish to recognize those individuals that encouraged me to persevere in the laboratory. Chris Johnson, Dr. Karin Berry, and Dr. Joseph Hankin, I thank you for your input

and support. You have all been invaluable to my training, and I would not be the scientist I

am now without your influence. I have to thank my closest Colorado friends, Dr. Emma

Murphy, Deborah Beckworth, and Dr. Michael Buendia. My day‐to‐day life was heavily

impacted by all of you, and I thank you for helping me grow into the person that I am today.

I would like to thank all of my committee members for their extensive input on my thesis

work.

Finally, I wish to recognize my mentor, Dr. Robert Murphy, for all of the hours he

has dedicated to my scientific training. My scientific ability has grown substantially because

of those countless hours, and I thank you for your willingness to sit and explain things to me

until I understood, which at times was significant. Your knowledge and passion for science

is extraordinary, and it has been a privilege to train with you.

vi TABLE OF CONTENTS

CHAPTER Page

I. INTRODUCTION 1

Arachidonic Acid and Phospholipase A2 1

The Discovery, Synthesis, and Activity of the Major Cyclooxyganse 7 Products

The Discovery of Leukotrienes 13

The Synthesis of Leukotrienes 13

The Biologic Role of Leukotrienes 18

The Development of Pharmacologic Intervention of Eicosanoids 20

Phospholipid Synthesis and Remodeling 23

Additional Modifications of Phosphatidylinositol 33

Thesis Goal 36

II. LIPID MASS SPECTROMETRY 37

Introduction 37

Techniques for the Ionization of Lipids 37

Mass Analyzers 42

Measuring Exact Mass by Mass Spectrometry 44

MALDI Imaging Mass Spectrometry 46

Tandem Mass Spectrometry 49

III. MEASUREMENT OF LYSOPHOSPHOLIPID ACYLTRANSFERASE 54 ACTIVITY USING SUBSTRATE COMPETETITION

Introduction 54

Experimental Procedures 55

Materials 55

vii Cell Culture and Microsomal Preparation 56

Single LPAT Assay 57

Dual Substrate Choice Assay 58

Liquid Chromatography and Mass Spectrometry 58

Results 61

NP‐HPLC‐MS/MS Detection and Quantitation of 61 Phospholipid Products

Determination of Enzymatic Parameters of the Dual 63 Substrate Choice Assay in Microsomes from RAW 264.7 Cells

The Measurement of Endogenous Microsomal Phospholipids 65 from the RAW 264.7 as a Means for Normalization

The Products of the Dual Substrate Choice Assay Reflects 67 the Presence of Many LPATs in RAW 264.7 Microsomes

Dual Substrate Choice Assay Comparison to Alternative 71 LPAT Assays

Discussion 73

IV. MEMBRANE BOUND O‐ACYLTRANSFERASE 5/ 76 LYSOPHOSPHATIDYLCHOLINE ACYLTRANSFERASE 3 ROLE IN ARARCHIDONIC ACID REGULATION

Introduction 76

Experimental Procedures 79

Materials 79

Cell Culture and Microsomal Preparation 80

Targeted Decrease of MBOAT5/LPCAT3 gene Expression 80 in RAW 264.7 Cells

RAW 264.7 Cell Stimulation and Eicosanoid Sample Preparation 81

LC‐MS/MS for Fatty Acids and Eicosanoids 82

Dual Substrate Choice Assay 82

LC‐MS/MS for Phospholipids 83

viii Results 84

Thimerosal Does Not Have the Same Affect on the RAW 264.7 Cells 84

Pioglitazone Treated RAW 264.7 Cells Upregulates 86 MBOAT5/LPCAT3

Activity of MBOAT5/LPCAT3 was Significantly and Specifically 88 Decreased in RAW 264.7 Cells Using a Targeted shRNA

The Depletion of MBOAT5/LPCAT3Does Not Lead to Changes in 92 Leukotriene Production

Discussion 94

V. THE AFFECT OF MEMBRANE BOUND O‐ACYLTRANSFERASE 7 AND 99 ACYL‐COENZYME A SYNTHETASE LONG‐CHAIN 4 ON ARACHIDONIC ACID AND LEUKOTRIEN PRODUCTION

Introduction 99

Experimental Procedures 101

Materials 101

qRT‐PCR in RAW 264.7 Cells 103

Dual Substrate Choice Assay and Microsomal Preparation 103

LC‐MS/MS for Phospholipids 104

RAW 264.7 Cell Stimulation and Eicosanoid Sample Preparation 105

LC‐MS/MS for Fatty Acids and Eicosanoids 106

Thin‐layer Chromatography and Incorporation of [3H] AA 106 into RAW 264.7 Cells

Results 107

Confirmation of MBOAT7/LPIAT1 shRNA Targeted Decrease in the 107 RAW 264.7 Cell

Targeted Decrease of MBOAT7/LPIAT1 Led to Decrease in 112 LTC4 Production

An Early Decrease in AA Release Led to a Decrease in 5‐LO 114 Products

ix Reduction in ACSL4 Led to Increases in AA Release but No Change 116 in 5‐LO Products

Targeted Decrease of MBOAT7/LPIAT1 Does Not Change other 118 Lands’ Cycle or Eicosanoid mRNA Levels

Loss of MBOAT7/LPIAT1 is Not Recovered by Adding Exogenous 120 AA or by Bypassing cPLA2α Activity

Molecular Species of PI Do Not Change with the Depletion of 122 MBOAT7/LPIAT1

Exogenously Added AA is Almost Entirely Recycled into PI, PC, 125 and PE in the Parental RAW 264.7 Cells

Targeted Analysis of AA Containing Global Phospholipid Pools 127 Do Not Change with MBOAT7/LPIAT1 Depletion

Discussion 128

IV. ACYL‐COENZYME A SYNTHETASE LONG CHAIN 1: 134 THE REGULATION OF PHOSPHOLIPIDS

Introduction 134

Experimental Procedures 137

Sample Preparation for Mass Spectrometry 137

Liquid Chromatography/Mass Spectrometry 138

Preparation and Analysis of Tissue for Matrix Assisted Laser 139

Desorption Ionization/ Imaging Mass Spec (MALDI‐IMS)

Results 140

Formation of Long‐Chain Fatty Acyl CoA Esters is Diminished 140 in the Heart when ACSL1 is Depleted

Total Mass of Cardiolipin Does Not Change but is Remodeled 140

Cardiolipin Content is Remodeled Throughout the Right and 143 Left Ventricle

Loss of DHA Containing Phosphatidylcholines in the ACSL1T‐/‐ 145

Two Major Molecular Species Lost with Depletion of ACSL1 in the 146 Heart Contained 22:6 or DHA

x

Phosphatidylcholine Containing DHA is Lost Throughout the 148 Right and Left Ventricle in the ACSL1T‐/‐

Phosphatidylinositol, Phosphatidylenthanolamine, and 150 Phosphatidylcholine Experienced Chain Remodeling with the Depletion of ACSL1 in the Heart

Cardiolipin Content is Recovered with High Levels of Linoleic but 153 No Recovery of Mitochondrial Function

Bis (Monoacylglycerol) Phosphate is Increased in the ACSL1T‐/‐ 155

Discussion 158

VII. CONCLUSIONS AND FUTURE DIRECTIONS 165

Summary of Major Findings

Chapter III 165

Chapter IV 165

Chapter V 166

Chapter VI 166

Future Directions 167

The Complex Role of Thimerosal 167

The Regulation of MBOAT5/LPCAT3 by RXR and the 168 LPCAT3T‐/‐ Mouse

ACSL4 in the Regulation of AA in the RAW 264.7 Cell 170

MBOAT7/LPIAT1 in the Regulation of AA Containing 170 Phosphatidylinositols in the RAW 264.7 Cell

MBOAT7/LPIAT1 in the Regulation of Mead Acid in the 171 RAW 264.7 Cell

MBOAT7/LPIAT1 in the Regulation of Phosphoinositides 172 in the RAW 264.7 Cell

Concluding Remarks 173

REFERENCES 175

xi LIST OF TABLES

TABLE Page

2‐1 Exact Mass and Natural Isotope Calculations for Common Organic Atoms. 46

3‐1 Multiple Reaction Monitoring Transitions for the Substrate Choice Assay. 60

5‐1 Semi‐quantitation of abundant AA containing phospholipids in 128 RAW 264.7 cells.

xii LIST OF FIGURES

FIGURE Page

1‐1 Biochemical synthesis pathway of the major biologically active prostanoids 11

1‐2 Biochemical synthesis pathway of leukotrienes. 17

1‐3 The de novo synthesis of phospholipids modified from D.E. Vance 26 and J.E. Vance, 2008.

1‐4 Summary of the known activities of the lysophospholipid acyltransferases 28

1‐5 Reacylation of lysophospholipids leads to incorporation of 32 polyunsaturated fatty acids and phospholipid asymmetry

1‐6 Phosphorylation states phosphatidylinositol. 35

2‐1 Scheme of the mechanism by which ions are transformed into the gaseous 40 by matrix assisted laser desorption ionization (MALDI).

2‐2 Scheme of the mechanism by which ions are transformed into the gaseous 41 by electrospray ionization.

2‐3 Scheme of the mechanism by which ions are selected for in the 44 quadrupole mass spectrometer.

2‐4 Workflow of how MALDI‐IMS (imagine mass spectrometry) is able 48 to obtain spatially distribution of lipids in a tissue.

2‐5 Tandem quadrupole experiments performed during the course 52 of this thesis work.

3‐1 Separation and quantitation of phospholipids 62

3‐2 Reaction conditions of the dual substrate choice acyltransferase assay. 64

3‐3 Endogenous microsomal phospholipids analyzed during the dual 66 substrate choice acyltransferase assay

3‐4 Acyl chain preference of RAW 264.7 microsomal acyltransferases 68

3‐5 Polar head group preference of RAW 264.7 microsomal acyltransferases. 69

3‐6 Effect of substrate competition on acyltransferase activity. 72

4‐1 Thimerosal inhibits MBOAT5/LPCAT3 and leads to increased leukotriene 78 production in human neutrophils.

xiii 4‐2 Thimerosal (Thim) has a dramatic affect on leukotriene production in 85 human neutrophils but does not have an effect on leukotriene formation in RAW 264.7

4‐3 Pioglitazone treatment for 24 hr leads to increased MBOAT5/LPCAT3 87 expression and decreases free AA and LTC4 levels.

4‐4 Phospholipid containing AA are reduced in microsomal extracts from 89 cells with an shRNA targeted against MBOAT5/LPCAT3.

4‐5 Effect of reduced expression of MBOAT5/LPCAT3 91 on acyltransferase activity.

4‐6 Phospholipid molecular species production during reduced 93 expression of MBOAT5/LPCAT3 in microsomes.

4‐7 Targeted decrease of MBOAT5/LPCAT3 in RAW 264.7 cells does change 94 leukotriene production.

5‐1 MBOAT7/LPIAT1 shRNA targeted cells have a decrease in mRNA 109 and enzymatic activity.

5‐2 Phospholipid molecular species production during reduced expression 111 of MBOAT7/LPIAT1 in microsomes.

5‐3 LTC4 production is reduced in RAW 264.7 cells that have decreased 114 MBOAT7/LPIAT1 expression when stimulated by ATP.

5‐4 Specific changes in AA release at early time points mirror later 117 reduced production in LTC4 and 5‐HETE in RAW 264.7 cells that have decreased MBOAT7/LPIAT1 expression.

5‐5 Targeted decrease of ACSL4 led to increases in freed AA after stimulation 119 but no change in leukotriene formation.

5‐6 Targeted decrease of MBOAT7/LPIAT1 did not lead to changes in 121 expression of other reacylation or eicosanoid enzymes.

5‐7 Deficiency in LTC4 production in the targeted decrease of MBOAT7/LPIAT1 123 could not be recovered with addition of exogenous AA or by non‐receptor mediated stimulus.

5‐8 Quantitation of PI molecular ions does not reveal any changes due to the 124 loss of MBOAT7/LPIAT1.

5‐9 Addition of exogenous [3H]‐AA is almost exclusively incorporated 126 into the phospholipid pool.

5‐10 The regulation of PI molecular species is a mixture of the reacylation 131

xiv pathway and the PI cycle.

6‐1 Tetra linoleic cardiolipin (18:2/18:2/18:2/18:2‐CL or m/z 1447.9) 137 is the most abundant cardiolipin species in mammalian cardiac muscle.

6‐2 Loss of ACSL activity in ACSL1T‐/‐ but no change in total 141 phospholipid measurement.

6‐3 The remodeling of acyl chain composition of cardiolipin in 142 ACSL1T‐/‐ heart tissue.

6‐4 MALDI imaging of cardiolipin in the ventricles from control and ACSL1T‐/‐ 144 mice shows global changes in acyl composition

6‐5 The remodeling of acyl chain composition of phosphatidylcholine in 147 ACSL1T‐/‐ heart tissue by LC‐MS/MS reveals major loss of DHA.

6‐6 MALDI imaging of PC in the ventricles from control and ACSL1T‐/‐ mice 149 shows global loss of 40:6 and 38:6‐PC and gain of 36:1 and 34:1‐PC..

6‐7 Semi‐quantitative changes in acyl composition of PC, PI, and PE in 151 the heart tissue of the ACSL1T‐/‐ compared to control animals reveals major changes in acyl chain composition.

6‐8 Cardiolipin shows significant changes in molecular species 155 with a high safflower oil.

6‐9 Safflower oil does repair most of the changes made to the 156 phospholipid molecular species in the ACSL1T‐/‐.

6‐10 Many BMP molecular ions are increased in the heart tissue of ACSL1T‐/‐. 157

6‐11 BMP molecular ions are also increased in the livers of ACSL1T‐/‐. 159

6‐12 High linoleate diet did not improve mitochondrial function in 160 Acsl1T‐/‐ hearts.

xv

CHAPTER I

BIOACTIVE LIPIDS

Arachidonic Acid and Phospholipase A2–

Arachidonic acid (AA) is the precursor for a family of bioactive lipid mediators that have been termed eicosanoids. This twenty‐carbon, four‐double bond, omega‐6 fatty acid is an essential fatty acid that is predominantly stored in phospholipids. The history behind discovering all of these characteristics of arachidonic acid took several decades of research to elucidate. In this section, we review the history of the structural characterization of arachidonic acid and how it was found to be a precursor to a large family of bioactive lipids.

The role of dietary fat was the topic of interest of Percival Hartley (Lister Institute,

London) for his doctorate of science degree from 1906 to 1909. During his studies of modified diets in animals, iodine was used to determine values to characterize the number of double bonds in the esterified fatty acids as well as how the number of double bonds found in lipids from tissues fluctuated with diet (1). To determine the structural characteristics of the different fatty acids, Hartley used lecithin from pig liver to obtain a high yield of fatty acids that contained a high number of double bonds (2). For the initial isolation of AA, Hartley went through an exhaustive protocol to first extract phospholipids from tissues, and then purified the free AA from the saponified mixture of fatty acids in the work he published in 1909 (2).

Hartley’s first step utilized differential solubility extraction techniques developed earlier by Roaf and Edie in 1905 to isolate lecithin and other similar molecules (3). Once the phospholipid fraction was precipitated from an ethanol extraction solution, it was subjected to base hydrolysis to free all of the acyl chains which were esters contained in lecithin. Hartley, knowing that lead salts of unsaturated acids were soluble in ether, was

1

able to separate the unsaturated fatty acids from other fatty acids. The previously mentioned initial purification steps enriched the sample in fats that contained unsaturated fatty acids as evident by the iodine value increasing from 97 to 144, and then to a range of

165 to 175. To remove cholesterol and unsaponified matter, potassium salt forms of the fatty acids were extracted with ether. The mean molecular weight of the extracted matter was determined to be between 308 and 312 using titration of weighed extracted lipid.

To further purify the unsaturated fatty acids, Hartley then used vacuum distillation of the methyl esters and identified four unique fractions. Fractions 3 and 4 had boiling temperature values of 180‐190°C, iodine values of 250, and contained fatty acids with molecular weights calculated to be 313 Da. The unsaturated acids in fraction 3 and 4 were then reacted with molecular bromine under acidic conditions to form a white precipitate.

To separate hexabromide compounds from suspected octobromide compounds, the precipitate was resuspended in benzene. Octobromide compounds are not soluble in benzene, which was the key step in the purification of AA. Using the same initial extraction protocol, he subjected the fatty acids in fraction 3 and 4 to alkaline permanganate oxidation to form hydroxy acids. He performed elemental composition analysis on the octobromo and hydroxy acid products and found a twenty carbon acid. His conclusion was that he had identified a C20H32O2 fatty acid, similar to arachidic acid, however, this new fatty acid contained four double bonds (2). The extracted fatty acid constituted approximately 10% of the total fatty acids that were derived from fractions of lecithin and similar complex structures (2). Since the saturated 20‐carbon fatty acid that had already been named as arachidic acid, the name arachidonic acid (AA) was first coined and published in 1913 (4) for the C20H32O2 fatty acid originally described by Hartley.

Additional groups supported the observation made by Hartley, and described a 20‐ carbon, 4‐double bond fatty acyl group in lecithin (phosphatidylcholine), kephalin or

2

cephalin (mixture of phosphatidylethanolamine and phosphatidylserine), neutral fat, and the free acid form in many different tissues from animal studies (5‐11). Isolation of large quantities of AA proved to be difficult due to the exhaustive protocol that was originally developed by Hartley. To further characterize the structure of AA, Laurence Wesson utilized the extraction of the highly insoluble octabrominated compound (7). After isolation of the octobromoarachidic acid, he performed a debromination reaction using a copper coated zinc dust. The AA was readily dehalogenated to a tetraunsaturated ethyl ester or free acid. Wesson described AA as a colorless liquid with a tenacious fishy odor

(7). He could not confirm the position of the four double bonds due to the extensive chemistry that had to be performed during isolation. However, his method was used in many experiments up until the 1950’s when chromatographic separation began to replace

Wesson’s technique for the isolation of AA.

The work to determine the double bond positions of AA was taken up by investigators in the United States, Japan, and England. They used ozonolysis and various oxidation techniques that generated chained‐shorten products. These products were isolated and compared to known short chain acids to rebuild the structure of AA. Toyama and Tsuchiya from Japan were the first to publish their studies and suggested the double bond position of AA as a Δ4, 8, 12, 16‐eicosatetraenoic acid (12). But their structure was not supported by data reviewed by Bull that suggested no double bonds could be present before carbon‐5 or ‐6 because the theoretical iodine value for AA was obtained, and any bonds present before the carbon‐5 position would have altered the iodine value (13).

In 1940, work performed by Shinowara and Brown at The Ohio State University led to the misidentification of the four double bonds as a Δ6, 9, 12, 15‐eicosatetraenoic acid

(14). In that same year, Dolby and colleagues in London used aqueous alkaline permanganate solutions to oxidize then cleave at each double bond converting the vinyl

3

carbon atom to a carboxylate group (15). They identified five chained shorten products: oxalic, succinic, pentanoic, hexanoic, and pentanedioic acid. With the identification of fragments common to linoleic acid, knowledge that the double positions were in a skipped pattern, and the double bonds began at carbon 5 or 6 their data supported the double bond configuration of Δ5, 8, 11, 14‐eicosatetraenoic acid. However, the authors stated they could not detect all of the product molecules necessary to confirm the structure, specifically they were not able to find malonic acid (15). Additional information was obtained after the initial Dolby paper to show that there was little conjugation (less than 5%) in double bonds of AA (16).

Ultimately, Brown and colleagues (17) performed additional ozonolysis and oxidization with potassium permanganate two years after publishing the wrong structure.

Their initial experiments yielded insufficient product formation and led to incorrectly identifying some acids as a product that form from the ozonolysis of arachidonic acid (17).

Using ozonolysis followed by melting point comparative analysis they identified hexanoic, acetic, pentanedioic, butanedioic, and malonic acids. The permanganate oxidation products that they identified were hexanoic, pentanedioic, butanedioic, and oxalic acids. These product identifications led them to the correct double bond configuration of Δ5, 8, 11, 14‐ eicosatetraenoic acid (17), which supported the work published by Dolby (15).

All of the structural work that had been performed up to the early 1940s did not address the double bond stereochemistry. In part, information about this important structural detail of AA came from biological investigations. Data was published in 1951 that showed supplementation of trans geometric isomers of linoleic acid could not recover the cession of growth and skin issues observed in animals fed diets without omega‐6 fatty acids (18). Using infrared spectroscopy, work with monounsaturated and diunsaturated fatty acids revealed a unique absorption in the spectra range from 970‐990 cm‐1 that was

4

characteristic of the cis‐trans carbon‐carbon double bond bending (19,20). The initial experiments performed by Jones and colleagues (21) using infrared spectroscopy and AA isolated by the octobromo method supported a trans double bond geometry in AA. They had used the old extraction techniques developed by Wesson, which is now known to isomerize cis double bonds into a trans isomer, and led to them to misidentifying AA as contain one or more trans doubles bonds in the molecule (21).

In 1956, a infrared spectrum of AA was published by Holman and Edmondson that identified an absorption at 4650 and 4570 cm‐1, which they state is unique to cis double bond configuration and not trans (22). However, they do not state their source of the fatty acid or the conditions under which they extracted AA to preserve the all cis configuration.

In 1960 and 1961, the all cis configuration of Δ5, 8, 11, 14‐eicosatetraenoate or arachidonic acid was confirmed using synthetic chemistry to selectively form the pure all cis isomer by reduction of alkynes with a Lindlar catalyst. The synthesized material was then compared by IR spectroscopy to an AA standard that had been purchased from Nutritional

Biochemicals Corp (23).

Prior to the structural characterization of AA, many synthetic routes were hypothesized for the formation of AA including unsaturation of arachidic acid or formation directly from carbohydrates (24). However, data published after these initial hypotheses began to suggest a relationship between linoleic and arachidonic acid. Extensive analysis of lipids from animals that were fed different diets found that the fatty acyl composition of glycerolipids was dependent on the fatty acids provided in the fat, specifically diets high in linoleate provided increased levels of arachidonate (25‐29). Burr and Burr discovered that the presence of highly unsaturated fatty acids were essential for growth (30). Specifically, they found that diets supplemented with linoleate and linolenate were essential for proper growth and health of rats; their data supported these two fatty acids as essential fatty acids

5

(31). However, the association of AA and linoleate was not validated until experiments were performed using rats on a fat‐free diet followed by treatment with methyl linoleate which led to increased arachidonate content (32).

Although the essential role of AA was established in the context of health, the link between AA and eicosanoids was not established until 1964 when two independent groups discovered that AA could be converted into prostaglandins (33‐36). Both the Samuelsson and Von Dorp groups used [3H]‐AA incubated with sheep vesicular gland homogenate and observed robust production of a tritium labeled molecule that co‐eluted with purified prostaglandin E2 [PGE2]. With the discovery of the central role of AA, Samuelsson continued to identify novel oxidative products derived from the enzyme cyclooxygenase including endoperoxides, thromboxanes, and the prostaglandins products PGD2 and PGI2

(prostacyclin) (37). Using [14C]‐AA, Samuelsson and Borgeat identified a novel monohydroxy derivative of [14C]‐AA made by circulating neutrophils that came from a family of lipoxygenases, not cyclooxygenases (38). The link between the 5‐lipoxygenase products, leukotrienes, and AA was validated in work published by Murphy and Borgeat in

1979 (39,40).

Non‐activated cells do not have high levels of free AA nor do they produce eicosanoids. Many groups in the 1970’s characterized mechanisms by which to stimulate different types of cells to initiate the release of AA from phospholipids (37,41‐43). When cells are activated (through a variety of mechanisms) the cytosolic concentration of calcium increases, leading to the activation of cytosolic phospholipase A2α (cPLA2α). When cPLA2α is activated by calcium, it causes structural changes within the C2 domain of the protein to reveal hydrophobic residues that facilitate translocation of the protein to membranes (44). Once at the membrane, cPLA2α catalyzes the release of free fatty acids, preferentially AA, from the sn‐2 position of phospholipids (45,46). While cPLA2α has been

6

shown to translocate to the endoplasmic reticulum [ER] and Golgi for the production of prostanoids, it is also translocated to the nuclear envelope for the formation of leukotrienes (47,48). While free AA has many enzymatic and non‐enzymatic fates, the research performed in this thesis has focused on leukotriene formation and reincorporation of AA into phospholipids. However, because prostaglandins have closely tied biologic activity, a brief review of the production and regulation of prostanoids is discussed in the following section.

The Discovery, Synthesis, and Activity of the Major Cyclooxygenase Products–

The term prostaglandin comes from the research performed in 1935 in which two groups discovered a substance that influenced smooth muscle cells and came from human seminal fluid (49,50). It was not until 1960, that Bergström and Sjövall were able to isolate and characterize prostaglandin E and F from sheep prostate glands (51,52). As previously mentioned, the link between AA and prostaglandins was not discovered until 1964. It was these [3H]‐AA experiments that allowed for the identification of 34 products that formed through an initial step by a cyclooxygenase (35). Although it was known that an enzyme in tissue extracts had this function, cyclooxygenase‐1 (COX1) or prostaglandin H2 synthase‐1

(PGHS‐1) was not purified until 1976 and not cloned until 1988 (53‐57). A second cyclooxygenase gene was identified in 1991, which was expressed after certain cellular stimuli (58). The novel gene product was given the name COX2 or PGHS‐2 because of the functional similarity with COX1 (58).

These two COX enzymes share approximately 60% sequence homology with the major difference occurring in a 19 amino acid sequence located just before the C‐terminal tail (59). COX1 is constitutively expressed in most cell types, and located on the ER membrane (60). COX2 is usually a short‐lived protein and induced under a variety of

7

stimuli (60). COX2 has a weak ER retention sequence and can be transported to other membranes through modifications that occur in the Golgi (59). It is well established that

COX1 and 2 are homodimers, however evidence has been published to suggest that the homodimers function as heterodimers (61‐64).

The functional subunit of COX1/2 has a strong affinity for heme which is required for the cyclooxygenase and peroxidase reactions to form prostaglandin H2 (PGH2) (61). The inactive subunit has a weaker binding affinity to heme and is thought to act as an allosteric modulator of the active subunit (61‐64). Some allosteric modifications occur through the binding of fatty acids or other small molecules in the enzymatic pocket in the non‐active subunit. Once bound, these molecules cause conformational changes in the inactive subunit, which directly effects the confirmation of the active subunit. The binding of many fatty acids has been tested to determine how each positively or negatively affects the activity of the active subunit (65).

The following section is a summary of the enzymatic activity of COX and the prostanoid generating enzymes that has been reviewed in much more detail by Smith and colleagues (66‐68). Each monomer contains three structural domains: an epidermal growth factor domain, membrane binding domain, and a C‐terminal globular domain (67).

The active site of COX1/2 is formed between the membrane binding domain and extends into the core of the C‐terminal globular domain (67). The COX1/2 active site functions first as a cyclooxygenase to form prostaglandin G2 (PGG2) and then later performs peroxidase activity at another region of the enzyme which reduces the 15‐hydroperxide group of PGG2 to form prostaglandin H2 (PGH2) (Figure 1‐1). The first step of the COX activity is the removal of the 13‐pro‐S hydrogen from AA leading to a carbon‐centered radical at carbon‐

11. Molecular oxygen attacks the radical leading to a hydroperoxyl radical at carbon 11.

The 11‐hydroperoxyl radical attacks the double bond at carbon‐9 leading to the

8

rearrangement of the carbon‐centered radical at carbon‐15 that reacts with oxygen. This leads to the intermediate product PGG2 that is reduced to form PGH2.

From the production of PGH2, there are five prostanoids that have profound bioactivity: prostaglandin E2 (PGE2), prostacyclin (PGI2), prostaglandin D2 (PGD2), prostaglandin F2α (PGF2α), and thromboxane A2 (TXA2). The enzyme(s) that synthesize each of the prostanoids from PGH2 are shown in Figure 1‐1. The detail of the synthesis of each product is not included, but information about the discovery and function of each enzyme is provided as a reference. PGE2 is synthesized by one of three proteins: microsomal PGE2 synthase‐1, mPGES‐1, m‐PGES‐2, or cytosolic PGE2 synthase (cPGES) (69). Microsomal

PGES‐1 is an inducible enzyme that is part of the membrane‐associated proteins involved in eicosanoid and glutathione metabolism superfamily (MAPEG) and is a glutathione dependent enzyme. Details on mPGES‐1 have been reviewed by Sampey, Monrad and

Crofford (69). Cytosolic PGES is almost universally and constitutively expressed, it is also glutathione dependent, and generally couples to COX1 (70). Microsomal PGES‐2 is ubiquitously expressed; it can also be induced under certain conditions, and is dependent on glutathione or other SH‐reducing agents for activity (69,70). These three enzymes function independently with COX1 and/or COX2 to form PGE2. PGD2 can be synthesized by hematopoietic PGD2 synthase (H‐PGDS), which is a glutathione dependent enzyme, or by glutathione independent lipocalin PGD2 synthase (L‐PGDS) (68). Information regarding H‐

PGDS has been reviewed by Kanaoka and Urade regarding the molecular biology and biochemistry of the enzyme (71). L‐PGDS is thought to be important in the brain. L‐PGDS has been isolated and characterized from brain tissue (72).

The synthesis of PGF2α is performed by human prostaglandin F synthase (PGFS)

(68). PGFS was first isolated in bovine lung and later crystallized; it is thought to be globally expressed in most mammalian tissues (73,74). Prostacyclin is formed by

9

prostaglandin I2 synthase (PGIS) that is primarily located in vascular smooth muscle; it is part of the cytochrome p450 superfamily, also named CYP8A1 (75). The crystal structure of PGIS was published by Chan and colleagues in 2006, which provided evidence for the essential amino acids and its structural homology to the cyp450 family (76). TxA2 is synthesized by TXA2 synthase (TxAS) which was first isolated from platelets in 1976 (77) and fully cloned in 1992 (78). Although the structure of the protein has not been solved, work has been review by Kulmacz and colleagues on the known essential catalytic residues, co‐factors, and secondary structure necessary for proper enzymatic function

(79).

Each product elicits its own physiological response that will be discussed briefly in the following section, but more generally these molecules work in an autocrine or paracrine fashion to affect processes in acute inflammation and immune cell recruitment

(80). Prostanoids act directly on a subfamily of eight rhodopsin‐like seven transmembrane spanning G‐protein coupled receptors. The receptors and their functions have been reviewed in detail (80‐82), but will be briefly described. Within the family of eight receptors there are additional splice variants of some of the receptors. There are four PGE2 receptors with their corresponding G‐proteins provided in parentheses: EP1 (Gq), EP2

(Gs), EP3 (Gi, G12), and EP4 (Gs). Unlike PGE2, the other prostanoids only have one known receptor. The product that binds the receptor is listed first followed by the name of the receptor and finally the G‐protein that it is coupled to: PGD2 (DP1, Gs), PGF2α (FP, Gq), prostacyclin or PGI2 (IP, Gs), and TxA2 (TP, Gq or G13).

10

Arachidonic*Acid*(AA)*

HO

O Cyclooxygenase*1/2*(COX1/2)*

COOH O

O Prostagladin*G2*(PGG2)*

OOH

Cyclooxygenase*1/2*(COX1/2)*

COOH O Prostagladin*H2*(PGH2)* O HBPGDS* LBPGDS* OH TxAS* Thromboxane*A2*(TxA2)*

COOH mPGESB1* O mPGESB2* O cPGES* PGIS* HO Prostagladin*D2*(PGD2)* OH COOH PGFS* COOH O

OH Prostagladin*E *(PGE )* O O 2 2 Prostacyclin*(PGI )* COOH 2

HO HO

OH OH HO Prostagladin*F2α*(PGF2α)* COOH

HO

OH

Figure 1‐1: Biochemical synthesis pathway of the major biologically active prostanoids. Figure is modified from (68). Enzymes are underlined and products with their common abbreviations are shown in parenthesis. Hematopoietic PGD synthase (H‐PGDS), lipocalin‐ type PGD synthase (L‐PGDS), microsomal PGE synethase‐1 (mPGES‐1), cytosolic PGE synthase (cPGES), PGF synthase (PGFS), prostacyclin synthase (PGIS), TxA synthase (TxAS).

11

The biochemical affects of each of the prostanoids can somewhat vary between tissues, however the following is a review of the more general, well‐known functions of each of the prostanoids. Prostaglandin E2 is probably the most discussed prostanoid. Since

PGE2 can act on four different receptor subtypes, it can have a large variety of biologic effects. Some of the well established functions of PGE2 include the induction of the female reproduction, vascular hypertension, tumorigenesis, pain hypersensitivity, and fever (83).

Similar to PGE2, PGD2 has many biologic effects although it is only known to act upon one receptor the DP1 receptor. PGD2 has been suggested to oppose PGE2 in temperature regulation, cause vascular and non‐vascular smooth muscle cell relaxation, induce allergic responses, and inhibit platelet aggregation (84).

Prostaglandin F2α was identified in 1960 (52) and was later described for its ability to induce uterine contraction, smooth muscle cell contraction in the intestines, and bronchoconstriction (85‐87). Thromboxane A2 and prostacyclin are thought to have opposing affects. Samuelsson and colleagues first identified thromboxane A2 in 1975 as an unstable intermediate synthesized by platelets that induced irreversible aggregation in platelets (88). Vane and colleagues were the first to isolate and describe prostacyclin in

1976 (89). In this report they determined that prostacyclin was able to act on gastrointestinal smooth muscle, relax certain isolated blood vessels, and inhibit platelet aggregation (89). The structure was deduced the same year by Vane and colleagues and found to be the intermediate responsible for the production of the more stable but inactive prostaglandin F1α (90). Collectively the prostanoids have diverse as well as opposing effects that mediate many different physiologic responses.

12

The Discovery of Leukotrienes–

The history of leukotriene research began around 1940, when Kellaway and

Trethewie observed that histamine was not sufficient to induce the contraction of the smooth muscle cells that line the airways of the lung (91). This led them to predict that there must be another compound produced to yield this physiologic effect, which they named slow‐reactive substance of anaphylaxis (SRS‐A) (91). It was not until 1960, with the advent of anti‐histamine compounds, that SRS‐A could be isolated and validated as a stimulus independent of histamine (92). Over the next 20 years, numerous investigators found SRS‐A to be produced in different injury models (93).

In 1976, Borgeat and Samuelsson gave exogenous arachidonic acid to neutrophils and observed a novel product 5‐L‐hydroxyl‐6, 8, 11, 14‐eicosatetraenoic acid, now known as 5‐HETE (38). The production of 5‐HETE was not blocked by indomethacin, a cyclooxygenase inhibitor, and therefore was determined to be a product of a lipoxygenase

(38). In 1979, three years after the discovery of the 5‐HETE, the structural identifications were made of two additional 5‐lipoxygenase products: leukotriene B4 (a 5(S),12(R)‐ dihydroxy‐eicosatetraenoic acid) and leukotriene C4 (a cysteine‐containing derivative of 5‐ hydroxy‐7,9,11,14‐eicosatetraenoic acid) (39,40). In 1988, the 5‐lipoxygenase gene was cloned and sequenced (94). After gene identification, the field expanded rapidly, revealing details about synthesis of leukotrienes, development of inhibitors, and the creation of a 5‐ lipoxygenase null animal. This work will be summarized in the next few sections.

The Synthesis of Leukotrienes–

The release of AA is the first critical step in leukotriene formation (Figure 1‐2).

Without the activation and expression of cPLA2α, leukotrienes fail to be produced (95).

Translocation of cPLA2α to the nuclear envelope is critical for the formation of leukotrienes

13

because 5‐lipoxygenase activating protein (FLAP), a protein critical to the proper function of 5‐lipoxygenase (5‐LO), resides in the nuclear envelope (96,97). It is essential that these three proteins localize in close proximity to produce the leukotriene precursor molecule, leukotriene A4 (LTA4). Like cPLA2α, 5‐LO is enzymatically inactive in the cytosol, but through increased cytosolic calcium, 5‐LO translocates to membranes increasing the availability of substrates. Translocation of 5‐LO requires 4‐10 μM cytosolic calcium levels compared to the 0.3‐1 μM for cPLA2α (98‐100). Similarly, the C2 domain of 5‐LO undergoes a conformational change exposing hydrophobic residues which enable the anchoring of 5‐

LO to phosphatidylcholine rich membranes of the outer nuclear bilayer (101). The enzymatic activity of 5‐LO and cPLA2α is independent of the calcium, however, availability of substrates makes calcium binding a critical step in the formation of products for these two enzymes.

The catalytic pocket of 5‐LO has at least six critical amino acid residues that assist in the formation of LTA4. Non‐heme iron is essential for the catalytic activity of 5‐LO. Two histidine (His) residues, His‐372 and His‐550, were found to be important in the coordination of the iron in the active site (102,103). From the crystal structure of 5‐LO, it was suggested that His‐367 and isoleucine‐673 also participate in the coordination of iron in the active site (104). In addition, residues glutamate‐376 and glutamine‐558 are known to be critical to the enzymes’ ability to synthesize leukotrienes, however, the function of these glutamate residues for enzymatic activity is not understood (103,105). Entry of AA into the catalytic pocket of 5‐LO is not completely understood. There are three hydrophobic amino acids that have been suggested to block entry into the enzymatic pocket. Two mechanisms have been proposed, that either entry of AA occurs from the carboxy or from the omega end, both lead to AA being in the proper alignment with the catalytic iron (106).

14

Once AA is in the active site, 5‐LO performs two catalytic events on the AA substrate. The first is the conversation of AA to 5‐hydroperoxyeicosatetraenoic acid (5‐

HpETE) (Figure 1‐2). During this initial step, a proton is removed from carbon‐7 of AA leading to the formation of a radical on AA. The catalytic Iron (III) gains an electron from the removal of the proton converting it into the Iron (II) state. This radical can then react with molecular oxygen to create the hydroperoxy radical which subsequently gains an electron from the iron. Thus oxidizing Iron (II) back to the Iron (III) state for the second enzymatic step of 5‐LO. If 5‐HpETE is exposed to aqueous environments, it can be converted to the more stable 5‐hydroxyicostetraenoic acid (5‐HETE) by the action of various peroxidases.

If 5‐HpETE is released from 5‐LO at this stage of the catalytic cycle, it can undergo a second catalytic event to produce LTA4 (Figure 1‐2). In this secondary reaction, a hydrogen atom is removed from carbon‐10 leading to the reduction of the iron in the active site leaving a radical on AA that can delocalize over multiple carbons. The new radical site on carbon‐10 destabilizes the 5‐hydroperoxide group, driving a reaction of a nascent hydroxyl radical with Iron (II) in the 5‐LO active site. The Iron (II) now donates an electron to the hydroxyl radical driving the reaction forward to form the conjugated triene and epoxide at the 5,6 position, producing LTA4. These radical‐mediated steps also regenerate Iron (III).

The mechanism of self‐regeneration of the iron in the active site allows the enzyme to immediately engage the next catalytic cycle.

The role of FLAP in the formation of LTA4 is poorly understood, however it is essential in the production of leukotrienes (96). This small integral membrane protein,

FLAP, was discovered accidently when inhibitors were being developed against 5‐LO

(97,107). Discovery and development of these inhibitors will be discussed in the next section, however, their discovery has allowed more insight into 5‐LO stabilization of

15

reactive intermediates (108). Current understanding of FLAP supports the idea that it has no enzymatic activity itself, it may present AA to 5‐LO, and that it is critical for the in vivo formation of leukotrienes (109,110).

Once LTA4 is formed by 5‐LO in coordination with FLAP, it can undergo further reactions by two different enzymatic pathways. One pathway engages an enzyme called

LTA4 hydrolase to produce LTB4 (111). Unlike 5‐LO, LTA4 hydrolase is a metalloenzyme expressed in almost all cell types (112). LTA4 hydrolase is a soluble enzyme that specifically uses LTA4, a highly reactive intermediate, as a substrate (113,114). With the use of zinc in the enzymatic pocket, LTA4 hydrolase catalyzes the conversion of LTA4 to

LTB4 with the use of two water molecules. The first water molecule attacks at carbon‐12 causing the destabilization of the 5,6‐epoxide. The second water molecule is coordinated near the catalytic zinc to open the epoxide to form a hydroxyl at carbon‐5. This leads to the production of the biologically active LTB4 (Figure 1‐2).

Alternatively, LTA4 can be a substrate for a small hydrophobic protein, LTC4 synthase. Although not a classical glutathione (S) transferase (a protein that covalently attaches the tripepetide glutathione (γ‐glutamyl‐cysteinyl‐glycine) to substrates), LTC4 synthase is specific for the addition of glutathione onto LTA4 (68). It is structurally unique from the other proteins that catalyze the addition of glutathione, but surprisingly shares around 30% identity with FLAP (115). The enzyme functions as a homo‐trimer. Based on the crystal structure of the protein complex, it is believed that LTA4 can sit at the interface between dimers (116). This confirmation allows arginine‐104 to be in perfect alignment to activate the glutathione. This provides a thiolate anion that attacks carbon‐6 of the epoxide allowing the opening of the ring. The resultant alkoxide anion is then poised to remove a proton from Arg‐31, leading to the production of LTC4 (Figure 1‐2). Leukotriene C4 can then be further converted to leukotriene D4 and subsequently leukotriene E4 by γ‐glutamyl

16

transpeptidase, and a membrane‐bound dipeptidase (117). LTC4, LTD4, and LTE4 are collectively referred to as the cysteinyl leukotrienes.

Arachidonic*Acid*(AA)*

HO

O 55Lipoxygenase*(55LO)* 55LO*Ac>va>ng*Protein*(FLAP)*

HO

O 55Hydroperoxide*(55HpETE)* O OH 55Lipoxygenase*(55LO)* 55LO*Ac>va>ng*Protein*(FLAP)* HO O O Leukotriene*A4*(LTA4)* O LTC4*Synthase* LTA4*Hydrolase* NH O O H N OH OH HO NH2 HO S O HO HO HO

O Leukotriene*B4*(LTB4)* O Leukotriene*C *(LTC )* 4 4

Figure 1‐2: Biochemical synthesis pathway of leukotrienes. Enzymes are underlined and products with their common abbreviations are shown in parenthesis.

17

LTC4 synthase is widely expressed in cells such as platelets, eosinophils, basophils, mast cells, vascular smooth muscle cells, endothelial cells, and monocytes, as well as in lung tissue (118,119). This raised an interesting question in the field as to why cells that do not express 5‐LO, which is required for the formation of LTA4, express enzymes that are in the final synthetic pathway for leukotrienes. Remarkably, cells are able to pass LTA4, which is unstable in water from cell‐to‐cell without exposing it to the aqueous environment.

Therefore, multiple cells can participate in the final synthesis of leukotrienes. An example of this cell‐cell coordination was supported by a paper that showed certain cell types could not synthesize a leukotriene product by themselves, but could biochemically cooperate with another cell type to synthesize a leukotriene (120). The mechanism by which LTA4 is transported between cells is not known, but it is suggested that there are LTA4 binding proteins that allow the shuttling of this reactive intermediate between cells (120).

The Biologic Role of Leukotrienes–

The regulation of leukotriene metabolism is important to many disease processes.

The following section provides a background of the known biological activity of these molecules and the effects they have once bound to their receptors. Once the leukotrienes are produced, through the mechanisms previously described, they have to be exported to initiate a biological effect, acting in an autocrine or paracrine fashion. Leukotrienes molecules are exported through facilitated transport using ATP‐binding cassette (ABC) efflux pumps (121). The transporter that is known to export LTC4 is ABCC1 or multidrug resistance‐associated protein 1 (MRP1) (122). The transporter known to efflux LTB4 is

ABCC4 or MRP4 (123). Once outside of the cell, these lipid mediators can bind to their respective receptors to induce an effect.

18

There are two receptors responsible for the recognition of LTB4: leukotriene B4 receptor 1 and 2 (BLT1 and BLT2). These proteins are GPCRs that have similar homology with the fMLP receptor family members (124). The BLT1 receptor has about a 20‐fold higher LTB4 binding affinity than that of the BLT2 receptor (125,126). The BLT2 receptor has a broader ligand recognition including molecules made by other lipoxygenases compared to BLT1 (126). Using genetically null animals for BLT1, it was determined that the receptor was essential for the chemotaxis of neutrophils and peritoneal macrophages

(127,128). The role of LTB4 binding to BLT2 is not as well understood. However, since

BLT2 does have a broader substrate specificity and global expression, it may have a different functional role compared to that of BLT1 (129). Some work has been done with the BLT2 null animal, which has led to the suggestion that this receptor has an anti‐ inflammatory role in inflammatory colitis (130). The emerging view as to the role of BLT1 and BLT2 receptors suggests that they have opposing effects. BLT1 has been shown to be pro‐inflammatory, while BLT2 is suggested to be anti‐inflammatory (131).

There are several GPCRs for the cysteinyl‐leukotrienes with the two first being named cysteinyl‐leukotriene receptor 1 and 2 (CysLT1 and CysLT2) (106). These two

GPCRs are thought to have different affinities for each of the cysteinyl‐leukotrienes. CysLT1 preferentially binds LTD4, LTC4, and with the lowest affinity LTE4 (132). CysLT1 has been found in many experimental models to drive smooth muscle cell contraction and increased vascular permeability (106). The drug montelukast, which will be discussed in a different section, was developed to block the action of CysLT1 in chronic asthma (133). The CysLT2 has a similar ligand preference, binding LTC4 and LTD4 with equally affinity, but LTE4 with lesser affinity (134). The role of CysLT2 appears to be more critical in small vessels, modulating vascular permeability (106). Other than these two widely studied receptors, there are three additional receptors that are activated by the cysteinyl‐leukotrienes: gpr17,

19

P2Y12, and CysLTE (106). The role of these additional receptors in the response to leukotrienes is not as established as the role of CysLT1 and CysLT2, but suggests additional complexity of signaling events and biologic role for the cysteinyl leukotrienes.

Understanding the role of leukotrienes in most disease processes is expanding, but in some instances contradicting effects have been observed. Many experiments have employed a 5‐LO null animal or inhibitors of the 5‐LO pathway, but finding a positive effect by this strategy does not provided insight into which leukotriene is the critical mediator. 5‐

LO has been implicated in many disease processes such as asthma, atherosclerosis, traumatic brain injury, stroke, and several types of cancer (106,135‐138). Our understanding of the exact mechanism by which leukotrienes are involved in the progression of such diseases is still in its infancy, but it is now clear that these eicosanoids have a critical role in many disease processes.

The Development of Pharmacologic Intervention of Eicosanoids–

Ancient therapies, now known to reduce bioactive lipid biosynthesis, were developed during the time of the Egyptians when mixtures of willow leaves were placed on inflamed wounds. Later, Hippocrates, following the Egyptian finding, used willow bark circa 460‐377 BC (139). He discovered that he could create a powder from the leaves and bark of willow trees and alleviate pain, headaches, and fever. However, it was not until

1829 when scientist, Johann Buchner, isolated a compound called salicin found within the willow plants that caused these pharmacologic effects. Raffaele Piria in 1838 was able to converting salicin to salicylic acid. By 1900, Felix Hoffmann working for Bayer patented acetylated salicylic acid and call this product aspirin which is still used today (139).

Since the original discovery of willow bark extract and synthesis of salicylic acid and aspirin, understanding how these drugs exerted their effect was unknown. It was not

20

until the work of Piper and Vane that the connection to inhibition of prostaglandin biosynthesis was made and the mechanism of action of aspirin as an inhibitor cyclooxygenase was determined (140). There are two classes of COX inhibitors: nonsteroidal anti‐inflammatory drugs (NSAIDs) and anti‐inflammatory steroids. In 1982,

Sune Bergström and Bengt Samuelsson won the Nobel prize in physiology or medicine for their original structural identifications of prostaglandins, and shared the Nobel prize with

John Vane who was the first to describe aspirin’s and other NSAIDs ability to inhibit the production of prostaglandin synthesis in 1971 (141,142).

The emergence of new bioactive eicosanoids, including SRS‐A or what is currently known as the cysteinyl‐leukotrienes, yielded additional drug development (143). Known for their ability to induce smooth muscle cell contraction and edema, blocking cysteinyl‐ leukotriene biosynthesis became of interest in chronic asthma (143). One class of drug that eventually was approved by the FDA was a receptor antagonist by Merck and the other a direct inhibitor of 5‐lipoxygenase by Abbott Pharmaceuticals. In 1989, Merck published a paper focusing on a potent inhibitor of the CysLT1 receptor, MK‐571. MK‐571 later went to market under the generic name montelukast or the proprietary name Singulair, where it is still currently being used to treat moderate to severe asthmatic patients (133).

During the same time, development of inhibitors against the protein 5‐ lipoxygenase was successful. In 1991, Abbott Pharmaceuticals identified an inhibitor in which they named zileuton (144). In 1996, the drug was approved by the FDA, but had notable liver toxicity and poor compliance due to the dosing regimen. In addition to the compliance issues, both montelukast and zileuton have reported side effects. These side effects were sufficiently serious that in 2008 the FDA mandated a review of neurological profile of patients currently on montelukast (Singulair) (146). While both of these drugs are efficacious, these side‐effect profile and dosing regimens have remained problematic.

21

An alternative strategy was pursued that involved inhibition of leukotriene were found to block biosynthesis, but not by the direct inhibition of 5‐lipoxygenase. Drugs such as MK‐886 and MK‐591 inhibit leukotriene formation by blocking FLAP (147). Even though the FLAP inhibitor program was abandoned by Merck because of the success of montelukast, this avenue of drug development was continued by a small biotech company, which later found drugs with a much high affinity and fewer side effect profiles than the previous FLAP inhibitors, with drugs AM‐103 and AM‐803 (148,149).

The rights to the patent for AM‐803 and similar analogs were purchased by

GlaxoSmithKline. The analog, renamed GSK2190915, completed phase II trials (150).

GSK2190915 can be taken orally on a daily dosing schedule and has been one of the most promising for leukotriene drugs thus far. Johnson & Johnson has also completed Phase II clinical trials of a LTA4 hydrolase inhibitor (JNJ‐40929837) (151).

Even with the success of these current treatment strategies, the question still remains: Is the most successful strategy inhibiting the production of one or a set of molecules completely? Many of these bioactive lipids oppose each other, which was brought to light during the development of cyclooxygenase‐2 inhibitors. Instead of directly inhibiting eicosanoid enzymes, an alternative approach could be taken to increase or decrease leukotriene production through modulating the AA substrate pool in phospholipids. AA is incorporated into phospholipids through the reacylation pathway. A better understanding of phospholipid synthesis and remodeling could possibly reveal alternative mechanisms in the regulation of eicosanoid biosynthesis. The following section will discuss the current understanding of the regulation, remodeling, and modification of phospholipid molecular species that contain esterified arachidonate.

22

Phospholipid Synthesis and Remodeling–

The first identification of phospholipids was made by Theodore Gobley in 1847 when he described a component in egg yolk which he eventually termed lecithin (152).

Gobley identified the components of the phospholipids that he isolated, as containing two esterified fatty acids, a glycerol, a phosphate, and a choline head group (153). It is now known that phospholipids are essential components of cellular membranes, which have amphipathic properties important in creating selective barriers against the free movement of molecules. There are seven major phospholipid classes, and several minor classes. These seven classes are phosphatidic acid (PA), phosphatidylcholine (PC), phosphatidylethanolamine (PE), phosphatidylglycerol (PG), phosphatidylinositol (PI), phosphatidylserine [PS], and cardiolipin (CL).

In the 1950’s, it was found that there were two branches, the Kennedy pathway and Lands’ cycle, that have overlapping enzymes to dictate the quantity and diversity of phospholipids synthesized across multiple phyla of organisms. The de novo assembly of phospholipids, the Kennedy Pathway, relies upon the specificity of glycerophosphate‐ and acylglycerophosphate‐acyltransferases to dictate the fatty acyl composition of downstream phospholipid products (154,155). All of these phospholipids species originate from the biosynthetic pathway shown in Figure 1‐3. The details of phospholipid synthesis and remodeling will be briefly reviewed, however, details of some individual steps will be elaborated on in later sections.

The first committed step in the synthesis of all phospholipids comes from the acylation of glycerol‐3‐phosphate (G‐3‐P) by three enzymes termed glycerol‐3‐phosphate acyltransferases (GPATs) (Figure 1‐3). GPATs catalyze the reaction between G‐3‐P and fatty acyl coenzyme A (CoA) esters. The initial stages of phospholipid synthesis can occur in the endoplasmic reticulum (ER) or the mitochondrion (156). Studies have shown that

23

GPAT3 is located on the ER and is non‐specific for the addition of an acyl chain at the sn‐1 position of the glycerol (157,158). GPAT1 and GPAT2 are located on the mitochondria

(159,160). GPAT1 more selectively adds palmitoyl‐CoA (16:0‐CoA) to the sn‐1 position of the glycerol leading to the production of a lysophosphatidic acid (lyso‐PA) (159).

GPATs, along with many other acyltransferase family members, require fatty acyl

CoA esters to facilitate the addition of fatty acyl chains. The formation of these amphipathic lipids is catalyzed by fatty acyl‐CoA synthetases (ACS). There are twenty‐six known family members that are separated into four distinct classes: short chain (2:0‐4:0), medium chain

(6:0‐10:0), long chain (14:0‐22:6), and very long chain (24:1 and above) (145). However, many of these enzymes do not fall perfectly into any specific category. All of these enzymes catalyze the covalent linkage of fatty acids to coenzyme A in a two‐step process consuming

ATP (161). Each isoform has a unique subcellular location and tissue specific distribution.

These fatty acyl CoA esters are critical in the de novo synthesis of phospholipids, remodeling of phospholipids, and β‐oxidation and elongation/desaturation of fatty acids.

During the formation of PA, a second fatty acyl CoA ester is used by a LPAT with the lysophosphatidic acid previously formed to complete the initial step in the de novo phospholipid synthesis pathway. Similar to the GPATs, there are LPATs located on the mitochondrion and ER. However, most of the activity is found in enzymes located on the

ER. There are four known enzymes that acylate the sn‐2 position of lysophosphatidic acid: lysophosphatidic acid acyltransferase 1 (LPAAT1), LPAAT2, LPAAT3, and lysophosphatidylcholine acyltransferase 1 (LPCAT1) (162). The known acyl chain specificities of these enzymes are summarized in Figure 1‐4. The initial acyl composition of all of the phospholipids comes from the activity and acyl substrate availability of these four enzymes. From the production of PA, phospholipid biosynthesis breaks into two separate pathways, both that take place in the ER.

24

One pathway, shown in Figure 1‐3, leads to the production of PE, PC, and PS. The first committed step in this process is the removal of the phosphate from PA by phosphatidic acid phosphatase (PP) to form a diacylglycerol [DAG] (163). Once the DAG has formed, phosphocholine or phosphoethanolamine can be added by CDP‐choline: DAG cholinephosphotransferase (CPT) or CDP‐ethanolamine: DAG ethanolaminephosphotransferase (EPT), respectively. It was in 1955 that Kennedy and

Weiss performed a series of experiments with different nucleotides and found that choline formed a CDP‐choline intermediate that was critical in the formation of PC (164). As shown in Figure 1‐3, the use of CTP has been determined to be a critical component for the formation of all phospholipids downstream of PA. Three protein isoforms have been found in humans that encode for the CDP‐choline and/or CDP‐ethanolamine 1,2‐DAG activity.

These transmembrane proteins have a DAG binding domain, CDP‐choline or CDP‐ ethanolamine binding domain, and a cation in the active site to function properly (165).

The products of the reaction are PC or PE, CMP, and a proton.

The newly formed PC and PE can remain, thus contributing to the 50% and 25% of total phospholipid pools, respectively. Alternatively PC or PE can be converted to PS through PS synthases (PSS). There are two known isoforms: PSS1 uses PC and PSS2 uses

PE to catalyze the head group exchange (166). The functional moieties of these two integral membrane proteins are not well understood, but many residues have been suggested to be critical to the serine‐exchange function (167). Once PS is synthesized it can be decarboxylated by phosphatidylserine decarboxylase (PSD). PSD uses a pyruvate cofactor to catalyze the decarboxylation of PS producing PE and releasing CO2 (168).

25

O GPAT% O

O O P O HO O P OH O OH OH

OH OH Lyso%PA% Glycerol)3)phosphate% Fa#y%Acyl) CoA% CoA%Ester%

O

O O P O

O O OH

O LPAT%

Fa#y%Acyl) CoA% CoA%Ester% Phospha4dic% Acid%(PA)% NH2 N

O N O O O H3C O O P O OH H C 3 N P OH O H3C O OH O HO OH O CPT% CDP)choline% PP%

DAG% Phospha4dylcholine%(PC)% CDP)DAG% Synthase% PEMT%

NH2

N CDP)DAG%

O N O Phospha4dylethanolamine%(PE)% O O O P P O

O O OH OH NH2 HO OH R1 O EPT%

N R2 PSD%

O N O O PG)P%Synthase% PG)P% O H O O O P H N P OH HO O O P OH H O OH PSS% HO OH PI%Synthase% P OH OH CDP) O O O ethanolamine%

R1 O NH2

R2 N Phospha4dylserine%(PS)% Phospha4dylinositol%(PI)% O N O O O O P P O

O O OH OH HO OH R1 O

R2 CDP)DAG% Phospha4dylglycerol%(PG)% Cardiolipin%(CL)%

Figure 1‐3: The de novo synthesis of phospholipids modified from D.E. Vance and J.E. Vance, 2008 (169).

26

PE can be converted to PC through methylation as an alternative pathway to form

PC. This alternative pathway constitutes the production of approximately 30% of total PC content in the liver, and is critical in the regulation of very low‐density lipoprotein formation (170,171). The enzyme that catalyzes this reaction is PE methyltransferase

(PEMT). This protein is able to place three subsequent methyl groups on the amine head group eventually leading to the formation of PC (172). The production and exchange between PC, PE, and PS suggests that these phospholipid molecular species are co‐ regulated to provide the proper balance of each.

Alternatively, the phosphatidic acid that is made in the de novo pathway can be diverted to form PI, PG, or CL. Similar to the formation of PC and PE, the use of CTP is critical in the production of PI, PG, and CL. However, it is not the head group that gets the

CDP covalent linkage, but rather PA is converted to CDP‐DAG. This step is thought to be the rate‐limiting step in the formation of PI and CL (Figure 1‐3).

Although previous groups had described inositol‐containing phospholipids in bacteria and soybeans, inositol containing phosphatides were first discovered in mammalian brain and spinal cord tissues by Folch and Wooley in 1942 (173‐175). PI comprises approximately 10% of total cellular phospholipids in most cells. The formation of PI comes from the covalent linkage between inositol and PA from a CDP‐DAG donor through the actions of PI synthase (shown in Figure 1‐3). The inositol head group, predominately the myo‐inositol isoform, can be taken in by diet, recycled or made by de novo synthesis from glucose‐6‐phosphate. The availability of inositol or PI synthase does not appear to change the rate of synthesis, however, deletion of PI synthase has been shown to be lethal in yeast and mammals (176). Recently, it was shown that the PI synthase does not have acyl chain specificity for the addition of the inositol group (177).

27

4:0$% 2:0% 14:0%16:0%18:0% 18:1%18:2%20:4%20:5%22:6% 12:0% LPAAT1%

LPAAT2%

PA% LPAAT3%

LPCAT1%

LPCAT1%

LPCAT2%

PC% LPCAT3% LPCAT4%

LPCAT3%

LPCAT4% PE% LPEAT1%

LPEAT2%

LPCAT1%

LPGAT1% PG% LPEAT2%

LPIAT1% PI% LPAAT3%

LPCAT3%

LPEAT1% PS% LPEAT2%

Figure 1‐4: Summary of the known activities of the lysophospholipid acyltransferases. Figure modified from Hishikawa D. et al 2007. Boxes that are shaded indicate that the activity has been found for the enzyme, unshaded boxes indicate no activity or the activity has not been tested.

28

The other use of CDP‐DAG is for the synthesis of PG and ultimately cardiolipin (CL).

CL was first discovered in bovine heart in 1942, and its structure was determined in 1964

(178,179). PG was not found until 1958 and was isolated from Scenedesmus algae, and its structure and biosynthetic pathway was determined in 1963 (180‐182). PG is a minor phospholipid component in most cells except for cells in the lung and in pulmonary surfactant. In most cells, PG is predominately used as a precursor of CL. PG synthase uses

CDP‐DAG and phosphoglycerol to form a phosphatidylglycerol‐phosphate intermediate.

This molecule is dephosphorylated by phosphatidylglycerophosphatase to form the PG. PG is a short‐lived species that is converted to CL by the transfer of PA from CDP‐DAG to PG to form CL (183).

All of these phospholipids come from the de novo synthesis pathway through a PA intermediate. The acyl chain composition of most phospholipids is dictated by LPAAT1,

LPAAT2, LPAAT3, and LPCAT1. Although these three LPATs have broad substrate specificity, the majority of the phospholipids from the de novo synthesis route contain saturated acyl chains like palmitate (16:0) and stearate (18:0) or monounsaturates like oleate (18:1). However, when analyzing phospholipids, it is quickly revealed that after the initial production of these phospholipids, the majority of phospholipids from the de novo synthesis pathway undergo remodeling to introduce asymmetry and unsaturation. This remodeling pathway is referred to as the Lands’ cycle.

The Lands’ Cycle remodels the products of the Kennedy pathway usually through sequential action of a phospholipase and a lysophospholipid acyltransferase (LPAT) (184).

Thus, the Lands’ cycle produces a different repertoire of phospholipid molecular species from those derived from the Kennedy Pathway (185). Described during the same time period as the Kennedy pathway, W. E. Lands observed that there was an independent pathway of phospholipid biosynthesis that did not require the de novo synthesis of the

29

lysophospholipids. Rather, a fatty acyl group was removed, then replaced or added back to the lysophospholipid. He suggested that it was through the activity of phospholipases that a fatty acid was removed, and that it could be added back via a coenzyme A/ATP dependent mechanism (185). It is now known that the acyl chain composition of phospholipids is the result of a combination of multiple enzymatic activities, including phospholipases, acyl CoA synthetases (ACS), and lysophospholipid acyltransferases

(LPATs), which lead to the large diversity of phospholipid species necessary for different membrane characteristics (186).

Although the initial enzymatic characterization of the remodeling pathway occurred over 50 years ago, many of the genes that encoded these enzymes were not discovered until recently. Important enzymes in the remodeling pathway were finally identified, only after various complete genomes were sequenced and collections of candidate gene cDNAs were empirically tested for their enzymatic functions. Since 2005, several groups have performed extensive analysis of the function of these enzymes. It was discovered that LPATs belonged to two different protein families; represented by the lysophosphatidic acid acyltransferases (LPAATs) and the membrane bound O‐ acyltransferases (MBOATs) (162,184,187‐197). In addition to the LPCATs there are also acyltransferases both AA dependent and independent that remodel phospholipids (186).

The details of the Lands cycle enzymes and products involved will be discussed in the following section.

Each acyltransferase’s substrate preference reveals much about certain phospholipid molecular species, but some synthetic schemes are still not known. This body of work will focus primarily on the incorporation of two fatty acyl groups, arachidonoyl (AA) and docosahexaenoyl (DHA). The incorporation of these two acyl chains is critical for numerous cellular functions, including regulation of lipid mediators,

30

functionality of proteins, and membrane fluidity (198).

Knowledge concerning the route of incorporation of DHA into phospholipids has remained elusive. Only two acyltransferases have been identified to use docosahexaenoyl‐

CoA as a substrate: LPCAT1 and LPCAT2 (188,199). However, the DHA‐CoA activity in overexpression systems was not robust with either enzyme. Also, LPCAT1 and LPCAT2 have activities to incorporate DHA‐CoA into lyso‐PC. PE and PS are the major phospholipids in vivo to contain docosahexaenoyl acyl chains. The route for how DHA is incorporated into PE and PS is still not understood.

In the mid‐2000s, several groups identified six proteins that are able to incorporate

AA into phospholipids. Through the de novo synthesis of PA, AA can be esterified at the sn‐2 position via LPAAT1, LPAAT2, or LPAAT3 (200‐202). This would explain how any phospholipids could contain AA at the sn‐2 position. However, this mechanism of AA addition through the de novo pathway does not appear to be the predominate means of adding AA to phospholipids since only specific types of phospholipids contain this acyl chain. The alternative pathway of incorporating AA into phospholipids through the Lands’ cycle emerged as a potential mechanism.

Numerous phospholipase A2s can remove the acyl chain from the sn‐2 position of phospholipids (203). Some phospholipases will selectively remove acyl chains based on the associated head group and or acyl composition, but others are less specific (203). Once the acyl chain is removed, these lysophospholipids are available to be reacylated by LPATs

(example shown in Figure 1‐5). Each LPAT enzyme has specificity for the head group and the fatty acyl group that it will utilize. This leads to specific phospholipids that are formed.

There are three acyltransferases thought to be critical in the incorporation of AA into phospholipids: membrane bound O‐acyltransferase 5/ lysophosphatidylcholine acyltransferase 3 (MBOAT5/LPCAT3), membrane bound O‐acyltransferase 7/

31

lysophosphatidylinositol acyltransferase 1 (MBOAT7/LPIAT1), and lysophosphatidic acid acyltransferase 3 (LPAAT3) (192,202). These three enzymes that utilize AA‐CoA as a substrate are thought to be essential in the regulation of AA containing phospholipid molecular species in PC/PE/PS (MBOAT5/LPCAT3) and PI (MBOAT7/LPIAT1 and

LPAAT3).

O O O

O O P O R O O P O R O O P O R 3 3 3 O O O O OH OH OH O OH O O

PLA2% LPAT%

FA% Fa(y%Acyl, CoA% CoA%Ester%

Figure 1‐5: Reacylation of lysophospholipids leads to incorporation of polyunsaturated fatty acids and phospholipid asymmetry. Enzymes with this function are displayed above the arrows. Substrates and byproducts are listed below the arrows.

32

Additional Modification of Phosphatidylinositol–

The complex synthesis and remodeling of phospholipids was described in the previous section of this thesis. However, the inositol head group of phosphatidylinositols can be phosphorylated which plays an essential role in many biological functions. In the

1950s, it was discovered that certain phospholipids were decreased when cells were stimulated. Eventually a group of phosphorylated phosphatidylinositols were identified

(204). There are currently seven known phosphorylated forms of phosphatidylinositols in three classes: monophosphates, bisphosphates, and triphosphates (shown in Figure 1‐6).

Myo‐inositol is coupled to diacylglycerol phosphates via phosphodiester linkage to the hydroxyl group at carbon‐1, leaving the five other hydroxyl groups available for phosphorylation. Due to steric‐hindrance from the phosphate group at carbon‐1, the hydroxyls at carbon‐2 and ‐6 are usually not phosphorylated. However, the other three‐ hydroxyl groups are readily available for phosphorylation (204).

There are three phosphatidylinositol kinases that phosphorylate different hydroxyls of the inositol ring with the use of ATP as a phosphate donor. They were aptly named for the hydroxyl group that they phosphorylate: phosphatidylinositol 3‐kinases

(PI3K), phosphatidylinositol 4‐kinases (PI4K), and phosphatidylinositol 5‐kinases (PI5K).

These phosphorylations can then be reversed by a group of phosphatases and myotubularins (MTMs). The initial phosphorylation is used both as an intermediate step for additional phosphorylation and to generate specific phosphoinositides to engage protein domains that assist in many different cellular processes (204).

There are specific protein domains that associate with phosphorylated and non‐ phosphorylated PI’s. Many of these protein domains need additional anionic lipids to strengthen their interaction with the phosphoinositides. The protein domains known to associate with phosphoinositides are FYVE (Fab1/YOTB/Vac1/EEA1), Phox homology

33

(PX), glucosyltransferases, Rab‐like GTPase activators and myotubularins (GRAM), Four‐ point one/Ezrin/Radixin/Moesin (FERM), ENTH/ANTH, C2 domains, pleckstrin homology

(PH), and plant homeodomain (PHD) (204,205).

Each of these domains is critical in recognizing the different phosphorylated form of phosphatidylinositol. Furthermore, each of these phosphorylated forms and protein domains have unique functions. With the use of modified protein domains with conjugated fluorescent tags, the subcellular locations of the different phosphoinositides species have been described (206). However, as a whole, the previously listed phosphoinositides have been shown to be critical components for protein recruitment and docking in apoptosis, cell cycle regulation, endocytosis, exocytosis, ion channel activity, and actin filament assembly (204).

Recently, more sophisticated methods have been developed to measure the different phosphoinositides using mass spectrometry (207). These studies have shown enrichment of the acyl chains corresponding to an 18 carbon fatty acyl chain and a 20 carbon‐4 double bond fatty acyl chain (18:0/20:4) in the phosphorylated forms of PI. It has also been shown that many of the enzymes that regulate the formation of phosphoinositides prefer to use the 18:0/20:4‐PI species (208). The enrichment of this acyl composition and the enzymes that rely on this composition for identification of PI has shed light on the critical regulation of phospholipid acyl chain composition in the phosphatidylinositol class. The data further supports the essential role of the phospholipid reacylation pathway in the correct formation of phosphoinositides for biological functions.

34

PI#

3%PIP# 4%PIP# 5%PIP#

4,5%PIP2#

3,4%PIP2#

3,5%PIP2#

3,4,5%PIP3# DAG+IP3#

Figure 1‐6: Phosphorylation states phosphatidylinositol. Phosphatidic acid (PA).

35

Thesis Goal–

Since the discovery of SRS‐A and its effect on the lung, it has been the target of inhibition. There has been significant effort placed on the direct inhibition of 5‐ lipoxygenase products. However, the history of blocking bioactive lipid production completely has been plagued with complexities. Our laboratory has previously worked on the modulation of leukotriene production through controlling the substrate availability.

The goal of this thesis work is to further advance the understanding of the regulation that phospholipid remodeling plays in the regulation of acyl chain composition of phospholipids and ultimately the effect that this maneuver has on leukotriene production.

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CHAPTER II

LIPID MASS SPECTROMETRY

Introduction–

Originally developed in the first decade of the twentieth century, mass spectrometry’s origins were in measuring the mass of atoms (209). However, since its advent it has been a key in the discovery of many bioactive compounds. Many techniques like NMR and IR spectroscopy and elemental analysis require large quantities of highly pure material to be available. For many biologically active compounds that activate high affinity receptors it is not critical to reach high concentrations. Mass spectrometry has been fundamental in finding compounds that are made in low quantities that are difficult to isolate from biologic matrices. This chapter will focus on the use of two ionization techniques that have been essential in the field of biochemistry: electrospray ionization

[ESI] and matrix‐assisted laser desorption ionization (MALDI). Also, two types of mass analyzers will be discussed that are often coupled to these ionization sources: quadrupole and time of flight (TOF) analyzers. Finally this chapter will cover the different types of experimental methods that were implemented during this thesis work including tandem mass analyzer experiments and MALDI imaging mass spectrometry.

Techniques for the Ionization of Lipids–

Until the discovery of soft‐ionization techniques (which will be discussed in this section), molecules that were analyzed by mass spectrometry were often limited to small, non‐polar molecules, that had to be volatile so as to enter the mass spectrometer as a gas.

The inability to ionize large biomolecules because of very low or no vapor pressure inhibited the analysis of molecules by electron, chemical, and photo ionization techniques

(210).

37

MALDI was first developed in 1985 by Franz Hillenkamp and Michael Karas (211).

They discovered that when they used a laser that had a wavelength equivalent to the absorption maximum of tryptophan they could more readily ionize this and other amino acids co‐mixed with tryptophan (211). This finding led to the use of a matrix, which was added to the sample, to absorb the energy of the laser and more readily lead to formation of ions. The unique aspect of MALDI was it bypassed the need to make a gaseous molecule prior to ionization; instead, ions were desorbed directly from a surface. The laser technology was the key to obtaining improved ionization. In 1987 Koichi Tanaka combined

30 nm cobalt particles with a 337 nm nitrogen laser for ionization. Many lasers have been shown to work extremely well, the small inexpensive nitrogen lasers remained popular until diode lasers emerged (209,212).

The matrix application and the matrix itself have been extensively studied, but what is critical is that the sample molecule of creates co‐crystals with the matrix. The common matrices that are used for the analysis of lipids are organic molecules such as 2,5‐ dihydroxy benzoic acid (DBA) for positive ion analysis or 2' 5'‐dihydroxyacetophenone

(DHAP) for negative ion analysis (213,214). The MALDI samples are placed in a chamber adjacent to an inlet orifice to the mass spectrometer in a position where a laser beam can be focused to effect ion desorption. This chamber can be at atmospheric pressure or under vacuum, but for the experiments in this thesis the chamber was at a vacuum pressure of approximately 0.1 mTorr. When the laser hits the crystalline structures, it leads to thermal explosion of ions (temperatures reaching 1500 K) from the surface of the laser ablation

(Figure 2‐1A and B). Several mechanisms of ion formation have been suggested, and the current thought is that there are multiple mechanisms that lead to ion formation in the gaseous state (212). However, once ionized the ions move towards ground potential

38

(labeled as grid/lens in Figure 2‐1C) since ions are in a high electric field, as seen in Figure

2‐1C and enter into the mass analyzer on the other side of the inlet orifice.

Another way to ionize non‐volatile, biological molecules is to use electrospray ionization. The advantage of electrospray ionization is that it can be coupled to many different chromatographic techniques that enable facile transfer of samples in solution to the mass spectrometer ion sources. Once chromatographed, the liquid enters a small metal capillary needle usually charged at a few kilovolts (215,216). At the needle tip, a Taylor cone forms due to the high electric field at the needle tip. This cone stretches outwards towards the ground potential from the needle tip, forming a jet, due to repulsive Coulombic forces in the cone structure. The larger droplets form smaller progeny droplets as they are ejected into a plume (depicted in Figure 2‐2A) (215).

However, these droplets are still heavily solvated and have to be stripped of the solvent so that bare ions (containing no solvent) can be analyzed by mass spectrometry. To desolvate the droplets, several engineering advances have been made from heating the electrospray chamber to infusing a counter current drying gas to assist desolvation of the molecules. Three different mechanisms have been proposed for the desorption of the ions from the solvated droplets. The evidence for each of these mechanisms has been reviewed in (216) and have been referred to as ion evaporation model, charge residue model, and chain ejection model (Figure 2‐1B). Once desolvated, the ions are in the gaseous state and are funneled into the mass analyze through a series of focusing elements that make up the ion source device.

39

A B

hν

Matrix(Crystal(La.ce(( C

Matrix(Crystal(La.ce((

Figure 2‐1: Scheme of the mechanism by which ions are transformed into the gaseous state by matrix assisted laser desorption ionization (MALDI). (A) Image from (217) that shows the co‐crystallization of analyte‐matrix lattice. (B) Image from (217) that shows thermal expansion of analyte‐matrix for the formation of ions. Immediately after the laser is fired. (C) Image showing the movement of ions from the surface of the MALDI plate into the mass spectrometer.

40

A

B

Figure 2‐2: Scheme of the mechanism by which ions are transformed into the gaseous state by electrospray ionization. (A) Image from (216) that shows the mechanism by which ions are made from the ESI source. (B) Image from (216) that shows the physical mechanisms suggested for ion formation. IEM (ion evaporation model), CRM (charge residue model), and CEM (chain ejection model).

41

Mass Analyzers–

Once the ions are in the gaseous form they are funneled through a series of offset focusing rings and quadrupolar electrical fields in an evacuated space (under 10‐4 Torr).

The advances in these technologies will not be described here, but have allowed for significantly lower detection limits of analytes of interest through the elimination of neutral molecules that create noise at the ion detector, and better focusing of the ion beam.

After these initial steps in the mass spectrometer, the ions enter the first mass analyzer.

For many tandem mass spectrometers, the first mass analyzer is a quadrupole mass filter.

Quadrupoles are made up of four electrically independent hyperbolic or cylindrical rods. Each rod is linked to RF (radio frequency) and DC (direct current) voltages (Figure 2‐

3A). The combination of the RF and DC voltages creates a time dependent hyperbolic electrical field between the rods, which can be altered very quickly to obtain a stable trajectory for specific ions depending on their mass‐to‐charge ratio (218). The mathematical expression that explains the behavior of ions within a quadrupolar hyperbolic field was first described by Émile Léonard Mathieu in the 1860’s who derived a mathematical function while studying the vibration of elliptical drumheads (219).

The Mathieu equation can then be transformed using the relationship between force, mass and acceleration (F=ma), and be solved to determine values for DC and RF voltages that are required for an ion to undergo a stable transit through the length of the quadrupole mass filter. The mathematical solution for each ion used in commercial mass spectrometers is actually a small area, which has been termed the region of stability

(Figure 2‐3B). These are regions of DC and RF voltages that allow the ion of the specific mass‐to‐charge [m/z] to remain stable in the hyperbolic field. As shown in the diagram, all ions are stable at low RF voltages in which no DC voltage is placed on the poles. The mathematical solutions of individual ions can be seen in Figure 2‐3. Two different scan

42

lines are shown in Figure 2‐3B. As can be seen in the figure, the slope of the scan line determines the resolution of that ion. When all ions are being focused through the quadrupole (total ion mode), it is referred to as an RF only function since all ions are stable at low RF voltages (218).

Quadrupoles are useful mass analyzers for tandem instruments because ions are separated while maintained in a continuous ion beam. However, commercial quadrupoles have only modest resolving power, providing mass accuracy measurements typically ±0.1

Da. The details of the importance of mass accuracy will be discussed later in this chapter.

Each quadrupole also has limitations for the stabilization of ions in the given hyperbolic field. Many commercial quadrupoles have m/z limits upwards of 4000 Da. Therefore, other mass analyzers are usually used to obtain higher mass range data.

The linear time of flight (TOF) mass sector has the potential for high mass accuracy measurements. Originally described and built in the 1950s, the linear TOFs had a resolving power of 20 (220). Resolving power is defined as the m/z of an ion divided by the width of the mass peak at half maximum height. In the first reported, the TOF experiment revealed a two mass unit wide peak measured for the argon ion at m/z 40 (220). TOF mass analyzers accelerate ions with known electrical field strength. This leads to ions with the same charged state to have the same amount of kinetic energy. Therefore, the velocity of the ion is only dependent on the m/z of that ion.

Since the ions have to travel a fixed path length, the time an ion takes to travel from the ion accelerator to the detector is directly proportional to the m/z ratio. The limitation of the mass accuracy is therefore directly related to how accurately time can be measured.

With the development of very sophisticated timing circuits and faster ion detectors, TOF mass analyzers continue to become increasingly accurate even at high m/z ratios. State‐of‐ the‐art TOF mass analyzers have mass accuracies to 0.001 Da to 0.0001 Da, so in the case

43

of phospholipid accurate measurement of 0.1 to 5 ppm. In addition to their mass accuracy, they can record an entire mass spectrum at μsec time scales, which is much faster than other mass analyzers.

Measuring Exact Mass by Mass Spectrometry–

Mass spectrometry measures a mass‐to‐charge ratio, a measurement based on the physical mass scale which is termed exact mass. Exact mass and formula mass differ because exact mass is a based on the mass of the most abundant isotope of carbon being exactly 12.000 Da. Alternatively, formula mass is a chemical scale taking into account the natural isotopes of elements such and 12C and 13C. AB

Figure 2‐3: Scheme of the mechanism by which ions are selected for in the quadrupole mass spectrometer. (A) Image from (218) showing a cross section of a quadrupole with the alternating charges placed on the each rod by the RF and DC voltages. (B) Image from (218) displays solutions from the Mathieu equation for three ions. The area under the m/z describes the RF and DC voltages at which that specific m/z is stable within the quadrupole. Two lines are drawn onto the solutions for the three ions, indicating two alternative lines of stability. The cross‐sectional area of where the lines of stability intersect is shown below.

44

Formula mass is based on the assembly of natural isotopes of carbon being added together (12C , 13C, and 14C) yielding a mass of 12.011 Da. Each naturally occurring isotope, which has been incorporated into the molecule, can be separated by mass spectrometry.

Table 2‐1 shows the exact mass calculation for common organic atoms at their natural isotope abundance. Information using exact mass calculations is critical in the interpretation and identification of molecules in a mass spectrum. Relative to 12C, 1H

(1.007825 Da) has excess mass where as 16O is mass deficient (15.9949 Da). Therefore, the mass defect that is measured for a given molecule provides information about the elemental composition of the given molecule of interest. Mass‐to‐charge is measured accurately enough (e.g. to ±0.001 Da) then elemental composition of that ion can be calculated. High‐resolution mass measurements are desired because they provide higher mass accuracy for more confidence in elemental composition calculations.

Lipid elemental composition consists of numerous carbon, oxygen, hydrogen atoms, and a few heteroatoms like nitrogen, phosphorous, and sulfur. If a molecule were only made of carbon atoms, it would have no mass defect. However, for every hydrogen that the molecule contains an additional mass defect of 0.007825 Da would be added to the mass and for every oxygen a mass of 0.0051 Da would be subtracted. Much of this thesis will focus on fatty acids and glycerophospholipids. A common phospholipid, 1‐palmitoyl‐

2‐oleoyl‐sn‐glycero‐3‐phosphocholine (16:0/18:1‐PC), has an elemental composition of

C42H82NO8P. With the numerous number of hydrogen atoms, the exact mass calculation of this compound comes out to be 759.578 Da, which is significantly higher than the nominal mass. Even larger glycerophospholipids, such as 1,1',2,2'‐tetra‐(9Z‐octadecenoyl) cardiolipin (18:1/18:1/18:1/18:1‐CL), which has a elemental composition of C81H148O17P2 and an exact mass of 1500.999 Da which is about 1 amu higher than nominal mass considering integer values for the mass of all elements, due to the presence of 148

45

hydrogen atoms that generates 149.158 Da in mass. As discussed above, quadrupoles have mass accuracy to the tenths digit, which sometimes leads to the rounding of numbers that are higher or lower than the true exact mass calculation. Later in this thesis, m/z ratios measurements will be rounded to the tenths digit, as a convenience when referring to a specific phospholipid molecular species.

MALDI Imaging Mass Spectrometry (MALDI‐IMS)–

Many techniques, based on optical microscopy, exist to determine localization of biomolecules such as proteins and nucleic acids in tissues, but until recently, techniques have lagged behind to determine spatial location of lipids in tissues. With the advances in soft‐ionization techniques, a recent field has immerged to create four‐dimensional molecular maps of lipids across biological tissues. Many of the advances in the MALDI‐IMS technique originated from work developed for proteins in the laboratory of Dr. Richard M.

Caprioli, however since the development of the technique, extensive validation in the field of lipids has also been performed (213,221,222).

Table 2‐1: Exact Mass and Natural Isotope Calculations for Common Organic Atoms

Atom# Exact#Mass# Natural#Isotope# Abundance#%# 1H# 1.007825# 99.985#

2H# 2.014101# 0.015#

12C# 12# 98.90#

13C# 13.00335# 1.10#

14N# 14.00307# 99.63#

15N# 15.000108# 0.37#

16O# 15.9949# 99.76#

17O# 16.99913# 0.04#

18O# 17.99916# 0.2# .

46

To collect tissue images, tissues are collected and frozen in a modified optimal cutting temperature (OCT) medium. The tissues are section at a thickness of 10‐20 μm and placed directly onto glass cover slips and then placed on MALDI plates or placed directly onto metal or glass MALDI plates (214). The surface of the tissue is coated by various techniques including sublimation (213,222) used in this thesis, by a matrix compound DBA for positive and DHAP for negative ion analysis) to enable molecules to be ionized by

MALDI.

A workflow of how the data is collected and converted is shown in Figure 2‐4. To expand on what is shown in the figure, the MALDI laser is programmed to raster across the tissue at a preset step size (usually 10‐50 μm). At each step the mass spectrum is recorded at that x‐y coordinate. The laser continues to collect individual mass spectra across the tissue at each movement of the laser (our laboratory has used a MALDI‐qTOF shown in

Figure 2‐4). These individual mass spectra can be reconfigured for the x‐y coordinates in which they were collected and the intensity of a given ion converted to a color‐coded pixel.

Collectively these pixels represent the relative signal intensity of a given ion, compared to that same ion captured the in mass spectra at other areas of the tissue. This relative intensity comparison of ions has enabled one to picture localization of specific lipids (as their positive or negative desorbed molecular ions) localized in certain regions of tissues.

Also, use of the MALDI‐qTOF instrument enables MS/MS experiments to be run on specific ions being generated at a local region of the tissue.

47

hν 355&nm&

MALDI&plate&

Pixels

Figure 2‐4: Workflow of how MALDI‐IMS (imagine mass spectrometry) is able to obtain spatially distribution of lipids in a tissue. Tissue sections (10‐20 μm) is sectioned onto a MALDI plate. The matrix is sublimated to coat the surface of the tissue. The MALDI laser then rasters across the tissue collection mass spectrum. Images shown above are approximately 50 μm resolution, but newer laser can obtain spatial resolution under 10 μm. A single ion is the collected, and its relative intensity can be transformed into a corresponding pixel. The intensity of the ion can be compared to the other mass spectra collected across the tissue. The tissue in the figure is then colored coded to represent the intensity of the specific ion of interest. In this figure blue represents higher ion intensity and red lower ion intensity.

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Tandem Mass Spectrometry–

Many mass spectrometers now use multiple mass analyzers, also termed sectors, in tandem to further structurally characterize ions generated by either electrospray or

MALDI. Our laboratory predominately uses two different tandem mass spectrometers: quadrupole‐time of flight (qTOF) and triple quadrupole mass spectrometers. Although the last mass analyzer differs, similar data can be collected from both of these instruments.

The tandem mass spectrometry experiments performed in this work will be described in detail using the triple quadrupole instrument which is depicted in Figure 2‐5, but experiments are directly applicable to the qTOF.

The classic mass spectrometry experiments have been with the use of one mass analyzer to observe all of the ionizable molecules in the sample. This technique utilizes a single quadrupole and ions are sequentially filtered (scanned) across a predetermined

(m/z) range. In the triple quadrupole, the first quadrupole mass filter is typically used to carry out this scanning through the m/z range. The second quadrupole is used as a collision cell with an RF‐only electric field. The third quadrupole is used as a mass filter for ions that exit the collision cell. In the diagram in Figure 2‐1, Q1 is displayed as the scanning quadrupole mass filter, but the third quadrupole can also be used as the scanning quadrupole (Q3 scans). During this scan, two quadrupoles focus all ions in the RF‐only mode while one quadrupole filters ions. This technique coupled to liquid chromatography

(LC‐MS) is very powerful in quickly identifying many different analytes in a sample in a less targeted fashion.

Commercial quadrupoles have only modest resolving power; therefore it is difficult to know the exact elemental composition based on the measured m/z ratio. With coupling to chromatography, assumptions can be made about the molecular composition of the molecule based on HPLC retention time. However without tandem mass spectrometry

49

experiments it is difficult to know much about the structural characteristics of the molecule. To gain insight, collision induced decomposition [CID] in the collision cell

(quadrupole 2) is used.

Collision cells use inert gases, such as nitrogen gas (N2), to fill the second quadrupole (Q2) of triple quadrupole instrument (Figure 2‐5). When the ions exit Q1, they are provided additional translational energy at the entrance of Q2 (collision energy). Once they enter the second quadrupolar region they begin to collide with the neutral gas. When the ion collides with the neutral gas molecule, the translational energy (velocity) given to the ion is converted into vibrational energy that eventually exceeds the covalent bond energy between two atoms, causing them to break or dissociate bonds causing the ion to break into smaller fragments. Ultimately, the molecule relaxes from the vibrationally excited state. The fragmentation of a molecule is dependent on the parameters of the instrument and on structural features of the molecule. However, some fragment ions produced can be unique identifiers of the structural groups. This unique pattern for each molecule is what makes tandem mass spectrometry coupled to liquid chromatography (LC‐

MS/MS) such a powerful technique in the analysis of analytes lipids as well as other biomolecules.

Product ion scans are used to determine the unique fragmentation of each analyte of interest. In this scan a single parent ion is selected by Q1 then enters the collision cell, all other ions are unstable in the first quadrupole sector and do not make it into the collision cell. Once in the collision cell, collisional activation takes place to form product ions.

Quadrupole 3 is then used to scan a m/z range to detect product ions formed and their abundance is determined using an electron multiplier detector.

Additional experiments that can be performed by the triple quadrupole mass spectrometer are precursor ion, neutral ion scanning, and multiple reaction monitoring

50

scans (MRM). These experiments use Q1 and Q3 as mass filters and Q2 as a collision cell to induce fragmentation. The goal of a precursor ion scan is to determine what molecules make a given product ion. Therefore, Q1 scans through a range of m/z ratios, and after the ion undergoes collision‐induced decomposition. Quadrupole 3 is set to select only one product ion. The data output then only displays ions that were scanned in Q1 that produced the specific product ion selected. This scan is often used to identify lipids of a certain class, for example any lipid that contains a phosphocholine head group will yield a positive ion product at m/z 184 after CID. Neutral loss scanning involves scanning both Q1 and Q3 to identify ions that produce a mass difference that corresponds to the mass of the neutral fragment found by CID.

The final scan type that was frequently used during this thesis work was multiple reaction monitoring (MRM). In this scan type, both Q1 and Q3 are set to a specific m/z ratio and the collision cell used to fragment ion selected by Q1. Q3 monitors for a specific product ion derived after CID. Once a molecule of interest has a known collisional behavior in terms of resultant product ions, this selective scan for the precursor‐product relationship is used as a specific criterion to identify an analyte. The quadrupoles dwell for a longer period of time (10‐100 msec) on a single precursor ion resulting in increased sensitivity of each measured analyte. An example of a common MRM transition for lipids would be the parent phospholipid ion at m/z 758 (16:0/18:1‐PC [M‐H]‐) being selected for in quadrupole 1, and the a fatty acyl carboxylate ions being measured in Q3 at m/z 255 or

281. These are referred to in this thesis as MRM ion transitions m/z 758281 and m/z

758255.

51

Q1#Q2# Q3#

O

O O P O O O OH O Q1#Scan#

Scans#All#Ions# No#Fragmenta

O

O O P O O O OH O MRM#Scan#

O

O

Selects#One#Ion# Fragmenta

O

O O P O O O OH Precursor#Ion# O

O

O

Scans#All#Ions# Fragmenta

O

O O P O O O OH O

O

O O P O O O Product#Ion# OH OH

O

Fragmenta

Figure 2‐5: Tandem quadrupole experiments performed during the course of this thesis work. The scan type is listed in the left had column, and each subsequent column corresponds to the action of each of the quadrupoles during that scan type.

52

The expanding technologies in mass spectrometry have allowed the field of lipid biochemistry to expand significantly. Few techniques are available for the analysis of lipids, especially at the sensitivity needed to detect novel bioactive lipid compounds. The field of soft‐ionization mass spectrometry will continue to expand the knowledge of lipid biochemistry as technologies in mass spectrometry become more readily available.

53

CHAPTER III

MEASUREMENT OF LYSOPHOSPHOLIPID ACYLTRANSFERASE ACTIVITY USING

SUBSTRATE COMPETITION

Introduction–

Enzyme activity assays have been a traditional biochemical means by which to assess the presence of a particular protein in the biological extract. These assays provide insight into the overall performance of this enzyme in terms of its synthetic capacity and substrate specificity. LPAT (lysophospholipid acyltransferase) activity assays which employ two substrates are often performed by exposing a tissue extract, containing the enzyme of interest, to a pair of pure substrates (one lysophospholipid and one acyl‐CoA ester), and measuring the reaction rate parameters for substrate conversion to product.

The result of this experiment is then compared to the results from an identical set of experiments, but with a different substrate. Eventually, a picture emerges of the preferred substrates. Thus, many substrates need to be individually studied. A separate concern with this approach is that any competition between substrates for the enzyme is not revealed

(192,223,224). If there are multiple enzymes that exert very similar biochemical activity and the enzymes utilizes multiple substrates, this greatly expands the total number of experiments that need to be carried out by this single substrate enzyme activity strategy.

An alternative biochemical assay for LPAT activity was developed that involves addition of a mixture of substrates to a microsomal extract containing this enzymatic activity. The products of the reaction reveals which different enzymatic activities are present (225). Unique phospholipid products are identified and quantified using liquid chromatography tandem mass spectrometry. This strategy of a substrate choice enzymatic assay is particularly useful for study of the LPATs. LPAT enzymes utilize fatty acyl‐CoA ester and lysophospholipids to make the final phospholipid product, which would require

54

a large number of single enzyme assays in a single experiment. This substrate choice enzyme activity assay presents microsomal enzymes with a mixture of different substrates for the same reaction, with the profile of product abundance indicating substrate preference. While absolute quantity of each product made by the substrate choice assay would differ from the absolute quantity of the same product in the single substrate assay, the dual substrate choice assay does reveal potential for the complexity of product formation that occurs in cells. The dual substrate choice assay is easily carried out due to advances in electrospray tandem mass spectrometry. In this chapter, experiments are presented that were used to determine the conditions for the dual substrate choice assay using microsomal extracts containing LPATs expressed in mammalian cells. The activities of the LPATs in microsomes were tested in a dual substrate choice model and compared to the results with the more traditional means of determining substrate specificity and enzymatic rate.

Experimental Procedures–

Materials– Fatty acyl‐CoA esters and phospholipids were from Avanti Polar Lipids

(Alabaster, AL): tetradecanoyl‐coenzyme A (myristoyl; 14:0‐CoA), hexadecanoyl‐coenzyme

A (palmitoyl; 16:0‐CoA), octadecanoyl‐coenzyme A (stearoyl; 18:0‐CoA), (9Z)‐ octadecenoyl‐coenzyme A (oleoyl; 18:1‐CoA), (9Z,12Z)‐octadecadienoyl‐coenzyme A

(linoleoyl; 18:2‐CoA), (5Z,8Z,11Z,14Z)‐eicosatetraenoyl‐coenzyme A (arachidonoyl; 20:4‐

CoA), (5Z,8Z,11Z,14Z,17Z)‐eicosapentaenoyl‐coenzyme A (20:5‐CoA),

(4Z,7Z,10Z,13Z,16Z,19Z)‐docosahexaenoyl‐coenzyme A (22:6‐CoA), 1‐(10Z)‐ heptadecenoyl‐2‐hydroxy‐lysophosphatidic acid (17:1‐LPA), 1‐(10Z)‐heptadecenoyl‐2‐ hydroxy‐lysophosphatidylcholine (17:1‐LPC), 1‐(10Z)‐heptadecenoyl‐2‐hydroxy‐ lysophosphatidylethanolamine (17:1‐LPE), 1‐(10Z)‐heptadecenoyl‐2‐hydroxy‐

55

lysophosphatidylglycerol (17:1‐LPG), 1‐(10Z)‐heptadecenoyl‐2‐hydroxy‐ lysophosphatidylinositol (17:1‐LPI), 1‐(10Z)‐heptadecenoyl‐2‐hydroxy‐ lysophosphatidylserine (17:1‐LPS), 1‐(9Z)‐octadecenoyl‐2‐hydroxy‐lysophosphatidic acid

(18:1‐LPA), 1‐(9Z)‐octadecenoyl‐2‐hydroxy‐lysophosphatidycholine (18:1‐LPC), 1‐(9Z)‐ octadecenoyl‐2‐hydroxy‐lysophosphatidyserine (18:1‐LPS), (2R)‐[2H31]‐3‐hexadecanoyl‐2‐

(9Z)‐octadecenoyl‐PA ([2H31]‐POPA), (2R)‐[2H31]‐3‐hexadecanoyl‐2‐(9Z)‐octadecenoyl‐PC

([2H31]‐POPC), (2R)‐[2H31]‐3‐hexadecanoyl‐2‐(9Z)‐octadecenoyl‐PE ([2H31]‐POPE), (2R)‐

[2H31]‐3‐hexadecanoyl‐2‐(9Z)‐octadecenoyl‐PG ([2H31]‐POPG), (2R)‐[2H31]‐3‐hexadecanoyl‐

2‐(9Z)‐octadecenoyl‐PI ([2H31]‐POPI), (2R)‐[2H31]‐3‐hexadecanoyl‐2‐(9Z)‐octadecenoyl‐PS

([2H31]‐POPS), 1‐heptadecanoyl‐2‐eicosatetraenoyl‐sn‐glycero‐3‐phosphate (17:0/20:4‐

PA), 1‐heptadecanoyl‐2‐(5Z,8Z,11Z,14Z)‐eicosatetraenoyl‐phosphatidylcholine (17:0/20:4‐

PC), 1‐heptadecanoyl‐2‐(5Z,8Z,11Z,14Z)‐eicosatetraenoyl‐phosphatidylethanolamine

(17:0/20:4‐PE), 1‐heptadecanoyl‐2‐(5Z,8Z,11Z,14Z)‐eicosatetraenoyl‐ phosphatidylglycerol (17:0/20:4‐PG), 1‐heptadecanoyl‐2‐(5Z,8Z,11Z,14Z)‐ eicosatetraenoyl‐phosphatidylinositol (17:0/20:4‐PI), and 1‐heptadecanoyl‐2‐

(5Z,8Z,11Z,14Z)‐eicosatetraenoyl‐phosphatidylserine (17:0/20:4‐PS). Dulbecco’s modified

Eagle’s medium (DMEM) was purchased from Corning Cellgro (Manassas, VA). RAW 264.7 cells were obtained from ATCC (Manassas, VA). EDTA‐free protease inhibitor cocktail was purchased from Roche (Madison, WI). All other chemicals and solvents were purchased through Fisher Scientific (Pittsburg, PA).

Cell Culture and Microsome Preparation– RAW 264.7 cells were cultured in DMEM

(with 4.5 g/L glucose and 100 µM sodium pyruvate) supplemented with 10 % heat‐ inactivated FBS. The cells were grown in humidified air with 5 % CO2 at 37 °C. For microsomal preparations, cells were pelleted at 300 x g, resuspended in homogenization buffer (50 mM Tris‐HCl, pH 7.4, 250 mM sucrose, 1 mM EDTA, 20% (w/v) glycerol, and

56

protease inhibitor cocktail), and lysed using a Sonics Vibra‐Cell probe sonicator (Newtown,

CT). Whole cells and cellular debris were pelleted at 12,000 x g for 20 min at 4 °C. The supernatant was transferred to an ultracentrifuge tube and centrifuged at 100,000 x g for

60 min at 4 °C. The microsomal pellet was resuspended in assay buffer (10 mM Tris‐HCl, pH 7.4, 150 mM NaCl, 1 mM EDTA), protein amount was determined using the bicinchoninic acid assay (Thermo Scientific, Rockford, IL), and microsomes were stored at ‐

20 °C until used.

Single Choice LPAT Assay– Stock solutions were made as follows: 60 µM arachidonoyl‐CoA in 100 % methanol; 200 µM lysophospholipids in assay buffer (prepared and sonicated immediately prior to the assay); 1.25 mM fatty acid‐free bovine serum albumin (BSA) in water; and internal standard mixture of 10 ng/µl each of

[2H31]16:0/18:1‐PA, [2H31]16:0/18:1‐PC, [2H31]16:0/18:1‐PE, [2H31]16:0/18:1‐PG,

[2H31]16:0/18:1‐PI, and [2H31]16:0/18:1‐PS in methanol. The final concentrations of the reaction components were: 10 µg total protein from microsomes, 3 µM LPC, LPE, or LPS, 3

µM arachidonoyl‐CoA, and 12.5 µM BSA, in assay buffer to a total volume of 200 µl. The

CoA ester solution was prepared in methanol, this solution was used during the assay which led to a concentration of 5 % methanol in the final reaction. When compared with reactions with no methanol, this 5 % level resulted in only minor differences found in 2 of the 48 products. The acyltransferase assay was performed at 37 °C for 10 min. The reaction was stopped with the addition of 750 µl of methanol: chloroform (2:1, v/v), internal standard mixture (2.5 µl) added and products were extracted by the Bligh and

Dyer method (226). After the samples were dried under a stream of nitrogen, they were resuspended in 100 µl of 75 % solvent A (isopropanol: hexanes 3:4, v/v) and 25 % solvent

B (isopropanol: hexanes: water 3:4:0.7, v/v/v, containing 5 mM ammonium acetate).

57

Samples were analyzed by liquid chromatography coupled to tandem mass spectrometry

(LC/MS/MS) as described below.

Dual Substrate Choice Assay– Acyltransferase activity was tested in microsomal preparations as described for the single substrate choice except for the following alterations. Stock solutions of reaction materials were made as follows: 6 µM, 20 µM, or 60

µM equimolar mixture of eight acyl‐CoAs (14:0, 16:0, 18:0, 18:1, 18:2, 20:4, 20:5 and 22:6) in methanol, 20 µM or 200 µM equimolar mixture of six lysophospholipids (LPA, LPC, LPE,

LPG, LPI and LPS) in assay buffer. Final lysophospholipid concentrations used were 0.3 µM,

1 µM, 3 µM or 10 µM, as indicated, with final concentrations of 50 ng/µl total protein from microsomes, 3 µM of each fatty acyl‐CoA ester, and 12.5 µM BSA, in a total volume of 200

µl. To assay different microsomal protein amounts, the assay was performed at final concentrations of 3 µM lysophospholipid mix, 3 µM fatty acyl‐CoA ester mix, and 12.5 µM

BSA, while using microsomal protein concentrations of from 5 ng/µl, 15 ng/µl and 50 ng/µl. To assay different fatty acyl CoA ester concentrations, 50 ng/µl total protein from microsomes, 3 µM of each lysophospholipid, and 12.5 µM BSA, was used while fatty acyl‐

CoA ester concentrations were varied to either 0.3 µM, 1 µM, or 3 µM each. The final volume of methanol was kept constant. To assay different time points the components were at final concentrations of 50 ng/µl total protein from microsomes, 3 µM of each lysophospholipid, 3 µM of each fatty acyl‐CoA ester, and 12.5 µM BSA. The assay was performed at 37 °C for 0 min, 1 min, 3 min, 10 min or 30 min. Samples were analyzed by liquid chromatography‐ tandem mass spectrometry (LC‐MS/MS) as described below.

Liquid Chromatography/Mass Spectrometry– For normal phase separation, samples were injected onto an Ascentis‐Si HPLC column (150 x 2.1 mm, 5 µm; Supelco with

Sigma Aldrich, St. Louis, MO) at a flow rate of 0.2 ml/min at 25 % solvent B. Solvent B was maintained at 25 % for 5 min, increased to 60 % over 10 min, and then to 95 % over 5 min.

58

The system was held at 95 % B for 20 min prior to re‐equilibration at 25 % for 14 min. For reversed phase separation, solvent C was methanol/acetonitrile/water, 60/20/20 (v/v/v), containing 2 mM ammonium acetate, and solvent D was methanol containing 2 mM ammonium acetate. The samples were injected onto an Ascentis‐C18 HPLC column (150 x

2.1 mm, 5 µm; Supelco) at a flow rate of 0.2 ml/min at 75 % solvent D. Solvent D was maintained at 75 % for 1 min and increased to 98 % over 5 min. The system was held at 98

% D for 20 min prior to re‐equilibration at 75 % for 10 min. Phospholipid products of the

LPAT assay were measured using an API3200 triple quadrupole mass spectrometer (AB

Sciex, Torrance, CA) in negative ion mode using multiple‐reaction‐monitoring (MRM) of the precursor‐product transitions shown in Table 3‐1. Quantification was performed using

AB Sciex MultiQuant software and using 17:0/20:4‐PA, 17:0/20:4‐PC, 17:0/20:4‐PE,

17:0/20:4‐PG, 17:0/20:4‐PI and 17:0/20:4‐PS reference standards for dilution curve analysis, as described previously (227).

59

Table 3‐1: Multiple Reaction Monitoring (MRM) Transitions Used for Detection of Products Made in the Substrate Choice Assay

Phosphatidylinositol (PI) Q1 m/z Q3 m/z2 17:1/16:0 821.5 255.2 17:1/14:0 793.5 227.2 Phosphatidic acid (PA) Q1 m/z Q3 m/z1 17:1/18:0 849.5 283.2 17:1/16:0 659.5 267.2 17:1/18:1 847.5 281.2 17:1/14:0 631.5 267.2 17:1/18:2 845.5 279.2 17:1/18:0 687.5 267.2 17:1/20:4 869.5 303.2 17:1/18:1 685.5 267.2 17:1/22:6 893.5 327.2 17:1/18:2 683.5 267.2 17:1/20:5 867.5 301.2 2 17:1/20:4 707.5 267.2 Phosphatidylserine (PS) Q1 m/z Q3 m/z 17:1/22:6 731.5 267.2 17:1/16:0 746.5 267.2 17:1/20:5 705.5 267.2 17:1/14:0 718.5 267.2 Phosphatidylcholine (PC) Q1 m/z Q3 m/z1 17:1/18:0 774.5 267.2 17:1/16:0 804.6 267.2 17:1/18:1 772.5 267.2 17:1/14:0 776.6 267.2 17:1/18:2 770.5 267.2 17:1/18:0 832.6 267.2 17:1/20:4 794.5 267.2 17:1/18:1 830.6 267.2 17:1/20:5 792.5 267.2 17:1/18:2 828.6 267.2 17:1/22:6 818.5 267.2 3 m/z m/z 17:1/20:4 852.6 267.2 Internal Standards Q1 Q3 2 17:1/22:6 876.6 267.2 [ H31]16:0/18:1-PA (sn1) 704.7 286.2 2 17:1/20:5 850.6 267.2 [ H31]16:0/18:1-PC (sn1) 849.8 286.2 2 m/z m/z1 [ H31]16:0/18:1-PE (sn1) 747.7 286.2 Phosphatidylethanolamine (PE) Q1 Q3 2 17:1/16:0 702.5 267.2 [ H31]16:0/18:1-PG (sn2) 778.7 281.2 [2H ]16:0/18:1-PI (sn2) 866.8 281.2 17:1/14:0 674.5 267.2 31 [2H ]16:0/18:1-PS (sn1) 791.7 286.2 17:1/18:0 730.5 267.2 31 4 m/z m/z 17:1/18:1 728.5 267.2 Endogenous Phospholipids Q1 Q3 17:1/18:2 726.5 267.2 16:0/18:1-PA (16:0 anion) 673.6 255.2 17:1/20:4 750.5 267.2 18:1/18:0-PA (18:0 anion) 701.6 283.2 17:1/22:6 774.5 267.2 16:0/18:1-PC (18:1 anion) 818.5 281.2 17:1/20:5 748.5 267.2 18:1/18:1-PC (18:1 anion) 844.5 281.2 Phosphatidylglycerol (PG) Q1 m/z Q3 m/z2 18:1/18:0-PC (18:1 anion) 846.5 281.2 18:1/18:1-PE (18:1 anion) 742.5 281.2 17:1/16:0 733.5 255.2 18:1/18:0-PE (18:1 anion) 744.5 281.2 17:1/14:0 705.5 227.2 18:1/18:1-PG (18:1 anion) 773.5 281.2 17:1/18:0 761.5 283.2 16:0/18:1-PI (18:1 anion) 835.5 281.2 17:1/18:1 759.5 281.2 18:1/18:1-PI (18:1 anion) 861.5 281.2 17:1/18:2 757.5 279.2 18:1/18:1-PS (18:1 anion) 773.5 281.2 17:1/20:4 781.5 303.2 16:0/18:1-PS (18:1 anion) 835.5 281.2 17:1/22:6 805.5 327.2 18:1/14:0-PA (18:1 anion) 861.5 281.2 17:1/20:5 779.5 301.2 18:1/14:0-PC (18:1 anion) 773.5 281.2 1"Q3"m/z"corresponds"to"the"loss"of"the"sn&1"acyl"anion." 18:1/14:0-PS (18:1 anion) 835.5 281.2 2 " "Q3"m/z"corresponds"to"the"loss"of"the"sn&2"acyl"anion. 18:1/22:6-PA (18:1 anion) 861.5 281.2 3"Q3"m/z"is"indicated"in"parenthesis"in"the"table." 4"Q3"m/z"is"indicated"by"the"specific"loss"of"a"carboxylate" 18:1/22:6-PC (18:1 anion) 861.5 281.2 anion"in"parenthesis"in"the"table."The"glycerol"position"of" 16:0/22:6-PS (16:0 anion) 760.5 255.2 the"acyl"chain"was"not"known"for"endogenous"lipids."

60

Results–

Normal‐Phase‐HPLC‐MS/MS Detection and Quantitation of Phospholipid Products–

The goal of the present chapter was to validate the dual choice assay by comparing it directly to the more traditional enzyme assay. This work allowed for the further development of the method as a tool to analyze the mixture of lysophospholipid acyltransferase activities present on the microsomes of the murine macrophage‐like cell

(RAW 264.7). The fatty acyl‐CoA esters chosen contained the acyl chains more commonly found in cellular membranes. All of the fatty acyl‐CoA esters in this thesis will be referred to using the carbon number: double bond number‐CoA nomenclature (e.g. 20:4‐CoA).

Three fatty acyl CoA esters of biological interest (20:4‐, 20:5‐ and 22:6‐CoA) were incorporated into the assay because their fatty acids forms are precursors for bioactive lipids along with five other common fatty acyl chains (14:0‐, 16:0‐, 18:0‐, 18:1‐, and 18:2‐

CoA). The lysophospholipids used in the assay all contained the same sn‐1 acyl chain, 17:1, which reduced the background signal from endogenous phospholipids present in the microsomal preparations.

The effectiveness of normal‐ and reverse‐phase liquid chromatography was evaluated by the ability to resolve both endogenous and newly synthesized phospholipids.

After chromatographic separation the HPLC effect was directed into an electrospray ion source and the resulting ions detected with a triple quadrupole mass spectrometer operated in the MRM mode. A typical normal‐phase chromatography profile of the products of the assay is presented in Figure 3‐1. With the ability to resolve phospholipids by their head group polarities in all subsequent experiments, normal phase separation was used over reverse phase chromatography. In those cases where chromatographic resolution overlapped (PE/PI and PA/PS), MRM mass spectrometry was used detect the unique molecular and fragment ions of each phospholipid products.

61

PI" PS" 175 100%" PA PC" AB150 PC 125 PE PA" PG" 100 PG PE" PI 75 PS 50 Rela3ve"Intensity" 25

Analyte/Internal Std. Area 0 2" 6" 10" 14" 18" 22" 26" 30" 34" 38" 0 25 50 75 100 125 150 Time,"min" phospholipid (ng)

Figure 3‐1: Separation and quantitation of phospholipids. A) Normal‐phase chromatographic separation of phospholipids by head group. B) Standard curves for the conversion of signal intensity to absolute quantity using a series of 17:0/20:4‐PL compared to [2H31]‐PL internal standards. Phosphatidylglycerol (PG), phosphatidylinositol (PI), phosphatidylethanolamine (PE), phosphatidylserine (PS), and phosphatidic acid (PA), and phosphatidylcholine (PC).

The deprotonated [M‐H]‐ precursor ions were selected as the target ions to quantitate all classes except for PC, in which case the acetate adduct [M+CH3COO]‐ ions were used (228). Upon collision‐induced decomposition, all precursor ions yielded abundant product ions corresponding to both the sn‐1 and sn‐2 carboxylate anions. In preliminary experiments using standards, the sn‐1 anion was found to be more abundant than the sn‐2 anion for PA and PS, whereas the sn‐2 anion was more abundant in the cases of PC, PE, PI and PG under the conditions used. Therefore, the logical choice was to use the most abundant fragment ion except for in the case of PC and PE. The transitions to the sn‐1 anion (m/z 267) were used for PC and PE because endogenous plasmenyl (18:0p‐ containing) PC and PE species are isobaric with the 17:1‐containing products derived from the substrates that were newly synthesized. Endogenous plasmenyl phospholipids do not generate a m/z 267 product ion. Therefore, the transitions to this sn‐1 acyl anion yielded robust signals that were not obscured by high background. Most of the m/z transitions

62

were specific for each phospholipid measured, with two exceptions: m/z 747→281

([2H31]16:0/18:1‐PE or 16:0/18:1‐PG) and m/z 774→267 (17:1/22:6‐PE or 17:1/18:0‐PS).

In these cases, quantitation was still possible because the corresponding phospholipids were resolved by normal‐phase chromatography. Quantitation of each of the phospholipid reaction products was performed by calculating the ratio of the integrated area of the phospholipid product peak (analyte) to that of the corresponding internal standard (IS).

Then, reference standard dilution curves were used to calculate the amount of phospholipid product made, as previously described (227). The calibration curves for the

17:0/20:4 phospholipid species of each of the six classes are presented in Figure 3‐1B.

Determination of Enzymatic Parameters of the Dual Substrate Choice Assay in

Microsomes from RAW 264.7 Cells– Single choice assay strategies to test LPAT activity used high levels of substrates with long incubation times to assay LPAT activity (223). In testing the dual substrate choice assay, reaction conditions were optimized to ensure that they were within or close to the linear range, to better appreciate the substrate preferences of the LPATs present in the sample.

Four major variables that dictate product formation were analyzed, namely time, lysophospholipid concentration, fatty acyl‐CoA concentration, and total microsomal protein. These reactions conditions were used because they were in the linear range of the reaction or allowed for detectable and reproducible levels of the phospholipid products. To illustrate the reaction kinetics for the dual substrate choice assay the formation of

17:1/20:4‐PC is presented in Figure 3‐2. The standard reaction conditions decided upon for all experiments were 3 µM of each lysophospholipid and fatty acyl‐CoA, 10 µg of microsomal protein, and a 10‐minute incubation.

63

7.5 A 6 B 5

5.0 4 3

2.5 2 /µg(microsomal(protein(

/µg(microsomal(protein( 1 pmol

pmol 0.0 0 0 10 20 30 0 2 4 6 8 10 Time((min)( Lysophospha7dylcholine((µM)( 3 C 30 D

2 20 ( pmol 1 10 /µg(microsomal(protein(

pmol 0 0 0.0 0.5 1.0 1.5 2.0 2.5 3.0 0 2 4 6 8 10 Arachidonoyl;CoA((µM)( Microsomal(Protein((µg)(

Figure 3‐2: Reaction conditions of the dual substrate choice acyltransferase assay. Microsomes from RAW 264.7 cells were used to assay for LPAT activity using a mix of six lysophospholipids and eight acyl‐CoA esters as described in the Methods section. Four variables were changed as shown in the different panels: time (A), lysophospholipid concentration (B), fatty acyl‐CoA concentration (C), and microsomal protein amount (D). Only the results for incorporation of arachidonoyl chain into phosphatidylcholine are shown, but all 48 possible products were quantitated using LC‐MS/MS. Three independent microsomal preparations were assayed in duplicate, and data shown are average ± SEM.

64

The product formation was clearly dependent on reaction time, concentrations of either substrate, and amount of protein. The standard reaction conditions used lysophospholipid concentrations that were significantly lower than concentrations used in previous biochemical assays. In part this was due to the inherent sensitivity of mass spectrometry but also it was felt important to use levels below the reported critical micellar concentration of lysophospholipids (229). The absolute amount of microsomal protein present in the assay was used to normalize the amount of each newly synthesized phospholipid product, as is commonly done in enzymatic assays.

The Measurement of Endogenous Microsomal Phospholipids from the RAW 264.7 as a Means for Normalization– Two common acyl compositions (16:0/18:1 and 18:1/18:1) found in most phospholipid classes were also measured for each phospholipid class (PA,

PC, PE, PG, PI, and PS) before and after the enzyme reaction. This was conducted to check for possible changes in endogenous microsomal phospholipids. Under each reaction parameter stated above (time, CoA ester concentration, lysophospholipid concentration, and microsomal protein content) the endogenous lipids were measured. Each of the phospholipid classes (16:0/18:1‐ and 18:1/18:1‐phospholipids) showed excellent correlation with the amount of microsomal protein (Figure 3‐3G‐H), which suggested that endogenous phospholipids could be used as an alternative normalization factor. No changes in endogenous lipids were observed over the incubation time or with different substrate concentrations (Figure 3‐3A‐F). A small increase in endogenous 16:0/18:1‐PC was observed, but when statistical analysis was performed between time 0 and time 3 min, a p‐value of 0.334 was found (Figure 3‐3A). When comparison was made for the formation of 17:1/20:4‐PC at time 0 and time 3 min a p‐value of 0.0006 was found (Figure 3‐2A), thus suggesting that the slight increase in 16:0/18:1‐PC of the 3 min time period was probably due to the error of microsomal addition to the samples.

65

16:0/18:1-PC 18:1/18:1-PC 16:0/18:1CPC* 18:1/18:1CPC* 10 AB15 8 10 6

4 5 2 /µg*microsomal*protein* /µg*microsomal*protein* pmol

0 pmol 0 0 10 20 30 0 10 20 30 Time*(min)* Time*(min)* 16:0/18:1-PC16:0/18:1CPC* 18:1/18:1CPC*18:1/18:1-PC 10 CD10 8 8

6 6

4 4

/µg*microsomal*protein* 2 2 /µg*microsomal*protein* pmol 0 pmol 0 0 1 2 3 0 1 2 3 Fa1y*Acyl*CoA*ester*(µM)* Fa1y*Acyl*CoA*ester*(µM)* 18:1/18:1-PC 18:1/18:1-PC 16:0/18:1CPC* 18:1/18:1CPC* 10 EF10 8 8

6 6

4 4

/µg*microsomal*protein* 2 2 /µg*microsomal*protein* pmol

0 pmol 0 0 2 4 6 8 10 0 2 4 6 8 10 Lysophospholipid*(µM)* Lysophospholipid*(µM)* 16:0/18:1CPC*16:0/18:1-PC 18:1/18:1CPC*18:1/18:1-PC 70 GH100 60 80 50 * 40 * 60 pmol 30 pmol 40 20 20 10 0 0 0 2 4 6 8 10 0 2 4 6 8 10 Microsomal*Protein*(µg)* Microsomal*Protein*(µg)*

Figure 3‐3: Endogenous microsomal phospholipids analyzed during the dual substrate choice acyltransferase assay. Microsomes from RAW 264.7 cells were used to assay for LPAT activity using a mix of six lysophospholipids and eight acyl‐CoA esters as described. The four variables indicated were changed as shown in the different panels. The phospholipids containing 16:0 and 18:1 chains or two 18:1 chains were quantitated. Three independent microsomal preparations were assayed in duplicate, and data shown are average ± SEM.

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The Products of the Dual Substrate Choice Assay Reflect the Presence of Many

LPATs in RAW 264.7 Microsomes– After determining optimal reaction conditions for the

LPATs present in microsomes, this assay was used to characterize the enzymatic activities in these microsomes. Figure 3‐4 shows the forty‐eight products analyzed after a 30‐min incubation. The data presented highlights different fatty acyl substituents at the phospholipid sn‐2 position, supporting the presence of several LPATs that specifically incorporated the specified acyl chain into each phospholipid class. In the PA class, most acyl chains were incorporated at high levels, with the exception of 20:5. The preferred acyl chains were 16:0 and 18:2, followed by 14:0 and 18:1. These results are consistent with the possible expression and activity of LPAAT1, LPAAT2 and LPAAT3 (162,196).

For lysophosphatidylcholine acyltransferase, the preferred acyl chain was 20:4.

There was also robust incorporation of 18:2 and 20:5 chains, consistent with the activity of

MBOAT5/LPCAT3 (162,192,196). The incorporation of saturated and monounsaturated chains suggested the activities of LPCAT1, LPCAT2 and/or LPCAT4 activity (162,196). The incorporation of 18:2 and 20:4 chains into PE was characteristic of the activity of LPEAT2

(162,196). LPEAT1 has been reported to incorporate 18:1‐CoA into LPE and LPS

(162,184,192,196). However, both of these PE species were produced at a minimal level, suggesting negligible activity of this enzyme in RAW 264.7 cells. The PS products generated were possibly a result of the LPSAT activity of MBOAT5/LPCAT3 (192). The 20:4 and 20:5 incorporation into PI was consistent with the activity of MBOAT7/LPIAT1 (192). Finally, there was not a robust production of any of the molecular species measured for PG, which may be due to low levels of LPGAT present in the RAW 264.7 cells.

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14:0 16:0 18:0 18:1 PA( 18:2 20:4 20:5 22:6 14:0 16:0 18:0 18:1 PC( 18:2 20:4 20:5 22:6 14:0 16:0 18:0 18:1 PE( 18:2 20:4 20:5 22:6 14:0 16:0 18:0 PG( 18:1 18:2 20:4 20:5 22:6 14:0 16:0 18:0 18:1 PI( 18:2 20:4 20:5 22:6 14:0 16:0 18:0 PS( 18:1 18:2 20:4 20:5 22:6 0 1 2 3 4 5 6 7 8 pmol/µg(microsomal(protein(

Figure 3‐4: Acyl chain preference of RAW 264.7 microsomal acyltransferases. Microsomes from RAW 264.7 cells were incubated with lysophospholipid and acyl‐CoA mixtures for 30 min. The 48 possible products of the reaction were quantitated, and data are presented to highlight the acyl chain preference within each of six phospholipid classes. Three independent microsomal preparations were assayed in duplicate, and data shown are average ± SEM.

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14:0% 16:0% 2.0 3

1.5 2 1.0

1 0.5 /µg(microsomal(protein( /µg(microsomal(protein( pmol 0.0 pmol 0 PA PC PE PG PI PS PA PC PE PG PI PS

2.5 18:0%2.25 18:1% 2.00 2.0 1.75 1.50 1.5 1.25 1.0 1.00 0.75 0.50 /µg(microsomal(protein(

0.5 /µg(microsomal(protein( 0.25 pmol 0.0 pmol 0.00 PA PC PE PG PI PS PA PC PE PG PI PS

18:2%3 20:4% 3.5 3.0 2.5 2 2.0 1.5 1 1.0 /µg(microsomal(protein(

/µg(microsomal(protein( 0.5 pmol

pmol 0.0 0 PA PC PE PG PI PS PA PC PE PG PI PS 20:5% 22:6% 1.75 1.00 1.50 0.75 1.25 1.00 0.50 0.75

0.50 0.25 /µg(microsomal(protein( 0.25 /µg(microsomal(protein( pmol 0.00 pmol 0.00 PA PC PE PG PI PS PA PC PE PG PI PS

Figure 3‐5: Polar head group preference of RAW 264.7 microsomal acyltransferases. Microsomes from RAW 264.7 cells were incubated with lysophospholipid and acyl‐CoA mixtures for 10 min. The 48 possible products of the reaction were quantified, and data are presented to highlight the head group preference for each of the acyl‐CoA esters tested. Three independent microsomal preparations were assayed in duplicate, and data shown are average ± SEM.

69

The incorporation of 18:0 chains into PE, PS, PI, and PC was surprisingly high

(Figure 3‐4), providing evidence for additional acyltransferase enzymes, since an acyltransferase with this activity or specificity has not yet been characterized. These phospholipid products could arise from other enzymes not previously described that could exhibit LPEAT, LPIAT, LPSAT, and LPCAT activities. It is also uncertain which enzyme incorporated 22:6 chains into PC and PI. Low activity with 22:6‐CoA esters has been reported for incorporation into LPA by LPAAT3, and LPC by LPCATs 1‐3 (162). The data demonstrated that many acyltransferase activities could be revealed by the dual substrate choice assay, which greatly enhances the chances of identifying previously unknown and novel enzymes. These data confirm that the use of the 17:1‐lysophospholipid in a dual choice setting recapitulates data obtained in the traditional enzymatic assays with natural lysophospholipid species.

An alternative presentation of the data makes it easier to illustrate phospholipid class specificity using each of the acyl chains (Figure 3‐5). Even though this experiment employed a 10‐min incubation, the apparent substrate preference of the combined LPAT activities in the microsomes remained very similar to that presented in Figure 3‐4. The incorporation of 20:4 and 20:5 into phospholipid molecular species had very similar distribution patterns in terms of relative phospholipid class formation, suggesting that the same LPATs (probably MBOAT5/LPCAT3 and MBOAT7/LPIAT1) were responsible for the incorporation of these acyl chains into phospholipids in microsomes from RAW 264.7 cells.

The distribution pattern of newly synthesized phospholipids incorporating 14:0, 16:0,

18:1, 18:2, and 22:6 were also quite similar. The incorporation of 18:0 displayed a unique pattern as compared to the other seven acyl chains, highlighting the possible involvement of an unknown acyltransferase activity.

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Dual Substrate Choice Assay Comparison to Alternative LPAT Assays– The dual substrate choice assay was compared to three other LPAT assays. Specifically, the levels of a single phospholipid molecular species, 17:1/20:4‐PC, were measured (Figure 3‐6). The single substrate assay (Single Substrate) employed the conditions of a traditional assay where only LPC and 20:4‐CoA were added to the microsomes, thus allowing the formation of only one phospholipid species. The lysophospholipid choice assay (Lyso Choice) included 20:4‐CoA and six lysophospholipids, therefore allowing the possible formation of six different phospholipid species. The acyl‐CoA choice assay (CoA Choice) used LPC and the eight different fatty acyl‐CoA esters allowing the potential formation of eight different phosphatidylcholine species. Figure 3‐6 shows an increased 17:1/20:4‐PC production with decreased diversity of substrate options. This finding suggested substrate competition for the active site of the LPATs present in the microsomes. Although the single substrate assay yielded a somewhat more abundant product formation, quantitative evidence for substrate selectivity would require numerous individual assays that would less accurately reflect the true in vivo milieu, since within the intact RAW 264.7 cell the LPATs likely reside in an environment with multiple substrate choices.

Importantly, the substrate preference exhibited for arachidonate incorporation into the different phospholipids was essentially identical using either a series of single substrate assays or the dual substrate choice (Fig. 3‐6). This indicates that the availability of a variety of substrates does not obscure the substrate preference of the enzymes present in the microsomes and supports the usefulness of this assay as a valid alternative to individual assays for each substrate.

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A" 11 10 9 8 7 6 5 4 3

/µg(microsomal(protein( 2 1 pmol 0 DualDual( Choice CoACoA( Choice LysoLyso Choice( SingleSingle( Choice Choice( (Choice( (Choice( Substrate(

B" Single Substrate Assay 11 10 9 8 7 6 5 4 3 /µg(microsomal(protein( 2 1 pmol 0 PC PE PI PS

C" Dual Substrate Choice Assay 3

2

1 /µg(microsomal(protein( pmol 0 PC PE PI PS

Figure 3‐6: Effect of substrate competition on acyltransferase activity. A) Microsomes from RAW 264.7 cells were incubated for 10 min with mixtures of lysophospholipids and CoA esters (Dual Choice), LPC and a mixture of fatty acyl CoA esters (CoA Choice), a mixture of lysophospholipids and 20:4‐CoA (Lyso Choice) or with LPC and 20:4‐CoA (Single Substrate). The production of 17:1/20:4‐PC was quantitated using LC‐MS/MS, and is compared between the four assay conditions. B) LPC, LPE, LPI, and LPS were incubated independently with 20:4‐CoA. The respective phospholipid products were quantitated using LC‐MS/MS. C) A dual choice assay was performed with 6 lysophospholipids and 8 fatty acyl CoAs. All products were quantitated by LC‐MS/MS, and four of the products are displayed. Three independent microsomal preparations were assayed in duplicate, and data shown are average ± SEM.

72

Discussion–

Glycerophospholipids are the major components of all biological membranes and specific molecular species within this family determine properties such as membrane fluidity and curvature (198). Glycerophospholipid molecular species composition also affects the activity of membrane‐associated enzymes and the production of lipid mediators of inflammation (198). The acyl chain composition of phospholipids is the result of a combination of the de novo synthesis route (Kennedy Pathway) and the remodeling of phospholipids through multiple enzymatic activities, including phospholipases, acyl CoA synthetases, and lysophospholipid acyltransferases (Lands Cycle). The combined action of these proteins leads to the large diversity of phospholipid species necessary for different membrane characteristics.

LPATs have been studied since the 1950s by employing assays in which the enzymatic activity was usually determined using substrates consisting of a single lysophospholipid and one fatty acyl‐CoA ester. In the last decade, several LPATs, belonging to two distinct protein families, have been described and characterized (162,184). Detailed enzymatic analysis of these proteins, after cloning and expression of the corresponding genes, has revealed a wide variety of activities for specific lysophospholipid and fatty acyl

CoA substrates (Figure 1‐4). Some activities are catalyzed by the same enzyme, which makes single substrate analysis less useful for characterizing biological samples. Previous work by this and other groups have used different lysophospholipids and/or acyl‐CoA esters to measure LPAT activity (192,224). This method had not been formally validated until the work presented in this chapter to compare the results of the dual choice assay to the results of the corresponding individual single substrate assays. Under the conditions used, the substrate competition did not significantly affect the relative incorporation of

14:0, 18:0, 20:4, or 22:6 into the different classes.

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In this chapter, absolute amounts of the phospholipid products were determined by creating and applying standard curves for each of the six phospholipid classes measured. Because ionization efficiencies can differ greatly among the different phospholipid classes, it was important to use separate internal standards and standard curves for each phospholipid class. An additional discovery from our experimentation was the finding that normalization of the activity data can be achieved by measuring endogenous molecular species within each class, since the absolute quantity of endogenous phospholipid was a direct measure of the quantity of microsomal preparation taken for assay. This study showed excellent correlation of the 16:0/18:1‐ and 18:1/18:1‐ phospholipids with the microsomal protein levels (Figure 3‐3).

The microsomal membranes from RAW 264.7 cells contained a mixture of different enzymes, thus the integrated reacylation activities observed in the dual choice assay reflect this complexity. However, it was possible to identify products that implied the presence of particular LPAT isoforms. In many cases the phospholipids produced in the reaction were not the most abundant phospholipids in the RAW 264.7 cell. An example of this would be

PE and PS that contain DHA. RAW 264.7 cells have appreciable levels of DHA containing PS and PE (230), however during the activity measurements no DHA containing PS or PE were produced. Currently no LPAT activity has been described at this point that can catalyze the formation of these phospholipids. It is possible in other cell types that there is an LPAT that can catalyze these reactions. An alternative explanation of the production of some abundant phospholipids not being formed in this reaction would be that it takes multiple different enzyme subtypes to catalyze these reactions, for instance involving CoA‐ independent transacylases (186). Since we are using microsomal extracts and incubating the reaction for a short period of time, we may be missing the formation of these lipids that would occur in a whole cell system and/or at longer time points.

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The dual choice assay provides a facile and more extensive biochemical approach to studying acyltransferase activity. This novel biochemical technique introduces substrate competition, which provides insight into the preference of substrates, since most biological systems have multiple fatty acyl CoA esters and lysophospholipids present. Perhaps most important is that this substrate choice assay is readily implemented using tandem mass spectrometry.

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CHAPTER IV

MEMBRANE BOUND O‐ACYLTRANSFERASE 5/ LYSOPHOSPHATIDYLCHOLINE

ACYLTRANSFERASE 3 ROLE IN REGULATING LEUKOTRIENE FORMATION THOUGH AA

INCORPORATION INTO PHOSPHOLIPIDS

Introduction–

The sn‐2 position of phospholipids is a readily available storage site of the bioactive precursor AA as an ester. When cells are stimulated, some phospholipases rapidly hydrolyze these phospholipids located in cellular membranes to liberate free arachidonate allowing it to be available for oxidization into eicosanoids. There are many regulatory steps engaged in the process of releasing AA due to the numerous enzymes that mediate the release, oxidation, reacylation, and degradation of AA and eicosanoids. A large body of literature has focused on the regulation of a phospholipase that releases AA (cPLA2α) or the enzymes that synthesize eicosanoids, 5‐LO or COX1/2. However, only AA that is liberated at or transported to the nuclear envelope would be in the correct subcellular location to be oxidized by the 5‐LO/FLAP complex. Work performed in human neutrophils with exogenously added [3H]‐AA showed that very little of the provided AA was converted to eicosanoids. The majority of the exogenously added [3H]‐AA was reacylated into phospholipids and neutral lipids (231). The acylation of labeled AA into phospholipids and neutral lipids suggested the reacylation pathway as a dominant pathway engaged in the production of eicosanoids. It also suggested a critical role for the incorporation and availability of AA in phospholipids as a regulator of eicosanoids production.

MBOAT5 was originally misidentified as a protein acyltransferase when the human genome was published (232). Later, a homologous isoform was identified in yeast (ale1), and when deleted the yeast had decreased lysophospholipid acyltransferase activity (233).

To determine the substrate specificity of MBOAT5, it was subcloned and expressed in a

76

yeast null for Ale1. The deletion of ale1 provided for a relatively clean background in order to determine MBOAT5 specificity as a lysophospholipid acyltransferase (192,233).

Through these experiments it was found that MBOAT5 preferentially incorporated AA into lyso PE, PS, and PC (192). In parallel studies, an enzyme was found that selectively incorporated 18:2‐CoA or 20:4‐CoA esters into lysoPC and was given the name lysophosphatidylcholine acyltransferase 3 (LPCAT3), which was later determined to be the gene mboat5 (162,234). However, the name LPCAT3 is not an accurate name given that it utilizes lyso PS equally as well as lyso PC, and lyso PE to a lesser extent. We will refer to this enzyme as MBOAT5/LPCAT3.

While performing these experiments, our laboratory determined that

MBOAT5/LPCAT3 was expressed in human neutrophils (192). By a series of accidental experiments, it was found that thimerosal, a mercury containing compound that is used as a antifungal and antibacterial preservative, inhibited the enzymatic function of the all

MBOATs/LPCATs (235). When neutrophils were activated by granulocyte macrophage colony‐stimulating factor (GM‐CSF) and then stimulated with N‐formylmethionyl‐leucyl‐ phenylalanine (fMLP), they released a significant amount of AA leading to 0.2 ng of leukotriene B4 (LTB4) per million cells (236). However, when the neutrophils were first treated with thimerosal (25 μM), and later stimulated, there was a dramatic increase in the amount of free AA and an increase of 9 ng of LTB4 to per million cells, a 45 fold increase

(236).

These findings led to the hypothesis that inhibiting the MBOATs/LPCATs would prevent reincorporation of AA‐CoA, which then can be hydrolyzed back to free AA (Figure

4‐1). When elevated concentrations of AA were available to 5‐LO, it could be converted to leukotrienes, thus driving the dramatic increase in leukotriene production. Therefore, the reacylation arm of the leukotriene pathway was thought to be a significant regulator of

77

eicosanoid production. To determine which enzymes(s) are important in the reacylation of

AA after cell stimulation, this chapter will focus on increasing MBOAT5/LPCAT3 expression to determine if that decreases eicosanoid formation as well as reciprocal experiments to decrease MBOAT5/LPCAT3 expression to determine if that increases eicosanoid formation by the availability of AA to 5‐LO.

Phospholipase*C* x DAG* DAG*Lipase*

Phospholipase*A

MAG* Lysophospholipid* - acyltransferase'

MBOAT5/LPCAT3' 2* MAG*Lipase* Sterol* MBOAT7/LPIAT1' *o>acyltransferase*

CE*

Diglyceride** Thimerosal' acyltransferase* Eicosanoids* Cyclooxygenase* TAG* Acyl>coenzyme*A* Lipoxygenase* Hydrolase' Cytochrome*P450s*

β>oxidaGon*

Fa=y*acyl*CoA*

Acyl>coenzyme*A* Synthetase*Long*chain**

(ACSL)'

Figure 4‐1: Thimerosal inhibits MBOAT5/LPCAT3 and MBOAT7/LPIAT1 leads to increased leukotriene production in human neutrophils. A modified scheme from (192) shows how blocking the MBOAT5/LPCAT3 and MBOAT7/LPIAT1 with thimerosal would lead to the increase in leukotriene formation.

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Experimental Procedures–

Materials– Fatty acyl‐CoA esters and phospholipids were from Avanti Polar Lipids

(Alabaster, AL): tetradecanoyl‐coenzyme A (myristoyl; 14:0‐CoA), hexadecanoyl‐coenzyme

A (palmitoyl; 16:0‐CoA), octadecanoyl‐coenzyme A (stearoyl; 18:0‐CoA), (9Z)‐ octadecenoyl‐coenzyme A (oleoyl; 18:1‐CoA), (9Z,12Z)‐octadecadienoyl‐coenzyme A

(linoleoyl; 18:2‐CoA), (5Z,8Z,11Z,14Z)‐eicosatetraenoyl‐coenzyme A (arachidonoyl; 20:4‐

CoA), (5Z,8Z,11Z,14Z,17Z)‐eicosapentaenoyl‐coenzyme A (20:5‐CoA),

(4Z,7Z,10Z,13Z,16Z,19Z)‐docosahexaenoyl‐coenzyme A (22:6‐CoA), 1‐(10Z)‐ heptadecenoyl‐2‐hydroxy‐lysophosphatidic acid (17:1‐LPA), 1‐(10Z)‐heptadecenoyl‐2‐ hydroxy‐lysophosphatidylcholine (17:1‐LPC), 1‐(10Z)‐heptadecenoyl‐2‐hydroxy‐ lysophosphatidylethanolamine (17:1‐LPE), 1‐(10Z)‐heptadecenoyl‐2‐hydroxy‐ lysophosphatidylglycerol (17:1‐LPG), 1‐(10Z)‐heptadecenoyl‐2‐hydroxy‐ lysophosphatidylinositol (17:1‐LPI), 1‐(10Z)‐heptadecenoyl‐2‐hydroxy‐ lysophosphatidylserine (17:1‐LPS), 1‐(9Z)‐octadecenoyl‐2‐hydroxy‐lysophosphatidic acid

(18:1‐LPA), 1‐(9Z)‐octadecenoyl‐2‐hydroxy‐lysophosphatidycholine (18:1‐LPC), 1‐(9Z)‐ octadecenoyl‐2‐hydroxy‐lysophosphatidyserine (18:1‐LPS), (2R)‐[2H31]‐3‐hexadecanoyl‐2‐

(9Z)‐octadecenoyl‐PA ([2H31]‐POPA), (2R)‐[2H31]‐3‐hexadecanoyl‐2‐(9Z)‐octadecenoyl‐PC

([2H31]‐POPC), (2R)‐[2H31]‐3‐hexadecanoyl‐2‐(9Z)‐octadecenoyl‐PE ([2H31]‐POPE), (2R)‐

[2H31]‐3‐hexadecanoyl‐2‐(9Z)‐octadecenoyl‐PG ([2H31]‐POPG), (2R)‐[2H31]‐3‐hexadecanoyl‐

2‐(9Z)‐octadecenoyl‐PI ([2H31]‐POPI), (2R)‐[2H31]‐3‐hexadecanoyl‐2‐(9Z)‐octadecenoyl‐PS

([2H31]‐POPS), 1‐heptadecanoyl‐2‐eicosatetraenoyl‐sn‐glycero‐3‐phosphate (17:0/20:4‐

PA), 1‐heptadecanoyl‐2‐(5Z,8Z,11Z,14Z)‐eicosatetraenoyl‐phosphatidylcholine (17:0/20:4‐

PC), 1‐heptadecanoyl‐2‐(5Z,8Z,11Z,14Z)‐eicosatetraenoyl‐phosphatidylethanolamine

(17:0/20:4‐PE), 1‐heptadecanoyl‐2‐(5Z,8Z,11Z,14Z)‐eicosatetraenoyl‐ phosphatidylglycerol (17:0/20:4‐PG), 1‐heptadecanoyl‐2‐(5Z,8Z,11Z,14Z)‐

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eicosatetraenoyl‐phosphatidylinositol (17:0/20:4‐PI), and 1‐heptadecanoyl‐2‐

(5Z,8Z,11Z,14Z)‐eicosatetraenoyl‐phosphatidylserine (17:0/20:4‐PS). Dulbecco’s modified

Eagle’s medium (DMEM) was purchased from Corning Cellgro (Manassas, VA). RAW 264.7 cells were obtained from ATCC (Manassas, VA). EDTA‐free protease inhibitor cocktail was purchased from Roche (Madison, WI). All other chemicals and solvents were purchased through Fisher Scientific (Pittsburg, PA).

Cell Culture and Microsome Preparation– RAW 264.7 and HEK293T cells were cultured in DMEM (with 4.5 g/L glucose and 100 µM sodium pyruvate) supplemented with

10 % heat‐inactivated FBS. The cells were grown in humidified air with 5 % CO2 at 37 °C.

For microsomal preparations, cells were pelleted at 300 x g, resuspended in homogenization buffer (50 mM Tris‐HCl, pH 7.4, 250 mM sucrose, 1 mM EDTA, 20% (w/v) glycerol, and protease inhibitor cocktail), and lysed using a Sonics Vibra‐Cell probe sonicator (Newtown, CT). Whole cells and cellular debris were pelleted at 12,000 x g for 20 min at 4 °C. The supernatant was transferred to an ultracentrifuge tube and centrifuged at

100,000 x g for 60 min at 4 °C. The microsomal pellet was resuspended in assay buffer (10 mM Tris‐HCl, pH 7.4, 150 mM NaCl, 1 mM EDTA), protein amount was determined using the bicinchoninic acid assay (Thermo Scientific, Rockford, IL), and microsomes were stored at ‐20 °C until used.

Targeted Decrease of MBOAT5/LPCAT3 Gene Expression in RAW 264.7 Cells–

HEK293T cells were transfected using Turbofect (Thermo‐Fisher, Pittsburg, PA) with lentiviral packaging vectors and a vector coding for either a non‐targeting shRNA

(SHC002) or pLKO.1‐puro vector coding for shRNA targeted for the mouse

MBOAT5/LPCAT3 sequence 5'‐CCGGGCCAATCTACTACGATTGTATCTCGAGATAC‐

AATCGTAGTAGATTGGCTTTTTG‐3') (Sigma Aldrich, St. Louis, MO). Medium containing the lentivirus was collected from the HEK293T cell cultures, and 1,5‐dimethyl‐1,5‐

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diazaundecamethylene polymethobromide (Polybrene) was added at a concentration of 8 mg/ml. The receiving RAW 264.7 cells were treated with 8 mg/ml polybrene for 30 min at

37°C. The medium was then replaced with the lentivirus‐containing medium and incubated overnight. The medium was replaced with standard medium containing 2 µg/ml puromycin. Cells were kept under constant selection of 2 µg/ml puromycin for at least three passages. Microsomes where prepared as described previously. Total RNA was extracted from non‐targeted and MBOAT5/LPCAT3 targeted cells using Life Technologies

(Grand Island, NY) Trizol reagent. Total RNA was converted to cDNA with the BioRad iScript reverse transcription supermix. Quantitative polymerase chain reaction (qPCR) was performed using the BioRad iTaq SYBR green supermix (Hercules, CA). Primers for mouse

GAPDH and MBOAT5/LPCAT3 were purchased from Integrative DNA Technologies

(Coralville, IA). Samples were analyzed with a BioRad iQ5 thermal cycler according to the instructions provided with the iTaq SYBR green supermix.

RAW 264.7 Cell Stimulation and Eicosanoids Sample Preparation– Prior to plating the cells, the cells were counted. Cells were plated in the normal growth medium at a density of 1 x106 cells per 6‐well, and allowed to attach overnight in humidified air with 5

% CO2 at 37 °C. Cells treated with pioglitazone were treated for 24 hours before stimulation or RNA measurements. After 16 hours, the cellular medium was removed, the cells were rinsed with 1x PBS, and 1 ml of 1x HBSS with Ca2+ and Mg2+ was placed on the cell. Cells that were treated with thimerosal were preincubated for 5 min before stimulation. Cell stimulated with ATP received a final concentration of 2 mM ATP. Cells were allowed to incubate with the ATP for 15 min. At the conclusion of the experiment the reaction was stopped by addition of 1 ml of 100% methanol that contained a mixture of the eicosanoid internal standards. Samples were stored overnight at ‐20°C to accelerate protein precipitation. Samples were spun at 3000 RPM to pellet the protein precipitants,

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and the supernatant was extracted using Strata‐X polymeric solid‐phase extraction cartridges (Phenomenex, Torrance, CA), per manufactures protocol. Samples were eluted in 100% methanol, dried under vacuum centrifugation, and resuspended in reverse phase starting conditions (67% solvent A: water with 0.05% acetic acid pH 5.7 with ammonium hydroxide; 33% solvent B: 35/65 methanol/acetonitrile).

Liquid Chromatography/Mass Spectrometry for Fatty Acids and Eicosanoids– For reverse phase separation, samples were injected onto a Kinetex C‐18 HPLC column (50 x

2.1 mm, 5 µm; Phenomenex, Torrance, CA) at a flow rate of 0.2 ml/min at 40 % solvent B.

Solvent B was maintained at 40% for 1 min, increased to 75 % over 5 min, and then to 98

% over 1 min. The system was held at 98 % B for 7 min prior to re‐equilibration at 40 % for 5 min. The following list of MRM transitions were monitored: LTC4 m/z 624.5 → 272.2,

LTD4 m/z 495.4 → 177.1, LTE4 m/z 438.3 → 333.3, PGE2 m/z 351.3 → 271.3, PGD2 m/z

351.3 → 233.2, 5‐HETE m/z 319.3 → 115.1, AA m/z 303.2 → 205.2, DHA m/z 327.3 →

283.2, d5‐LTC4 m/z 629.5 → 272.2, d5‐LTD4 m/z 500.4 → 177.1, d5‐LTE4 m/z 443.3 → 338.3, d4‐PGE2 m/z 355.3 → 275.3, d8‐5‐HETE m/z 327.3 → 116.1, and d8‐AA m/z 311.2 → 267.2.

Dual Substrate Choice Assay– Acyltransferase activity was tested in microsomal preparations as described for the single substrate choice except for the following alterations. Stock solutions of reaction materials were made as follows: 6 µM, 20 µM, or 60

µM equimolar mixture of eight acyl‐CoAs (14:0, 16:0, 18:0, 18:1, 18:2, 20:4, 20:5 and 22:6) in methanol, 20 µM or 200 µM equimolar mixture of six lysophospholipids (LPA, LPC, LPE,

LPG, LPI and LPS) in assay buffer. Final lysophospholipid concentrations used were 0.3 µM,

1 µM, 3 µM or 10 µM, as indicated, with final concentrations of 50 ng/µl total protein from microsomes, 3 µM of each fatty acyl‐CoA ester, and 12.5 µM BSA, in a total volume of 200

µl. To assay different microsomal protein amounts, the assay was performed at final concentrations of 3 µM lysophospholipid mix, 3 µM fatty acyl‐CoA ester mix, and 12.5 µM

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BSA, while using microsomal protein concentrations of from 5 ng/µl, 15 ng/µl and 50 ng/µl. To assay different fatty acyl CoA ester concentrations, we used 50 ng/µl total protein from microsomes, 3 µM of each lysophospholipid, and 12.5 µM BSA, while using fatty acyl‐

CoA ester concentrations 0.3 µM, 1 µM, or 3 µM each. The final volume of methanol was kept constant. To assay different time points the components were at final concentrations of 50 ng/µl total protein from microsomes, 3 µM of each lysophospholipid, 3 µM od each fatty acyl‐CoA ester, and 12.5 µM BSA. The assay was performed at 37 °C for 0 min, 1 min, 3 min, 10 min or 30 min. Samples were analyzed by liquid chromatography‐ tandem mass spectrometry (LC‐MS/MS) as described below.

Liquid Chromatography/Mass Spectrometry for Phospholipids– For normal phase separation, samples were injected onto an Ascentis‐Si HPLC column (150 x 2.1 mm, 5 µm;

Supelco, St. Louis, MO) at a flow rate of 0.2 ml/min at 25 % solvent B. Solvent B was maintained at 25 % for 5 min, increased to 60 % over 10 min, and then to 95 % over 5 min.

The system was held at 95 % B for 20 min prior to re‐equilibration at 25 % for 14 min. For reversed phase separation, solvent C was methanol/acetonitrile/water, 60/20/20 (v/v/v), containing 2 mM ammonium acetate, and solvent D was methanol containing 2 mM ammonium acetate. The samples were injected onto an Ascentis‐C18 HPLC column (150 x

2.1 mm, 5 µm; Supelco) at a flow rate of 0.2 ml/min at 75 % solvent D. Solvent D was maintained at 75 % for 1 min and increased to 98 % over 5 min. The system was held at 98

% D for 20 min prior to re‐equilibration at 75 % for 10 min. Phospholipid products of the

LAT assay were measured using an API3200 triple quadrupole mass spectrometer (AB

Sciex, Thornhill, CA) in negative ion mode using multiple‐reaction‐monitoring (MRM) of the m/z transitions shown in Supplemental Table 1. Quantitation was performed using AB

Sciex MultiQuant software and using 17:0/20:4‐PA, 17:0/20:4‐PC, 17:0/20:4‐PE,

17:0/20:4‐PG, 17:0/20:4‐PI and 17:0/20:4‐PS reference standards for dilution curve

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analysis, as described previously (227). Quantitative analysis of the reference standards was performed in the absence or presence of microsomes, and quantitative values did not differ significantly between the two conditions (not shown).

Results–

To confirm the previously published data from the human neutrophils, initial experiments mirrored those previously performed with thimerosal treated neutrophils

(236). The human blood neutrophils preparations were activated by GM‐CSF and then stimulated with fMLP. The non‐thimerosal treated cells release a significant amount of AA leading to the production of leukotriene B4 (LTB4) (Figure 4‐2A). However, when the neutrophils were first treated with thimerosal (25 μM), and later stimulated, there was a dramatic increase in the amount of free AA and an increase of LTB4 (Figure 4‐2A). These human neutrophil preparations also led to the production of leukotriene C4 (LTC4). A substantial amount of LTC4 can be measured due to platelet contamination of neutrophil preparation from blood. Through a transcellular mechanism, the stimulation led to the production of LTC4. The model was switched to a murine monocyte cell line, RAW 264.7.

Neutrophils are very short‐lived cells that perform very little protein synthesis from the gene level. Therefore, to selectively decrease the gene by targeting the mRNA levels would have not been possible. The RAW 264.7 cell is relatively easy to genetically manipulate and is known to produce leukotrienes.

Thimerosal Does Not Have the Same Affect on the RAW 264.7 Cell– From previously published data and the data shown in Figure 4‐2A, thimerosal has a profound effect on leukotriene formation in neutrophils (236). To determine if a similar effect could be seen in the RAW 264.7 cells, experiments were carried out with different concentrations of thimerosal and then cells were stimulated with ATP to activate purinergic receptors on

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the surface of the RAW 264.7 cell. When the eicosanoid profile was measured using tandem mass spectrometry, no change in the amount of free AA was observed with any of the concentrations of thimerosal that were tested (Figure 4‐2B). In contrast to the observations made in the neutrophils, increasing concentrations of thimerosal appeared to decrease the leukotriene C4 (LTC4) production in the RAW 264.7 cell. This decrease in LTC4 production could have been due to the mercury in thimerosal binding to the free sulfur group in glutathione. Glutathione is conjugated to LTA4 to produce LTC4, and therefore thimerosal binding glutathione would lead to decreased availability of glutathione for LTC4 synthase.

A LTB Production Arachidonic Acid LTC4 Production 4 2.0 10 125

100 1.5 8

6 75 1.0 4 50 0.5 2 25 Analyte/Internal Std. Area Analyte/Internal Std. Area 0.0 Analyte/Internal Std. Area 0 0 Not Stimulated No Thimerosal Thimerosal Not Stimulated No Thimerosal Thimerosal Not Stimulated No Thimerosal Thimerosal

Stimulated Stimulated Stimulated B# C# Free$AA$ LTC4$Produc6on$ 0.4 12.5

10.0 0.3

7.5 0.2

Area$Ra6o$ 5.0 Area$Ra6o$ 0.1 2.5

0.0 $ $ $ $ $ 0.0 $ $ $ $ $ Thim Thim Thim Thim Thim Thim Thim Thim Thim Thim Control$ Control$ 1$µM$ 3$µM$ 10$µM$ 30$µM$ 3$µM$ 100$µM$ 1$µM$ 10$µM$ 30$µM$ 100$µM$

Figure 4‐2: Thimerosal (Thim) has a dramatic affect on leukotriene production in human neutrophils but does not have an effect on leukotriene formation in RAW 264.7. A) Human neutrophil preparation with no treatment, stimulation with GM‐CSF and fMLP, or stimulation with GM‐CSF and fMLP in the presence of thimerosal. B & C) Vehicle (control) or different concentrations of thimerosal were preincubated with RAW264.7 cells that were then stimulated with ATP. Y‐axis is the area ratio of the analyte produced over the internal standard area (Area Ratio). n=3, Average ± SEM.

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Pioglitazone Treated RAW 264.7 Cells Upregulates MBOAT5/LPCAT3– Since the approach with thimerosal did not have an affect in the RAW 264.7 cell, the focus of the projected change to include experiments to increase the expression of MBOAT5/LPCAT3.

Previously published work suggested that retinoid X receptor (RXR) occupied the promoter of MBOAT5/LPCAT3, and through heterodimeric binding of liver X receptors

(LXRs) or peroxisome proliferator‐activated receptors (PPARs) MBOAT5/LPCAT3 could be upregulated (237). Therefore, we used RXR transcriptional binding partner agonists to induce an expression of MBOAT5/LPCAT3. LXR agonists (T0901317 and GW3965) had no effect on increasing the expression of MBOAT5/LPCAT3 in the RAW 264.7 cell because these cells express minimal amounts of LXRα (Tontonoz P, unpublished). However, activation of PPARϒ with pioglitazone (133) significantly increased expression of

MBOAT5/LPCAT3 within 24 hours of treatment (Figure 4‐3). There was no change in the expression of MBOAT7/LPIAT1 supporting that this regulation is specific for LPCAT3.

When cells treated with pioglitazone for 24 hours were then stimulated with ATP, there was a dose dependent decrease in the production of leukotrienes and an overall decrease in the amount of free AA at most concentrations of pioglitazone (Figure 4‐3). The data in Figure 4‐3 supported the hypothesis that increased expression of

MBOAT5/LPCAT3 would lead to decreased levels of free AA after cell stimulation and ultimately decrease LTC4 production. Pioglitazone is not specific in its target gene expression, therefore a more targeted approach was used by implementing shRNA that specifically target and lead to the degradation of the MBOAT5/LPCAT3 mRNA.

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MBOAT5/LPCAT3'Expression' MBOAT7/LPIAT1'Expression' 2 8

6

1 4 Fold'Change' Fold'Change' 2

0 0 DMSO' Pio' DMSO' Pio'

0.25 Free'AA' 10.0 LTC4'Produc4on'

0.20 7.5 0.15 5.0 0.10 Area'Ra4o' Area'Ra4o' 2.5 0.05

0.00 0.0 ' ' ' ' ' ' ' ' ' ' Pio Pio Pio Pio Pio Pio Pio Pio Pio Pio Control' Control' 1'µM' 3'µM' 1'µM' 3'µM' 10'µM' 0.1'µM' 0.3'µM' 10'µM' 0.1'µM' 0.3'µM'

Figure 4‐3: Pioglitazone, Pio, treatment for 24 hr leads to increased MBOAT5/LPCAT3 expression and decreases free AA and LTC4 levels. Vehicle (control) or different concentrations of Pio were preincubated with RAW264.7 cells that were then stimulated with ATP. Y‐axis is the area ratio of the analyte produced over the internal standard area (Area Ratio). n=3, Average ± SEM.

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Activity of MBOAT5/LPCAT3 was Significantly and Specifically Decreased in RAW

264.7 Cell Using a Targeted shRNA– To determine if MBOAT5/LPCAT3 expression and activity was specifically decreased, the LPAT activity assay described in Chapter 3 was used. RAW 264.7 cells containing non‐targeted shRNA cells and shRNA against

MBOAT5/LPCAT3 were used to make microsomes. The microsomal preparations are known to contain many membrane‐associated proteins, including a mixture of lysophospholipid acyltransferases. These microsomal preparations were incubated with six non‐endogenous lysophospholipids (17:1‐LPA, ‐LPC, ‐LPE, ‐LPG, LPI, and –LPS) and eight fatty acyl CoA esters (14:0‐, 16:0‐, 18:0‐, 18:1‐, 18:2‐, 20:4‐, 20:5‐, 22:6‐CoA), and the

48 possible unique phospholipid products were detected using liquid chromatography tandem mass spectrometry (LC‐MS/MS).

These microsomes underwent identical treatments to be able to directly compare the activity data. Figure 4‐4 shows the raw data of the incorporation of AA into the six lysophospholipids that were provided from the two microsomal preparations. To analyze all of the phospholipids precursor products in a single run, negative ion mass spectrometry was used. The traces in Figure 4‐4 show the raw negative ion counts from specific ion transitions detected by the electron multiplier in the mass spectrometer, with no quantitative corrections applied. From the raw data, synthesis of each class of lipid cannot be compared directly because ionization efficiency differs between phospholipid classes.

However, visual comparison between the trace of the non‐targeted shRNA microsomes and the MBOAT5/LPCAT3 shRNA targeted microsomes suggests decreased incorporation of AA into PC, PS, and PE with very little to no change in AA incorporation into PG, PI, or PA

(Figure 4‐4A, control compared to 4‐4B, shRNA targeted MBOAT5/LPCAT3 microsomes).

This decrease in AA incorporation into each of the phospholipid that MBOAT5/LPCAT3 utilized corresponded well to the decrease in the mRNA expression (Figure 4‐4, inset).

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9500 A" PE# PS# Intensity, cps

PI# PA# PC# Intensity,#cps#

PG#

2 6 10 14 18 22 26 30 34 38 Time, min 9500 LPCAT3 mRNA Levels 1.5 B" 1.0

0.5

PI# Fold#Change# Intensity, cps 0.0 PS# CNTL# KD# PA#

Intensity,#cps# PG# PE# PC#

2 6 10 14 18 22 26 30 34 38 Time, min

Figure 4‐4: Phospholipids containing AA are reduced in microsomal extracts from cells containing an shRNA targeted against MBOAT5/LPCAT3. RAW 264.7 cells were transduced with either a non‐targeting shRNA construct (A; CNTL) or an shRNA targeted against the MBOAT5/LPCAT3 gene (B; KD). The levels of mRNA expression of MBOAT5/LPCAT3, as determined by quantitative PCR, are shown in the inset. The microsomes were incubated with mixtures of 17:1‐lysophospholipids (6 species) and fatty acyl‐CoA esters (8 species). The negative ion LC‐MS/MS profiles of the six phospholipids containing the 20:4 acyl chain are shown. The data shown are representative of a total of nine experiments performed with three independent microsomal preparations. The inset data are average ± SEM.

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The raw data was then converted to total quantity of phospholipid newly synthesized and normalized to microsomal protein content amount using the ratios of the precursor‐product ion transition to the corresponding internal standards and applying calibration curves. This transformation enabled the direct comparison of newly synthesized phospholipid products in each phospholipid class. Microsomes from RAW

264.7 cells were found to incorporate arachidonate into all six classes of phospholipids tested in this assay, albeit to very different extents. In the MBOAT5/LPCAT3‐targeted microsomes, there was a substantial decrease in the incorporation of 20:4, specifically into

PC, PE, and PS, but no decrease in 17:1/20:4‐PA, ‐PG, or ‐PI production was observed

(Figure 4‐5A). These changes were clearly indicative of a decrease in MBOAT5/LPCAT3 activity, while other LPAT activities for AA were unchanged in this microsomal preparation.

The incorporation of 16:0 (Figure 4‐5B) acyl chains into the six different phospholipids was found to be unchanged between the control and MBOAT5/LPCAT3 shRNA microsomes. This finding was consistent with this 16:0‐CoA being a poor substrate for MBOAT5/LPCAT3, and other enzymes being responsible for this lysophospholipid activity on the microsomes. Changes in MBOAT5/LPCAT3 activity were also revealed by the incorporation of 20:5‐CoA and 18:2‐CoA (Figure 4‐5C&D). It had been previously shown that MBOAT5/LPCAT3 could incorporate 18:2 into lyso‐PC, PE and PS, therefore the finding that these products were decreased was an expected finding (Figure 4‐5D). The incorporation of 20:5‐CoA was not previously tested but showed similar effects to the incorporation of AA into phospholipids (Figure 4‐5A&D). The decrease in specific phospholipid product formation supported that the shRNA selectively decreased the activity of MBOAT5/LPCAT3 while the other LPAT activities remained the same. These

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data also supported MBOAT5/LPCAT3 as the major LPAT to incorporate AA into lyso‐PC,

PE and PS in the RAW 264.7 cell.

A" 20:4(( B" 16:0( NS( * * NS NS NS NS NS NS NS NS 7 * 4

6 3 5

4 2 3

2 /µg(microsomal(protein( 1 /µg(microsomal(protein( 1 pmol pmol 0 0 CNTLKD CNTLKD CNTLKD CNTLKD CNTLKD CNTLKD CNTLKD CNTLKD CNTLKD CNTLKD CNTLKD CNTLKD PA PC PE PG PI PS PA PC PE PG PI PS

C" 20:5(( D 18:0(

NS * * NS NS NS * * NS NS * 4 * 5

4 3

3 2 2 /µg(microsomal(protein(

/µg(microsomal(protein( 1 1 pmol pmol 0 0 CNTLKD CNTLKD CNTLKD CNTLKD CNTLKD CNTLKD CNTLKD CNTLKD CNTLKD CNTLKD CNTLKD CNTLKD PA PC PE PG PI PS PA PC PE PG PI PS

Figure 4‐5: Effect of reduced expression of MBOAT5/LPCAT3 on acyltransferase activity. RAW 264.7 cells were transduced with either a non‐targeting shRNA construct (CNTL) or an shRNA targeted against the MBOAT5/LPCAT3 gene (KD), and microsomes were used in the dual substrate choice acyltransferase assay. Newly synthesized phospholipid molecular species containing (A) 20:4 acyl chains, (B) 16:0 acyl chains, (C) 20:5 acyl chains, or (D) 18:2 acyl chains. Four independent microsomal preparations were assayed for a total of nine experiments, and data shown are average ± SEM, two‐tailed Students t‐test, *=p<0.05.

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To determine if there were any other changes in LPAT activity, the non‐targeting shRNA control and MBOAT5/LPCAT3 shRNA derived microsomes were compared to one another. In this figure, the dual‐choice assay results are presented as the quantity of newly synthesized phospholipid molecular species catalyzed by the LPCATs knockdown microsomes after subtraction of the quantity made by the non‐targeting shRNA RAW 264.7 cell microsomes when the dual‐choice assay was performed. The largest drop in molecular species occurred in the acylation of LPC and LPS with 18:2‐, 20:4‐, 20:5‐CoA chains, which would be expected for a loss of MBOAT5/LPCAT3 activity (Figure 4‐6). To a lesser extent this occurred with acylation of LPE using these same polyunsaturated CoA esters. There were a few minor changes in other products, however none of the products pointed at a single LPAT that would have been changed with the depletion of MBOAT5/LPCAT3 and are probably due to variability in the assay.

The Depletion of MBOAT5/LPCAT3 Does Not Lead to Changes in Leukotriene

Production– Three independent constructs that all led to similar depletion of

MBOAT5/LPCAT3 mRNA were used to determine if there were changes in leukotriene formation. Cells were stimulated with 2mM ATP, the eicosanoids were extracted, and analysis was performed by tandem mass spectrometry. The raw data traces for the leukotriene production are shown in Figure 4‐7. The blue traces are that of the leukotriene

C4 (LTC4) produced by the cells, and the red traces are the internal standard that was added to each sample at equal concentrations. Leukotriene formation was unchanged in two of the three shRNA constructs against MBOAT5/LPCAT3 albeit similar depletion of mRNA. This finding was consistent among multiple experiments showing that depletion of

MBOAT5/LPCAT3 in RAW 264.7 cells did not have an effect on leukotriene biosynthesis.

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14:0 16:0 18:0 18:1 PA" 18:2 20:4 20:5 22:6 14:0 16:0 18:0 18:1 PC" 18:2 20:4 20:5 22:6 14:0 16:0 18:0 18:1 PE" 18:2 20:4 20:5 22:6 14:0 16:0 18:0 18:1 PG" 18:2 20:4 20:5 22:6 14:0 16:0 18:0 18:1 PI" 18:2 20:4 20:5 22:6 14:0 16:0 18:0 18:1 PS" 18:2 20:4 20:5 22:6 -7 -6 -5 -4 -3 -2 -1 0 1 2 3 4 5 6 7 Δ"pmol/µg"microsomal"protein""

Figure 4‐6: Phospholipid molecular species production during reduced expression of MBOAT5/LPCAT3 in microsomes. RAW 264.7 cells were transduced with either a non‐ targeting shRNA construct (CNTL) or an shRNA targeted against the MBOAT5/LPCAT3 gene (KD), and microsomes were used in the dual substrate choice acyltransferase assay. Four independent microsomal preparations were assayed. All 48 phospholipids products were quantitated by tandem mass spectrometry and normalized by microsomal protein content. Each phospholipid molecular species produced by KD cells was subtracted from that measured in CNTL cells. The results are expressed as a histogram for each phospholipid molecular species as a less abundant product (bars going left of center) or more abundant product (bars going right of center).

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Non1targe7ng' LPCAT3' LPCAT3' LPCAT3' shRNA' shRNA'#1' shRNA'#2' shRNA'#3'

4.0e4 4.0e4 4.0e4 4.0e4

3.0e4

LTC4' LTC4' LTC4' LTC4' 2.0e4 Intensity, cps Intensity,

1.0e4 d51 d51 d51 d51 LTC4' LTC4' LTC4' LTC4'

4.0 5.0 6.0 7.0 4.0 5.0 6.0 7.0 4.0 5.0 6.0 7.0 4.0 5.0 6.0 7.0 Time, min Time, min Time, min Time, min

Figure 4‐7: Targeted decrease of MBOAT5/LPCAT3 in RAW 264.7 cells does not change leukotriene production. Non‐targeted shRNA and MBOAT5/LPCAT3 shRNA containing RAW 264.7 cells were stimulated with ATP and leukotriene formation was measured by LC‐MS/MS. LTC4 produced by the cells (blue traces) were compared to the internal standard d5‐LTC4 (red traces).

Discussion–

MBOAT5/LPCAT3 has been shown to be a lysophospholipid acyltransferase critical for the incorporation of AA into PC, PE, and PS. In C. elgans MBOAT5/LPCAT3 is critical in normal larval development and in mice MBOAT5/LPCAT3‐/‐ is lethal at birth due to hypoglycemia ((238) and P. Tontonoz unpublished). It has been previously shown that

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thimerosal inhibits the activity of MBOAT5/LPCAT3 (192). When neutrophils were acutely treated with thimerosal, before the cells were stimulated, there was a dramatic increase in the production of leukotrienes compared to cells not treated with thimerosal (236).

Although the RAW 264.7 cell has both expression and activity for MBOAT5/LPCAT3, thimerosal did not have an effect on the leukotriene production in stimulated cells. One caveat was that thimerosal could act on the human isoform of MBOAT5/LPCAT3 and not the mouse isoform. Between the mouse and human isoforms there is 87% identity and

93% similar amino acid match. Since the protein is highly conserved from mouse to human, it is unlikely that thimerosal cannot elicit an effect on MBOAT5/LPCAT3, although it has not been tested.

Previous groups had showed that RXR transcriptional binding partners regulate

MBOAT5/LPCAT3 at the gene level (237). Treatment of the parental RAW 264.7 with the

PPARΥ agonist (pioglitazone) elicited a five‐fold increase in the expression of

MBOAT5/LPCAT3 at the mRNA level, but there was no change measured in the expression of MBOAT7/LPIAT1. After 24 hours of treatment with pioglitazone, the cells were stimulated with ATP. There was a dose dependent decrease in the production of leukotrienes compared to the vehicle treated cells. The decrease in leukotrienes supported the idea that MBOAT5/LPCAT3 is critical in the regulation of free AA, and that increasing the expression of the gene would lead to faster reacylation of AA into phospholipids and decrease leukotriene production.

To determine if the opposite was true, that decreasing MBOAT5/LPCAT3 would lead to a decrease in leukotriene production, shRNAs were transduced into the RAW 264.7 cells to selectively decrease MBOAT5/LPCAT3. Initially the most robust shRNA construct was used in the characterization of the cells. To confirm the selective decrease in

MBOAT5/LPCAT3, mRNA measurements were made comparing the targeted shRNA

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against the non‐targeted shRNA cells. The cells displayed a 60% decrease in mRNA expression compared to the cells with the non‐targeted shRNA construct. The 60% decrease in mRNA message led to a 2 to 8 fold decrease in the activity of MBOAT5/LPCAT3 in microsomal activity assays. The activity assay and qPCR data supported a significant decrease in the MBOAT5/LPCAT3 compared to the non‐target shRNA control cells.

Once the decrease in MBOAT5/LPCAT3 expression was confirmed for the three shRNA constructs (data not shown), the cells were stimulated with ATP to determine the effect of MBOAT5/LPCAT3 deficiency in leukotriene formation. Non‐targeted shRNA and

MBOAT5/LPCAT3 targeted shRNA cells were stimulated for 30 min with ATP and the eicosanoids were measured by LC‐MS/MS. The raw data from the mass spectrometer revealed that two of the three shRNA constructs had no effect on leukotriene formation compared to the non‐targeted control cells, and one of the constructs appeared to slightly decrease leukotriene formation.

A possible explanation for why the targeted decrease of MBOAT5/LPCAT3 in the

RAW 264.7 cell did not have an effect was because the AA content in phospholipid content could not be lowered. Cultured cells generally have a decreased level of AA esterified within phospholipids compared to the levels in in vivo isolates. Human neutrophils have a plethora of AA available to be incorporated into phospholipids taken in from the diet and circulating in plasma at a typical concentration of 3 μM (239). Cultured cells only get the fatty acids provided in their medium that contains 10% fetal bovine serum or from fatty acid synthesis. The cells are undergoing rapid division in culture, and their membranes are expanding rapidly which requires fatty acids for de novo phospholipid synthesis. Cell culture serum is a finite resource, therefore the cells have to synthesize their own fatty acids through acetic acid and malonyl‐CoA. Through the de novo route, mammalian cells can only produce ω‐9 unsaturated fatty acids. AA is an ω‐6 fatty acid that is an essential

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fatty acid, meaning that it has to be obtained through diet (31). When ω‐6 fatty acids are depleted, as in the case for cell culture cells, they begin to compensate by elongation and desaturation of the 18:1, ω‐9 fatty acid to form the 20:3, ω‐9 fatty acid, Mead acid (240).

Although the presence of Mead acid has not been confirmed by identification of the double bond positions and cis‐trans stereochemistry, the fragment ion m/z 305 was observed from multiple phospholipid species that normally contain AA. The presence of m/z 305 fits as the carboxylate anion of Mead acid.

Cells in culture have very little AA containing phospholipids, therefore depleting an enzyme that makes a product that is already depleted may not yield as significant of an affect as a cell that has more AA available to incorporate into phospholipids. Additional experiments addressing the depletion of AA in these cells will be shown and supported in later chapters of this thesis, but addition of [3H]‐AA to RAW 264.7 cells directly labeled phospholipids where as in the neutrophil 55% of the [3H]‐AA was sequestered into the neutral lipid pool (231).

Enhanced expression of MBOAT5/LPCAT3 supported our initial hypothesis that cells treated with pioglitazone, which increased the expression of MBOAT5/LPCAT3, would lead to a decrease in LTC4 and AA. However, decreasing expression of MBOAT5/LPCAT3 had no effect on leukotriene formation. It is possible that decreases or inhibition of

MBOAT5/LPCAT3, especially chronically with shRNA, is not tolerated well in the RAW

264.7 cells. The cells may be at a minimum concentration of AA contain phospholipids, and decreasing the pool further is not feasible to maintain normal function. However, increases in expression of MBOAT5/LPCAT3 would allow for more robust reincorporation of AA into phospholipids. The free AA‐CoA formed after cell stimulation would be reacylated rapidly and not available for hydrolysis back to free AA. Therefore, overexpression of

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MBOAT5/LPCAT3 would support the working hypothesis that MBOAT5/LPCAT3 is critical in the regulation of AA into phospholipids and production of eicosanoids.

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CHAPTER V

THE EFFECT OF MEMEBRANE BOUND O‐ACYLTRANSFERASE

7/LYSOPHOSPHATIDYLINOSITOL ACYLTRANSFERASE 1 AND ACYL‐COENZYME A

SYNTHETASE LONG CHAIN 4 ON ARACHIDONIC ACID AND LEUKOTRIENE

PRODUCTION

Introduction–

The phospholipid source of AA, for the production of eicosanoids, has been a controversial topic. There are four major phospholipid classes that contain AA at the sn‐2 position: phosphatidylcholine (PC), phosphatidylethanolamine (PE), phosphatidylinositol

(PI), and phosphatidylserine [PS]. There is evidence to support the liberation of AA from

PC/PE and PI, however it has been difficult to determine the true phospholipid source because of numerous factors. The first issue is a small percentage of the AA containing phospholipid pool is utilized in the liberation of AA, which makes identification of changes in the total phospholipid pool difficult. The second difficulty has been with unidentified enzymes, transacylases, which transfer an acyl chain from a phospholipid to a lysophospholipid. Therefore, a lysophospholipid that is measured may not be one from which AA was removed from because of the activity of transacylases. This being said, a body of literature has supported PI as the source of AA for the production of eicosanoids

(241‐243). AA containing PI is also thought to be a critical lipid for the docking of cPLA2α for the liberation of AA (242,244,245).

Suggested mechanisms involved in the regulation of eicosanoids have largely focused on the enzymes that catalyze the formation of each of the products and corresponding receptors in which the eicosanoids bind. Previously published data suggested that the majority of the released AA is not made into eicosanoids, but rather it is sequestered by a series of enzymes to reacylate the AA into neutral lipids and

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phospholipids (231). This pathway involves the initial formation of an arachidonoyl‐ coenzyme A (AA‐CoA) that is formed by the action of acyl‐CoA synthetase long‐chains

(ACSLs). There are five ACSL isoforms that have been shown to utilize AA as a substrate:

ACSL1, 2, 4, 5, 6 (161). However, ACSL4 has been suggested to be critical in the regulation of AA in monocytes (246).

Once this thioester forms, the AA‐CoA can be used as a substrate by a wide variety of acyltransferase. There are two major families that reacylate lysophospholipids: 1‐ acylglycerol‐3‐phosphate O‐acyltransferases/lysophosphatidic acid acyltransferases

(AGPATs/LPAATs) and membrane bound O‐acyltransferases (MBOATs). In 2008, two separate groups, including our laboratory, discovered an acyltransferase that preferentially esterified AA‐CoA to PI namely MBOAT7 (191,192). In C. elegans the gene that encoded this protein was annotated mboa‐7, and in humans mboat7. After its discovery, the protein was renamed lysophosphatidylinositol acyltransferase 1 (LPIAT1) for the protein’s described preference for AA‐CoA and lysophosphatidylinositol (162). This enzyme will be referred to as MBOAT7/LPIAT1.

Since the discovery of MBOAT7/LPIAT1, it has been selectively deleted from C. elegans and mice. Deletion of mboa‐7 in C. elegans led to decreased maturation of the worms. Deletion of mboat7 in mice led to decreased birth rate of the null animals, and by day 30 there were almost no surviving animals (247). The major phenotypes that were observed were decreased body size and improper hippocampal development (247). All of these selective deletions of MBOAT7/LPIAT1 led to decreases in phosphatidylinositol lipids, and specifically, led to decreases in 4,5‐phosphatidylinositol bisphosphate

(PtdIns(4,5)‐P2) (247,248).

The work presented in the chapter describes the selected decrease of

MBOAT7/LPIAT1 and ACSL4 in the RAW 264.7 cell (a murine monocyte cell line) to

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determine the affect these enzymes have on the regulation of leukotriene biosynthesis. To perform these experiments, short‐hairpin RNAs (shRNAs) were used to selectively decrease the expression of MBOAT7/LPIAT1 and ACSL4. Once the targeted decrease in

MBOAT7/LPIAT1 and ACSL4 was established, cells were challenged to release AA and produce eicosanoids. Interestingly, decreases in ACSL4 led to increased levels of AA, but had no affect on eicosanoid formation. Whereas, decreases in MBOAT7/LPIAT1 led to decreased levels of free AA and a decrease in leukotriene formation that could not be rescued by addition of exogenous AA. All the data presented in this chapter supports the hypothesis that MBOAT7/LPIAT1 is a critical regulator of AA and LTC4 biosynthesis in these cells, but the mechanism of how this occurs is still yet to be discovered.

Experimental Procedure–

Materials– Fatty acyl‐CoA esters and phospholipids were from Avanti Polar Lipids

(Alabaster, AL): tetradecanoyl coenzyme A (myristoyl; 14:0‐CoA), hexadecanoyl coenzyme

A (palmitoyl; 16:0‐CoA), octadecanoyl coenzyme A (stearoyl; 18:0‐CoA), (9Z)‐octadecenoyl coenzyme A (oleoyl; 18:1‐CoA), (9Z,12Z)‐octadecadienoyl coenzyme A (linoleoyl; 18:2‐

CoA), (5Z,8Z,11Z,14Z)‐eicosatetraenoyl coenzyme A (arachidonoyl; 20:4‐CoA),

(5Z,8Z,11Z,14Z,17Z)‐eicosapentaenoyl coenzyme A (20:5‐CoA), (4Z,7Z,10Z,13Z,16Z,19Z)‐ docosahexaenoyl coenzyme A (22:6‐CoA), 1‐(10Z)‐heptadecenoyl‐2‐hydroxy‐ lysophosphatidic acid (17:1‐LPA), 1‐(10Z)‐heptadecenoyl‐2‐hydroxy‐lysophosphatidyl‐ choline (17:1‐LPC), 1‐(10Z)‐heptadecenoyl‐2‐hydroxy‐lysophosphatidylethanolamine

(17:1‐LPE), 1‐(10Z)‐heptadecenoyl‐2‐hydroxy‐lysophosphatidylglycerol (17:1‐LPG), 1‐

(10Z)‐heptadecenoyl‐2‐hydroxy‐lysophosphatidylinositol (17:1‐LPI), 1‐(10Z)‐ heptadecenoyl‐2‐hydroxy‐lysophosphatidylserine (17:1‐LPS), (2R)‐[2H31]‐3‐hexadecanoyl‐

2‐(9Z)‐octadecenoyl‐PA ([2H31]‐POPA), (2R)‐[2H31]‐3‐hexadecanoyl‐2‐(9Z)‐octadecenoyl‐

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PC ([2H31]‐POPC), (2R)‐[2H31]‐3‐hexadecanoyl‐2‐(9Z)‐octadecenoyl‐PE ([2H31]‐POPE), (2R)‐

[2H31]‐3‐hexadecanoyl‐2‐(9Z)‐octadecenoyl‐PG ([2H31]‐POPG), (2R)‐[2H31]‐3‐hexadecanoyl‐

2‐(9Z)‐octadecenoyl‐PI ([2H31]‐POPI), (2R)‐[2H31]‐3‐hexadecanoyl‐2‐(9Z)‐octadecenoyl‐PS

([2H31]‐POPS), 1,2‐didodecanoyl‐sn‐glycero‐3‐phosphate (12:0/12:0‐PA), 1,2‐ didodecanoyl‐phosphatidylcholine (12:0/12:0‐PC), 1,2‐didodecanoyl‐ phosphatidylethanolamine (12:0/12:0‐PE), 1,2‐didodecanoyl‐phosphatidylglycerol

(12:0/12:0‐PG), (8:0/8:0‐PI), 1,2‐ditetradecanoyl‐phosphatidylserine (14:0/14:0‐PS), and

(17:0/20:4‐PI). Dulbecco’s modified Eagle’s medium (DMEM) was purchased from Corning

Cellgro (Manassas, VA). RAW 264.7 cells were obtained from ATCC (Manassas, VA). EDTA‐ free protease inhibitor cocktail was purchased from Roche (Madison, WI). All other chemicals and solvents were purchased through Thermo‐Fisher Scientific (Pittsburg, PA).

Cell culture and knockdown of MBOAT7/LPIAT1 gene expression in RAW 264.7 cells‐ RAW 264.7 and HEK293T cells were cultured in DMEM (with 4.5 g/L glucose and

100 µM sodium pyruvate) supplemented with 10 % heat‐inactivated FBS. The cells were grown in humidified air with 5 % CO2 at 37 °C. HEK293T cells were transfected using

Turbofect (Thermo‐fisher) with lentiviral packaging vectors and a vector coding for either a non‐targeting shRNA (SHC002) (Sigma‐Aldrich, St. Louis, MO), pLKO.1‐puro vector coding for shRNA targeted for the mouse MBOAT7/LPIAT1 Construct #1 5’‐

CCGGCTGCTAACATCAGGGTATTACCTCGAGGTAATACCCTGATGTTAGCAGTTTTTTG‐3’

Construct #2 sequence 5' CCGGCTGGTTACTACCTAAGCTTCACTCGAGTGAAGCTTA

GGTAGTAACCAGTTTTTTG‐3'). Medium containing the lentivirus was collected from the

HEK293T cell cultures, and 1,5‐dimethyl‐1,5‐diazaundecamethylene polymethobromide

(Polybrene) was added at a concentration of 8 mg/ml. The receiving RAW 264.7 cells were treated with 8 mg/ml polybrene for 30 min at 37°C. The medium was then replaced with the lentivirus‐containing medium and incubated overnight. The medium was replaced with

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standard medium containing 2 µg/ml puromycin. Cells were kept under constant selection of 2 µg/ml puromycin for at least three passages.

Quantitative Reverse Transcriptase PCR in RAW 264.7 Cells– Total RNA was extracted from non‐targeted and MBOAT5/LPCAT3 targeted cells using Life Technologies

(Grand Island, NY) Trizol reagent. Total RNA was converted to cDNA with the BioRad iScript reverse transcription supermix (BioRad, Hercules, CA). Quantitative polymerase chain reaction (qPCR) was performed using the BioRad iTaq SYBR green supermix.

Primers for mouse GAPDH, MBOAT7/LPIAT1, MBOAT5/LPCAT3, ACSL1, ACSL4, 5‐LO, and cPLA2α were purchased from Integrative DNA Technologies. Samples were analyzed with a

BioRad iQ5 thermal cycler according to the instructions provided with the iTaq SYBR green supermix.

Dual Substrate Choice Assay and Microsome Preparation– For microsomal preparations, cells were pelleted at 300 x g, resuspended in homogenization buffer (50 mM

Tris‐HCl, pH 7.4, 250 mM sucrose, 1 mM EDTA, 20% (w/v) glycerol, and protease inhibitor cocktail), and lysed using a Sonics Vibra‐Cell probe sonicator (Newtown, CT). Whole cells and cellular debris were pelleted at 12,000 x g for 20 min at 4 °C. The supernatant was transferred to an ultracentrifuge tube and centrifuged at 100,000 x g for 60 min at 4 °C.

The microsomal pellet was resuspended in assay buffer (10 mM Tris‐HCl, pH 7.4, 150 mM

NaCl, 1 mM EDTA), protein amount was determined using the bicinchoninic acid assay

(Thermo Scientific, Rockford, IL), and microsomes were stored at ‐20 °C until used.

Stock solutions were made as follows: 60 µM equimolar mixture of eight acyl‐CoAs

(14:0, 16:0, 18:0, 18:1, 18:2, 20:4, 20:5 and 22:6) in 100% methanol, 200 µM equimolar mixture of six lysophospholipids (LPA, LPC, LPE, LPG, LPI and LPS) in assay buffer

(prepared and sonicated immediately prior to the assay); 1.25 mM fatty acid‐free bovine serum albumin (BSA) in water; and internal standard mixture of 10 ng/µl each of

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[2H31]16:0/18:1‐PA, [2H31]16:0/18:1‐PC, [2H31]16:0/18:1‐PE, [2H31]16:0/18:1‐PG,

[2H31]16:0/18:1‐PI, and [2H31]16:0/18:1‐PS in methanol. The final concentrations of the reaction components were: 10 µg total protein from microsomes, 3 µM LPC, LPE, or LPS, 3

µM arachidonoyl‐CoA, and 12.5 µM BSA, in assay buffer to a total volume of 200 µl. The

CoA ester solution was made in methanol, leading to a concentration of 5 % (v/v) in the final reaction. The acyltransferase assay was performed at 37 °C for 10 min (225). The reaction was stopped with 750 µl of methanol: chloroform (2:1, v/v), internal standard mixture (2.5 µl) was added and products were extracted by the Bligh and Dyer method

(226). After the samples were dried under a stream of nitrogen, they were resuspended in

100 µl of 75 % solvent A (isopropanol: hexanes 3:4, v/v) and 25 % solvent B (isopropanol: hexanes: water 3:4:0.7, v/v/v, containing 5 mM ammonium acetate). Samples were analyzed by liquid chromatography coupled to tandem mass spectrometry (LC/MS/MS) as described below.

Liquid Chromatography/Mass Spectrometry for Phospholipids– For normal phase separation, samples were injected onto an Ascentis‐Si HPLC column (150 x 2.1 mm, 5 µm;

Supelco with Sigma‐Aldrich) at a flow rate of 0.2 ml/min at 25 % solvent B. Solvent B was maintained at 25 % for 5 min, increased to 60 % over 10 min, and then to 95 % over 5 min.

The system was held at 95 % B for 20 min prior to re‐equilibration at 25 % for 14 min. For reversed phase separation, solvent C was methanol/acetonitrile/water, 60/20/20 (v/v/v), containing 2 mM ammonium acetate, and solvent D was methanol containing 2 mM ammonium acetate. The samples were injected onto an Ascentis‐C18 HPLC column (150 x

2.1 mm, 5 µm; Supelco with Sigma‐Aldrich) at a flow rate of 0.2 ml/min at 75 % solvent D.

Solvent D was maintained at 75 % for 1 min and increased to 98 % over 5 min. The system was held at 98 % D for 20 min prior to re‐equilibration at 75 % for 10 min. Phospholipid products of the LAT assay were measured using an API4000 triple quadrupole mass

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spectrometer (AB Sciex, Thornhill, CA) in negative ion mode using multiple‐reaction‐ monitoring (MRM) of the m/z transitions shown in Table 3‐1. Quantitation was performed using AB Sciex MultiQuant software and using 12:0/12:0‐PA, 12:0/12:0‐PC, 12:0/12:0‐PE,

12:0/12:0‐PG, 8:0/8:0‐PI and 14:0/14:0‐PS reference standards for dilution curve analysis, as described previously.

RAW 264.7 Cell Stimulation and Eicosanoids Sample Preparation– Prior to plating, the cells were lifted and counted. Cells were plated in the normal growth medium at a density of 2 x106 cells per 6‐well, and allowed to attach overnight in humidified air with 5

% CO2 at 37 °C. After 16 hours, the cellular medium was removed, the cells were rinsed with 1x PBS, and 1 ml of 1x HBSS with Ca2+ and Mg2+ was placed on the cell. Cell stimulated with ATP received a final concentration of 2 mM ATP. Cells were allowed to incubate with the ATP for 0, 1, 5, 15, 30, or 60 min. Cells were stimulated with a final concentration of 10

μM ionomycin (0.5% ethanol) for 15 min. Cells stimulated in the presence of AA were performed as followed: cells undergoing vehicle treatment were incubated with 0.5% ethanol whereas cells with AA were incubated with AA (final ethanol concentration of

0.5%) for 1 min. After the initial incubation, cells were stimulated with 2 mM ATP for 15 min. At the conclusion of the experiment the reaction was stopped by addition of 1 ml of

100% methanol that contained a mixture of the eicosanoid internal standards. Samples were stored overnight at ‐20°C to accelerate protein precipitation. Samples were spun at

3000 RPM to pellet the protein precipitants, and the supernatant was extracted using

Strata‐X polymeric solid‐phase extraction cartridges (Phenomenex, Torrance, CA), per manufacture’s protocol. Samples were eluted in 100% methanol, dried under vacuum centrifugation, and resuspended in reverse phase starting conditions (67% solvent A: water with 0.05% acetic acid pH 5.7 with ammonium hydroxide; 33% solvent B: 35/65 methanol/acetonitrile).

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Liquid Chromatography/Mass Spectrometry for Fatty Acids and Eicosanoids– For reverse phase separation, samples were injected onto a Kinetex C‐18 HPLC column (50 x

2.1 mm, 5 µm; Phenomenex) at a flow rate of 0.2 ml/min at 40 % solvent B. Solvent B was maintained at 40% for 1 min, increased to 75 % over 5 min, and then to 98 % over 1 min.

The system was held at 98 % B for 7 min prior to re‐equilibration at 40 % for 5 min. The following list of negative ion MRM transitions were monitored: LTC4 m/z 624.5 → 272.2,

LTD4 m/z 495.4 → 177.1, LTE4 m/z 438.3 → 333.3, PGE2 m/z 351.3 → 271.3, PGD2 m/z

351.3 → 233.2, 5‐HETE m/z 319.3 → 115.1, AA m/z 303.2 → 205.2, DHA m/z 327.3 →

283.2, d5‐LTC4 m/z 629.5 → 272.2, d5‐LTD4 m/z 500.4 → 177.1, d5‐LTE4 m/z 443.3 → 338.3, d4‐PGE2 m/z 355.3 → 275.3, d8‐5‐HETE m/z 327.3 → 116.1, and d8‐AA m/z 311.2 → 267.2.

Thin Layer Chromatography and Incorporation of [3H]‐AA into RAW264.7 Cell–

Prior to plating the cells, the cells were lifted and counted. Cells were plated in the normal growth medium at a density of 2 x106 cells per 6‐well, and allowed to attach overnight in humidified air with 5 % CO2 at 37 °C. After 16 hours, the cellular medium was removed, the cells were rinsed with 1x PBS, and 1 ml of 1x DMEM with 0.5 μCi of [3H]‐AA (100 Ci/mmol) was placed on the cell. The cells were incubated with the [3H]‐AA for 2 hours. At the conclusion of the experiment, the supernatant was collected, and 2:5 1x PBS: methanol was added to the adhered cells. Cells were scraped and each well was rinsed with 2:5 1x PBS: methanol. The two rinses were added together, and Bligh‐Dyer was performed (226).

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Results–

Confirmation of MBOAT7/LPIAT1 shRNA Targeted Decrease in the RAW 264.7

Cell– Two independent shRNA constructs were used to decrease the expression of

MBOAT7/LPIAT1 in the RAW264.7 cell. The decrease in mRNA expression was confirmed by qRT‐PCR. Compared to the RAW264.7 cell that contained a non‐targeted shRNA, there was a 50% decrease in mRNA expression with shRNA #1, and a 40% decrease with shRNA

#2 in regards to RNA expression (Figure 5‐1A, MBOAT7/LPIAT1 mRNA Expression).

Lysophospholipid acyltransferase enzymatic assay, previously developed in

Chapter 3, was used to measure changes in phospholipid biosynthesis from a mixture of lysophospholipids and acyl CoA esters. In this enzymatic assay, microsomal preparations known to contain several LPATs were made from each of the cell lines: non‐targeted shRNA, MBOAT7/LPIAT1 shRNA #1, and MBOAT7/LPIAT1 shRNA #2. These microsomal preparations were incubated with six non‐endogenous lysophospholipids (17:1‐LPA, ‐LPC,

‐LPE, ‐LPG, LPI, and –LPS) and eight fatty acyl CoA esters (14:0‐, 16:0‐, 18:0‐, 18:1‐, 18:2‐,

20:4‐, 20:5‐, 22:6‐CoA), and the 48 possible unique phospholipid products were detected using liquid chromatography tandem mass spectrometry (LC‐MS/MS) in the MRM mode

(225).

Collectively the newly synthesized phospholipid products were analyzed by normal‐phase LC‐MS/MS in multiple reaction monitoring (MRM) mode. To ensure that the correct identification of a product of the reaction, each phospholipid product was identified by their co‐elution with deuterated internal standards ([d31]‐palmitoyl‐oleoyl phospholipids) that were added at the termination of the experiment. Along with validation of each lipid class with co‐elution by chromatography, MRM experiments were used to validate precursor‐product relationships of the parent phospholipid ion to the carboxylate fatty acyl ion. A newly synthesized phospholipid had to fit all three criteria (co‐

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elution with a deuterated class standard, parent ion, and product ion) to be considered a valid product of the reaction. The deuterated internal standards were also used to quantitate the absolute amount of the newly synthesized phospholipids by comparison of the area of the analyte signal and internal standard signal and a standard curve separately determined. The internal standard was always added in the same known concentration to each sample, and separately compared to various quantities of non‐deuterated phospholipid standards for accurate quantitation [calibration curves].

The data in Figure 5‐1E contains the raw chromatographic traces from the production 17:1/20:4‐PI that was produced by the microsomes after they were incubated for 10 min with the mixture of the six 17:1 lysophospholipids and the eight fatty acyl‐CoA esters. The y‐axis displays raw ion counts measured by the detector in the mass spectrometer. The x‐axis displays the retention time for the corresponding MRM transition. At the end of the incubation, deuterated standards, 2H31‐16:0/18:1 PI, were added to allow for comparison and quantitation (blue trace).

The production of 17:1/20:4‐PI was then compared between microsomes from cells containing a non‐targeted shRNA and microsomes from cells that contained the two different shRNA against MBOAT7/LPIAT1 (red traces). After 10 min of incubation, the non‐ targeted shRNA microsomal preparations had made a significantly higher level of

17:1/20:4‐PI compared to the microsomal preparation from cells that contained an shRNA against MBOAT7/LPIAT1. The difference in production can be observed by the decrease in signal intensity counts (intensity, cps) as well a comparison of the ratio between the d31‐

16:0/18:1 PI standard (blue traces) and the 17:1/20:4‐PI peak (red traces). Note the signal for the internal standard, [d31]‐16:0/18:1‐PI, did not change.

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A" MBOAT7/LPIAT1.mRNA.Expression. B" 17:1/20:4+PI.ProducJon. 1.2 3

1.0

0.8 2 * ** ** 0.6 ***

0.4 product of 1 Fold Change pg 0.2

0.0 0 Non-Targeted shRNA #1 shRNA #2 Non-Targeted shRNA #1 shRNA #2 C" D" 1.5 17:1/20:4+PC.ProducJon. 20 Microsomal.POPC.

15 1.0

10 of product product of 0.5 of product product of pg 5 pg

0.0 0 Non-Targeted shRNA #1 shRNA #2 Non-Targeted shRNA #1 shRNA #2 E" Non+targeted.shRNA. MBOAT7/LPIAT1.shRNA.#1. MBOAT7/LPIAT1.shRNA.#2. 5915 5915 5915

17:1/20:4+PI.

[d31]16:0/18:1+PI. 17:1/20:4+PI. [d31]16:0/18:1+PI. [d31]16:0/18:1+PI. 17:1/20:4+PI. Intensity, cps Intensity,

10 15 20 25 10 15 20 25 10 15 20 25 Time, min Time, min Time, min Figure 5‐1: MBOAT7/LPIAT1 shRNA targeted cells have a decrease in mRNA and enzymatic activity. A) mRNA measurement relative to the non‐targeted control cells. B&C) Measurements of newly synthesized non‐endogenous phospholipids from the dual‐choice assay. D) Measurement of endogenous lipids found on the microsomes of the dual choice assay. E) Raw data traces showing the production of the non‐endogenous 17:1/20:4‐PI produced relative to the internal standard [d31]‐16:0/18:1‐PI that was used for quantitation. n=8, Average ± SEM. Students’ t‐test. ***= p<0.001, **= p<0.01, *= p<0.05.

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A semi‐quantitative approach was taken to validate the specific changes in the formation of 17:1/20:4‐PI. When the non‐targeted shRNA microsomes were compared over multiple samples, there was a significant decrease in the production of 17:1/20:4‐PI from the microsomes derived from cells with a targeted decrease of MBOAT7/LPIAT1 activity (Figure 5‐1B).

The formation of another AA contain phospholipid product that is not made by

MBOAT7/LPIAT1 was also analyzed in a similar manner. No significant change was found in the production of the 17:1/20:4‐PC in the microsomal preparations from the

MBOAT7/LPIAT1 shRNA targeted cells (Figure 5‐1C). This 17:1/20:4‐PC product could be made by MBOAT5/LPCAT3, but should not change with the MBOAT7/LPIAT1 shRNA targeted cells. An endogenous lipid found in the microsomes (16:0_18:1‐PC) was measured to confirm that there were no changes in the amount of microsomal preparation present in the reaction, but that the data was truly representative of the deletion of MBOAT7/LPIAT1 activity from the microsomal preparations derived from these cells. No significant changes were observed in the endogenous 16:0_18:1‐PC (Figure 5‐1D).

The data from the LPAT choice assay was graphed in an alternative fashion to reveal if there were any changes in all 48 phospholipid products between the non‐targeted microsomes and the MBOAT7/LPIAT1 shRNA #1 targeted cells (Figure 5‐2). The synthesis of the phospholipids in the MBOAT7/LPIAT1 shRNA #1 were subtracted from the phospholipid products of the non‐targeted microsomal preparations to determine the change in phospholipid products. All negative numbers supported decreases in the LPAT activity in the cells with an shRNA against MBOAT7/LPIAT1. All positive numbers supported the gain of LPAT activity with an shRNA against MBOAT7/LPIAT1.

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14:0 16:0 18:0 18:1 PA# 18:2 20:4 20:5 22:6 14:0 16:0 18:0 18:1 PC# 18:2 20:4 20:5 22:6 14:0 16:0 18:0 18:1 PE# 18:2 20:4 20:5 22:6 14:0 16:0 18:0 18:1 PG# 18:2 20:4 20:5 22:6 14:0 16:0 18:0 18:1 PI# 18:2 20:4 20:5 22:6 14:0 16:0 18:0 18:1 PS# 18:2 20:4 20:5 22:6 -2 -1 0 1 2 Δ#pg#of#product#

Figure 5‐2: Phospholipid molecular species production during reduced expression of MBOAT7/LPIAT1 in microsomes. RAW 264.7 cells were transduced with either a non‐ targeting shRNA construct or an MBOAT7/LPIAT1 shRNA #1, and microsomes were used in the dual substrate choice acyltransferase assay. Four independent microsomal preparations were assayed. All 48 phospholipids products were quantitated by tandem mass spectrometry and normalized by microsomal protein content. Each phospholipid molecular species produced by MBOAT7/LPIAT1 shRNA #1 cells was subtracted from that measured in non‐targeted shRNA cells. The results are expressed as a histogram for each phospholipid molecular species as a less abundant product (bars going left of center) or more abundant product (bars going right of center).

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MBOAT7/LPIAT1 shRNA #1 and #2 microsomal preparations showed specific changes in the incorporation of AA into PI, which was noticeably reduced by this graphical display (Figure 5‐2). The only other notable change was the decrease in the incorporation of 14:0, 16:0, and 18:2 into phosphatidic acid (PA). MBOAT7/LPIAT1 is not known to have a direct effect on the formation of PAs. Why or how shRNA against MBOAT7/LPIAT1 would alter activity of an LPAT that incorporates these acyl chains into lyso PI is not known, but since reacylation of AA into PA after cell stimulation is not a robust mechanism this decrease of activity was not concerning to test the affect on MBOAT7/LPIAT1 on leukotriene formation.

Targeted Decrease of MBOAT7/LPIAT1 Led to Decreases in Leukotriene C4

Production– Once the decrease in MBOAT7/LPIAT1 was confirmed for the two independent shRNA constructions by both qRT‐PCR and enzymatic activity assay, the cells were challenged to examine how a deficiency in MBOAT7/LPIAT1 would affect the production of leukotrienes. Cells were initially challenged with 2 mM ATP, activating purinergic receptors on the surface of the RAW 264.7 cell, which led to an increase in cytosolic calcium. This in turn activated cPLA2α and 5‐LO to release AA and produce leukotrienes.

At the termination of the ATP stimulation, eicosanoids and fatty acids were isolated and analyzed by liquid chromatography tandem mass spectrometry in MRM mode. To ensure that the correct analyte was measured, eicosanoid products were identified by their co‐elution with a precise amount of corresponding deuterated internal standard that was added at the termination of the experiment. The deuterated internal standards were also used to quantitate the production of eicosanoids by comparison of the area of the analyte and internal standard area. The internal standard was always added in the same known

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concentration to each sample, and separately compared to non‐deuterated reference standards for accurate quantitation.

The traces in Figure 5‐3 show the production of LTC4 (blue trace) that was found by the cells after they were stimulated with ATP for 30 min. In each case the LTC4 signal co‐ eluted with the deuterated standard that was added, d5‐LTC4 (red trace). The production of

LTC4 was then compared between cells that had a non‐targeted shRNA to those cells that had an shRNA against MBOAT7/LPIAT1. After 30 min of stimulation with ATP, the non‐ targeted shRNA cells made a significantly higher level of LTC4 as compared to those cells that contained an shRNA against MBOAT7/LPIAT1. The difference in production can be observed by the decrease in signal intensity counts (intensity, cps) as well as a comparison of the ratio between the d5‐LTC4 standard and the LTC4 peak.

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Non-targeted shRNA LPIAT1 shRNA #1 LPIAT1 shRNA #2

5.0e4 5.0e4 5.0e4

LTC4

LTC4 LTC4 Intensity, cps Intensity, d -LTC d5-LTC4 5 4 d5-LTC4

3.5 4.0 4.5 5.0 3.5 4.0 4.5 5.0 3.5 4.0 4.5 5.0 Time, min Time, min Time, min

Figure 5‐3: LTC4 production is reduced in RAW 264.7 cells that have decreased MBOAT7/LPIAT1 expression when stimulated by ATP. LTC4 production (blue trace) after 30 min of stimulation by ATP in the non‐targeted control cells and the MBOAT7/LPIAT shRNA targeted cells. After the experiment a d5‐LTC4 standard was added for validation and quantitation of LTC4 in the cells (red trace). The unprocessed data is displayed in detector counts (Intensity, cps) and relative to the retention time of LTC4 and the d5‐LTC4 standard (Time, min).

An Early Decrease in AA Release Led to a Decrease in 5‐lipoxygenase Products–

From the data presented in Figure 5‐3, stimulated cells that had decreased

MBOAT7/LPIAT1 activity made less leukotrienes at a single 30 min time point, but additional experiments were needed to determine if it was a change in the rate or overall production of leukotrienes. Non‐targeted shRNA cells and cells that contained shRNAs against MBOAT7/LPIAT1 were stimulated at various time points. In cells that contained shRNA targeted against MBOAT7/LPIAT1, there was a decrease in LTC4 formation at 5, 15,

30, and 60 minutes post‐stimulation with ATP as compared to the stimulated non‐targeted shRNA containing cells (Figure 5‐4A). The decrease in LTC4 formation correlated well with

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the decrease in mRNA and activity of MBOAT7/LPIAT1, and was more dramatic with shRNA construct #1. In addition to the measurement of LTC4, 5‐hydroxyicosatetraenoic acid (5‐HETE) was measured. 5‐HETE is a byproduct of 5‐LO that is produced from the hydrolysis of the first reaction product 5‐hydroperoxyeicosatetraenoic acid (5‐HpETE).

The production of 5‐HETE at 5 and 15 minute was significantly less in the cells that contained shRNA targeted against MBOAT7/LPIAT1 than that of the production of 5‐HETE in the non‐targeted shRNA cells (Figure 5‐4B). The corresponding change in 5‐HETE and

LTC4 supported an overall decrease of 5‐LO products when MBOAT7/LPIAT1 was decreased in the RAW 264.7 cell.

Since both 5‐HETE and LTC4 come from AA, the levels of freed AA were measured at various time points. There was a significant decrease in the amount of free AA at the 5 minute time point in the cells that contained shRNA targeted against MBOAT7/LPIAT1 compared to the non‐targeted shRNA control cells (Figure 5‐4C). The peak release of AA appeared to be between 1 and 15 minutes in the RAW 264.7 cell, so changes at the 5 minute time points may not completely define the reduction of free AA in the

MBOAT7/LPIAT1 shRNA cells compared to the non‐targeted shRNA cells. To confirm that this was a specific change to the release of AA, an additional fatty acid, docosahexaenoic acid (DHA) was measured. No significant changes were observed in levels of DHA at any of the time points after ATP stimulation (Figure 5‐4D). Decreased AA availability could lead to decreased 5‐LO product formation. The data in Figure 5‐4 suggested specific changes in

AA due to the decrease in MBOAT7/LPIAT1 expression that may explain the decrease in 5‐

LO product formation.

Reduction in ACSL4 Led to Increased Levels of Free AA but No Change in 5‐

Lipoxygenase Products– The previous finding that the targeted decrease of

MBOAT7/LPIAT1 led to decreased amounts of free AA after cell stimulation led to the

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hypothesis that RAW 264.7 cells were not able to reacylate AA (leaving it in the non‐ esterified form) would result in an increased leukotriene production. To test this hypothesis, shRNAs against ACSL4 were used to decrease expression of the major ACSL thought to regulate AA. The targeted decrease of ACSL4 was validated by measuring mRNA message using qRT‐PCR. Both shRNA constructs against ACSL4 showed a significant reduction at the mRNA level (Figure 5‐5A). ACSL4 shRNA #1 was approximately a 60% reduction and shRNA #2 was approximately a 70% reduction at the mRNA level compared to the non‐targeting control. Other genes in the pathway were tested to confirm that the effects were due to changes in ACSL4. No change in any mRNA expression level was observed for 5‐LO, MBOAT5/LPCAT3, MBOAT7/LPIAT1, ACSL1, cPLA2α (data not shown).

Once the decrease in mRNA expression in the RAW 264.7 cells was confirmed, the cells were challenged with ATP for different amounts of time. Eicosanoids were collected and confirmed as previously described. As anticipated, the stimulated cells had significantly higher amount of free AA and a delayed reacylation of AA (Figure 5‐5B).

Similar to AA, free DHA was significantly increased and there was a delay in the reacylation of the free fatty acid (Figure 5‐5C). However, the changes in the levels of AA did not translate into increased leukotriene production as assessed by LTC4 production (Figure 5‐

5D). 5‐HETE did significantly increase, only at the 15 min time point, in the stimulated cells that contained shRNA targeted against ACSL4, but at all other time points there was no describable difference (Figure 5‐5E).

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A" Leukotriene C B" 5-HETE 4 Non-Targeted shRNA #1 shRNA #2 Non-Targeted shRNA #1 shRNA #2 60 3.5 *** 3.0

2.5 40 * cells 6 * 2.0 *** cells 6 * 1.5 ng/1 x10 20 1.0 *** ng/1 x10 *** *** 0.5 * 0 0.0 0 10 20 30 40 50 60 0 10 20 30 40 50 60

Time, min Time, min

C" Arachidonic Acid D" Docosahexaeneoic Acid 80 Non-Targeted shRNA #1 shRNA #2 30 Non-Targeted shRNA #1 shRNA #2 ***

60 20 cells cells

6 40 6

10 ng/1 x10 20 ng/1 x10

0 0 0 10 20 30 40 50 60 0 10 20 30 40 50 60

Time, min Time, min

Figure 5‐4: Specific changes in AA release at early time points mirror later reduced production in LTC4 and 5‐HETE in RAW 264.7 cells that have decreased MBOAT7/LPIAT1 expression. Cells were stimulated for different lengths of time with ATP, and four analytes were measured. N=10‐12, Average ± SEM. Two‐way ANOVA with Tukey’s post‐test. ***= p<0.001, **= p<0.01, *= p<0.05.

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Targeted Decrease of MBOAT7/LPIAT1 Does Not Change Other Lands Cycle or

Eicosanoid mRNA Levels– Since the targeted depletion of MBOAT7/LPIAT1 led to decreases in AA, LTC4 and 5‐HETE, the levels of essential proteins in the pathway that form the leukotriene products were analyzed to confirm that they did not change. Since the first step in leukotriene synthesis is the release of AA, two pathways were evaluated to determine the dominant route of AA hydrolysis after cellular stimulation. The two most prevalent pathways that are used to release AA from phospholipids are cPLA2α and

DAG/MAG lipase pathway (249,250). Using the inhibitor pyrrolidine, it was determine that the predominate pathway for AA release in the RAW 264.7, by activation of the puringeric receptors with ATP, was through the activation of cPLA2α (data not shown). A secondary measurement of the MAG lipase pathway was measured by mRNA message and no mRNA message could be detected for MAG lipase in any of the RAW 264.7 cells.

Since the data supported cPLA2α as the dominant pathway for AA release in the

RAW 264.7 cells, mRNA message and protein was assessed. No change in the expression of mRNA or protein was observed in cPLA2α with the depletion of MBOAT7/LPIAT1 (Figure

5‐6A&B). Additional genes including 5‐LO, MBOAT5/LPCAT3, ACSL4, and ACSL1 also had no significant change in the mRNA expression with the depletion of MBOAT7/LPIAT1

(Figure 5‐6C). From the data presented in Figure 5‐6, no change of mRNA expression of the proteins essential for the formation of LTC4 or other enzymes in the reacylation pathway could be detected, and therefore did not explain the reduction in amount of free AA or LTC4 production.

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A" ACSL4 mRNA Expression 1.2

1.0

0.8

0.6 *** 0.4 Fold Change *** 0.2

0.0

ACSL4 shRNA 1 ACSL4 shRNA 2

Non-targeted shRNA B" C" Free DHA Free AA 70 Non-targeted shRNA 140 *** ACSL4 shRNA #1 *** 60 120 ACSL4 shRNA #2 Non-targeted shRNA 50 *** ACSL4 shRNA #1 100 *** *** ACSL4 shRNA #2

cells 40 6

cells 80 6 30 *** 60 *** ** ng/1x10

ng/1x10 ns 40 20 ns 20 10

0 0 0 10 20 30 40 50 60 0 10 20 30 40 50 60 Time, min Time, min D" E" 5-HETE production LTC4 production 4 70 Non-targeted shRNA ** ACSL4 shRNA #1 60 3 ACSL4 shRNA #2 50 cells 6

cells 40

6 2 30 ng/1x10

ng/1x10 Non-targeted shRNA 20 1 ACSL4 shRNA #1 10 ACSL4 shRNA #2

0 0 0 10 20 30 40 50 60 0 10 20 30 40 50 60 Time, min Time, min Figure 5‐5: Targeted decrease of ACSL4 led to increases in freed AA after stimulation but no change in leukotriene formation. A) ACSL4 mRNA measurements were made and compared to cells that had a non‐targeted shRNA. B‐E) Cells were stimulated for different lengths of time with ATP, and four analytes were measured. N=8‐12, Average ± SEM. Two‐ way ANOVA with Tukey’s post‐test. ***= p<0.001, **= p<0.01, *= p<0.05.

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Loss of MBOAT7/LPIAT1 is Not Recovered by Adding Exogenous AA or by

Bypassing cPLA2α Activity– Previous stimulations were all performed without external addition of AA, and using a purinergic receptor agonist to stimulate the cells. Since the activation of RAW 264.7 cells with decreased MBOAT7/LPIAT1 expression led to decreased AA release, experiments were performed to bypass the release of AA. If depletion of MBOAT7/LPIAT1 led to decreased AA containing phospholipids then adding high exogenous concentrations of AA just prior to cell stimulation should recover the LTC4 production to the level produced in the non‐targeted shRNA cells. Experimental controls were performed using ATP with no AA addition and compared to the stimulated cells that were treated with exogenous AA before stimulated with ATP. The AA was added immediately before the experiment to limit the incorporation of the exogenous AA into the cellular lipids. However adding exogenous AA to the cell was not sufficient to recover the production of leukotrienes in the MBOAT7/LPIAT1 cells compared to the non‐target shRNA containing cells (Figure 5‐7, A and C).

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A" B" MBOAT7# 1.4 #1# #2# 1.2 kDa# Control# shRNA 250# shRNA 1.0 150# 0.8 100# cPLA2α# 0.6 75#

Fold Change 0.4 50# 0.2 β?Ac@n# 0.0 37# Non- shRNA #1 shRNA #2 Targeted C" 1.2

1.0

0.8

0.6

Fold Change 0.4

0.2

0.0 NT #1 #2 NT #1 #2 NT #1 #2 NT #1 #2

5-Lipoxygenase ACSL1 MBOAT5 ACSL4

Figure 5‐6: Targeted decrease of MBOAT7/LPIAT1 did not lead to changes in expression of other reacylation or eicosanoid enzymes. A and C) mRNA measurements were made and compared to cells that had a non‐targeted shRNA. B) Protein analysis of cPLA2α. n=8, Average ± SEM. One‐way ANOVA with Tukey’s post‐test.

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Since the addition of exogenous AA did not recover the LTC4 production in the

MBOAT7/LPIAT1 targeted shRNA cells, experiments were performed to exclude the possibility that ATP acting on the purinergic receptor was the cause of the decrease in AA release and leukotriene formation. Ionomycin is a calcium ionophore that allows large amounts of calcium from the extracellular space to enter into the cytosol. Therefore, cPLA2α would be activated independently from the receptor‐mediated stimulus of ATP. Treatment of the RAW 264.7 cells with ionomycin, for the same amount of time as ATP, yielded no change in the production of LTC4 in the cells with decreased MBOAT7/LPIAT1 expression compared to the non‐targeted shRNA control (Figure 5‐7 A and E). A similar finding was observed when analyzing the other 5‐LO products, 5‐HETE (Figure 5‐7 B, D, F). Taken together the data supports that the reduction in released AA and production of LTC4 is not a signaling defect of the purinergic receptor nor can the defect be repaired by acute exposure to exogenous AA before stimulation.

Molecular Species of PI Do Not Change with the Depletion of MBOAT7/LPIAT1–

The finding from the experiments using the addition of exogenous AA were similar to the findings in the cells with the targeted decrease of ACSL4. In both experiments, increased levels of AA had no effect on leukotriene formation. This unexpected finding suggested that there may be a very specific class of lipid that is changing to account for the decreased production of leukotrienes. Since AA is released from numerous phospholipids and changes in MBOAT7/LPIAT1 could ultimately affect several phospholipid classes through the remodeling pathways, a global approach to analyze phospholipids was taken. Initial evaluation of PI (Figure 5‐8) supported that there was no change in any of the abundant PI molecular species. A notable finding however was that the most abundant PI anion was m/z 887. In most mammalian systems the m/z of 885 which corresponds to 18:0/20:4‐PI is the most abundant, constituting up to 70% of the total PI species mass.

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A" LTC4 ATP B" 5-HETE ATP

100 100

80 80

60 60

40 40

20 20 Percent of Non-Targeted Non-Targeted Percent of Percent of Non-Targeted Non-Targeted Percent of 0 0 Non- Targeted shRNA #1 shRNA #2 Non- shRNA #1 shRNA #2 Targeted

C" LTC4 AA ATP D" 5-HETE AA ATP 100 100

80 80

60 60

40 40

20 20 Percent of Non-Targeted Non-Targeted Percent of Percent of Non-Targeted Non-Targeted Percent of 0 0 Non- Non- Targeted shRNA #1 shRNA #2 Targeted shRNA #1 shRNA #2 E" F" 5-HETE Ionomycin LTC4 Ionomycin 100 100

80 80

60 60

40 40

20 20 Percent of Non-Targeted Non-Targeted Percent of Percent of Non-Targeted Non-Targeted Percent of 0 0 Non- Non- shRNA #1 shRNA #2 Targeted shRNA #1 shRNA #2 Targeted

Figure 5‐7: Deficiency in LTC4 production in the targeted decrease of MBOAT7/LPIAT1 could not be recovered with addition of exogenous AA or by non‐receptor mediated stimulus. Data was compared to the production of the non‐targeted cells. Cells were stimulated for 15 min under each condition, and two analytes were measured. n=8, Average ± SEM.

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Fragmentation of m/z 887 was performed and the fragment ions supported the most abundant species as an 18:0/20:3‐PI molecular species. The fatty acid 20:3 is not often observed in mammalian systems because of the abundance of ω‐6 fatty acids in the diet. However, cells in culture usually do not have sufficient amounts of essential fatty acids to sustain their rapid proliferation. When cells have insufficient amounts of essential fatty acids they will compensate by synthesizing a 20:3 ω‐9 fatty acid from the precursor oleic acid (240). This shift from the ion m/z 885 to 887 could potentially explain why all the cells, including the non‐targeted cells, have an abnormal distribution of PI lipids. From analysis and semi‐quantitation of the parent ion scan data, there appears to be no change in any of the phospholipid molecular species with the depletion of MBOAT7/LPIAT1.

RAW%264.7%Phospha'dylinositol%Species% 9" Non/targe5ng"shRNA" MBOAT7"shRNA"#1" MBOAT7"shRNA"#2"

8"

7"

6"

Area%Ra'o% 5"

4"

3"

2"

1"

0" 835" 847" 859" 861" 863" 871" 873" 875" 883" 885" 887" 889" 899" 901" Mass+to+Charge%Ra'o%

Figure 5‐8: Quantitation of PI molecular ions does not reveal any changes due to the loss of LPIAT1. Molecular ions of the six major phospholipid classes were analyzed relative to an internal standard of the same head group class (Area Ratio). The MBOAT7/LPIAT1 targeted shRNA cells (red and green bars) showed no difference compared to the non‐ targeting shRNA cells (blue bars). n=8, Average ± SEM.

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Exogenously Added AA is Almost Entirely Recycled into PI, PC, and PE in the

Parental RAW 264.7 Cell– Since there were no changes in molecular species of phospholipids observed due to the depletion of MBOAT7/LPIAT1, and addition of exogenous AA did not recover the leukotriene production compared to the non‐targeted shRNA control cells, experiments were performed using [3H]‐AA to determine the fate of exogenously added AA in the RAW 264.7 cell. In human neutrophils, 55% of exogenous AA is incorporated into neutral lipids (231). To determine if the RAW 264.7 cell behaves in a similar manner, exogenous [3H]‐AA was added to the parental cell line and allowed to incorporate for 2 hours. After the cells were incubated with the [3H]‐AA, the supernatant removed was counted. Approximately 20% of the counts were found in the supernatant.

The adherent cells were collected and extracted by the Bligh‐Dyer method (226). The counts of the aqueous phase of the Bligh‐Dyer were negligible. The majority of the counts were in the organic phase (80%).

The organic phase of Bligh‐Dyer was run on two different thin‐layer chromatography systems for the separation of neutral lipids and one for phospholipids.

After the plates were run, the radioactivity was counted and plotted based on migration.

Non‐radioactive standards were run to confirm the migration of each class of lipid and are plotted with arrows indicating the distance they migrated. With the two different thin‐ layer chromatography systems it appeared that almost all of the counts in the organic layer had been incorporated into the phospholipid pool (Figure 5‐9 A&B). There was no migration of the radioactivity from the origin of the neutral lipid thin‐layer chromatography system, supporting that the majority of [3H]‐AA had been incorporated into phospholipids.

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8000 A" 500 B" PE 400 6000 300 $ PA/PI/PS PG $ 4000 cpm cpm 200 PC 2000 100 DAG AA DAG TAG AA 0 0 0 50 100 150 200 0 50 100 150 200

Migra*on,$mm$ Migra*on,$mm$ 15000 C"

16:0 (x2)-PG 16:0/18:1-PS 10000 Soy PI 18:0/20:4-PA 16:0/18:1-PE $ cpm

5000 16:0/20:4-PC AA

0 0 1 2 3 4 5 6 7 8 9 10 11 12 13 14 15 16 17 18 19 20 21 22 23 24 25 26 27 28 29 30 31 32 33 34 35 36 37 38 39 40

Time,$min$

Figure 5‐9: Addition of exogenous [3H]‐AA is almost exclusively incorporated into the phospholipid pool. The organic cellular extracts were analyzed using thin‐layer chromatography for phospholipids (A) and neutral lipids (B). The counts per min (cpm) detected were plotted against the migration on the plate. Non‐labeled standards were used to confirm the migration point of each class of lipid and are indicated by the arrows/labels and the arrow under the axis indicates the origin. Further analysis of the phospholipids was performed using normal phase chromatography and collecting one‐minute fractions to determine the distribution of radioactivity. The cpm of each fraction is plotted over the chromatographic time (C). Traces for the non‐radioactive standards (each is labeled with the description on the trace) were run under the same conditions and detect using mass spectrometry are overlaid on the radioactive traces to confirm the identity of the phospholipids that were labeled by the [3H]‐AA.

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The phospholipid thin‐layer chromatography system does not allow for great separation between phospholipid classes as many of them co‐migrate from the trace in

Figure 5‐9A. Therefore, normal phase HPLC chromatography was used to determine the distribution of the [3H]‐AA in the phospholipid pool. The histogram plot in Figure 5‐9C shows the radioactivity counts collected in 1 min fractions. To confirm the identity of the radioactivity, non‐radiolabelled standards were run under the same chromatographic conditions and detected using mass spectrometry. The trace of each standard was overlaid onto the radioactivity bar graph and each standard was labeled. The mass spectrometry traces indicate relative intensity of the molecular ion of each of the phospholipid species.

The PC and AA peaks were expanded to make all the peaks visible. From the data in Figure

5‐9C it appears that most of the radioactive counts are distributed between PI, PE, and PC based on the retention with standards.

Targeted Analysis of AA Containing Global Phospholipid Pools Do Not Change with

MBOAT7/LPIAT1 Depletion– Since the RAW 264.7 cell esterified all of the AA into phospholipids, but there were no changes in the global phospholipid parent ion scans, we used a more sensitive and targeted approach to analyze the AA phospholipid species.

Global phospholipids from non‐targeted shRNA cells and cells that contain shRNA’s against

MBOAT7/LPIAT1 were analyzed using MRM transitions to specifically target lipids that contain AA. A 17:0/20:4‐PI standard was used to normalize all of the phospholipid data.

Table 5‐1 shows the values of the measured lipid compared to the 17:0/20:4‐PI standard.

None of the measured phospholipid molecular species had significant changes with the depletion of MBOAT7/LPIAT1.

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Table 5‐1: Semi‐quantitation of abundant AA containing phospholipids in RAW 264.6 cells.

Non$targeted+shRNA+ LPIAT+shRNA+#1+ LPIAT+shRNA+#2+

16:0/20:4(PC+ 0.14+±+0.07+ 0.13+±+0.05+ 0.14+±+0.07+

18:0/20:4(PC+ 0.12+±+0.05+ 0.11+±+0.06+ 0.12+±+0.05+

18:1/20:4(PC+ 0.13+±+0.06+ 0.12+±+0.07+ 0.13+±+0.05+

16:0/20:4(PE+ 2+±+1+ 1.6+±+0.6+ 2+±+1+

18:0/20:4(PE+ 7+±+2+ 6+±+1+ 6+±+2+

18:1/20:4(PE+ 3+±+1+ 2.6+±+0.7+ 3+±+1+

16:0/20:4(PI+ 0.4+±+0.2+ 0.4+±+0.1+ 0.5+±+0.2+

18:0/20:4(PI+ 5+±+2+ 4+±+2+ 5+±+2+

18:1/20:4(PI+ 1.2+±+0.7+ 1.3+±+0.6+ 1.3+±+0.6+

16:0/20:4(PS+ 0.03+±+0.01+ 0.02+±+0.01+ 0.03+±+0.02+

18:0/20:4(PS+ 0.5+±+0.3+ 0.5+±+0.2+ 0.4+±+0.2+

18:1/20:4(PS+ 0.04+±+0.02+ 0.04+±+0.02+ 0.04+±+0.02+

Numbers+are+displayed+as+area+measurement+of+the+phospholipid+signal+to+the+signal+of+17:0/20:4(PI.+ n=8,+average+±+standard+deviation.+

Discussion–

MBOAT7/LPIAT1 was previously described as an acyltransferase that almost exclusively used lysophosphatidylinositol and AA‐CoA as substrates (191,192). Previous work had demonstrated that non‐selective inhibition of MBOAT7/LPIAT1 by thimerosal in human neutrophils led to an increase in the production of eicosanoids (192). Other groups have also displayed the essential role of MBOAT7/LPIAT1 in the formation of 4,5‐PIP2

(248). To determine a potential role for MBOAT7/LPIAT1 in the production of leukotrienes

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in RAW 264.7 cells, a targeted shRNA approach was used to selectively decrease the mRNA of MBOAT7/LPIAT1.

Quantitation of mRNA, as well as acyltransferase activity assay, was implemented to validate the loss of MBOAT7/LPIAT1. As expected, there was a significant decrease in the production of the 17:1/20:4‐PI. No changes in other AA lysophospholipid activities were observed. However, three PA products (14:0, 16:0, and 18:2) were decreased in the

MBOAT7/LPIAT1 shRNA microsomes compared to the control. MBOAT7/LPIAT1 does not have known activity for PA. The possibility remained that the targeted decrease of

MBOAT7/LPIAT1 led to changes in other acyltransferases. From this data set and the known activity of some of the acyltransferases, it would appear that there might be changes in the expression and activity of LPAAT1 or LPAAT2. However, the analysis of the cell lines with a targeted depletion of MBOAT7/LPIAT1 was pursued because there were no changes in the esterification of AA into other lysophospholipids.

Originally, it was hypothesized that decreased expression of MBOAT7/LPIAT1 or

ACSL4 would lead to increased production of leukotrienes since both proteins are involved in AA recycling. The targeted decrease of ACSL4 had the predicted affect of increasing the amount of free AA. However, that increase in free AA did not translate into LTC4 production. There was an increase of 5‐HETE in the ACSL4 shRNA targeted cells that potentially had several possible explanations. 5‐HETE synthesis does not always correlate with LTC4 synthesis. Three explanations lend the most credence. The first is that free AA can be non‐enzymatically oxidized to 5‐HETE, and having increased amounts of free AA led to increased oxidation product, 5‐HETE. This reason is less likely because other HETE products, which should also be formed in non‐enzymatic oxidation, were not detected in the eicosanoid profile. Another possible reason for the increase in 5‐HETE is that the free

AA is interacting with 5‐LO that has not associated with FLAP. 5‐LO proteins that are free

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in the cytosol may be able to synthesize 5‐HETE, but are extremely inefficient at the production of LTA4 to produce LTC4. A third explanation would be that 5‐HETE itself can be esterified to CoA predominately by ACSL4. Since 5‐HETE is eventually recycled, there may be other ACSLs that have lower 5‐HETE activities but eventually lead to the esterification of 5‐HETE. Phospholipids have been documented to contain esterified 5‐HETE (251). The mechanism of how these phospholipids are generated is unknown, however it is thought to be through a CoA dependent mechanism (251).

Targeted reduction of MBOAT7/LPIAT1 led to decreased amounts of free AA and the production of LTC4. The catabolism and metabolism of PI lipids is complex because they are involved in two independent but related pathways. Once PI is generated from the de novo synthesis pathway, it can be remodeled by PLA2’s and as well as by

MBOAT7/LPIAT1 (pathway depicted in Figure 6‐10). MBOAT7/LPIAT1 and LPAAT3 are currently the only known LPIATs. From activity assays presented in Chapter 3, phospholipid products were formed that are not known products of LPAAT3 or

MBOAT7/LPIAT1 suggesting that there is/are other enzyme(s) that can catalyze acylation of lysophosphatidylinositol, but to date no other LPATs with this activity have been identified. However in the monocyte the most abundant PI molecular species are made through the synthesis by MBOAT7/LPIAT1 and usually contain AA at the sn‐2 position of the phospholipid (230).

After synthesis, AA containing PI can be phosphorylated to form seven different phosphorylated states of PI. The majority of these phosphorylation events take place on the plasma membrane. Many of the kinases that phosphorylate PI have been shown to favor the AA containing species (208). Some phosphorylated forms of PI will eventually be cleaved and used as secondary messengers. The generated DAG has to be recycled back to the endoplasmic reticulum, where the DAG gets the inositol head group. This process of

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phosphorylation and regenerations has been termed the PI cycle, and is depicted in the diagram in Figure 5‐10 (208).

5-LO* Leukotrienes ACSL Nuclear Envelope AA

AA-CoA cPLA2α*

Endoplasmic Reticulum/ trans Golgi LPI Inositol PIS

CDP-DG(18:0/20:4) PI(18:0/20:4) MBOAT7/LPIAT1*

CDS2*

PI4K PA(18:0/20:4)

DGKe* Plasma Membrane

PI(18:0/20:4)-4P DG(18:0/20:4) IP 3 PI4P5K* Second Messengers

PI(18:0/20:4)-4,5P * Specificity for AA Phospholipase C 2 PLC

Figure 5‐10: The regulation of PI molecular species is a mixture of the reacylation pathway and the PI cycle. Products are in black, enzymes specific for AA are in red, and other enzymes are in blue. Dashed lines indicate different compartments, which are labeled.

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To determine the cause of the decrease in AA and LTC4, numerous experiments were performed. One hypothesis was that since MBOAT7/LPIAT1 is involved in a cyclical pathway, decreasing its expression would lead to changes in other proteins in the cycle. No changes were found at the mRNA level in many of the genes that would have been involved in the reacylation pathway. Since cPLA2α was critical in the release of AA in the RAW 264.7 cell and the MBOAT7/LPIAT1 targeted cells had decreases in free AA after stimulations, further analysis was performed to show that there was no change in the protein expression of cPLA2α.

The activity changes in other acyltransferase were also assessed, but no activity changes were observed in the incorporation of AA into other phospholipids. A caveat was that many of the proteins that were measured are enzymes that are regulated through post‐translational modifications and external stimuli that allow for their optimal activity.

Additional analysis is needed to determine if the activity or protein levels of any of these enzymes, measured by mRNA (5‐LO, cPLA2α, ACSL1, and ACSL4), changed.

With no changes in the other proteins in the reacylation pathways, experiments switched focus to eliminate pathways that may involve signaling events through phosphoinositides. The RAW 264.7 cell shows a robust stimulation with the use of ATP, which activates a family of purinergic receptors (252). The purinergic receptor family is a mixture of ion channels and GPCRs that are both expressed on the surface of cells.

Simulation with ATP probably activates multiple receptors at the same time, amplifying the cytosolic calcium to high enough levels to activate 5‐lipoxygenase. To avoid using receptor mediated signaling events, which may depend on phosphoinositides, cells were stimulated with ionomycin, a Ca2+ ionophore. However, receptor independent stimulation of the cells also led to a decrease in the production of leukotrienes in the cells that contained a shRNA against MBOAT7/LPIAT1 compared to the non‐targeted shRNA cells.

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Phosphoinositides have also been suggested to be important for lowering the Ca2+ threshold needed for the translocation, docking, and retention of cPLA2α on the membrane of monocytes (245,253). Also, cPLA2α can hydrolyze AA from PI lipids (100). Exogenous AA was added before stimulation with ATP to bypass the need for the activation of cPLA2α.

Addition of exogenous AA did not restore the decrease in leukotriene production in the cells with a targeted decrease in MBOAT7/LPIAT1.

Since exogenous AA and the targeted decreases in ACSL4 led to no change in LTC4 formation, it seemed likely that very specific pools of AA contain phospholipids are utilized in the formation of leukotrienes. To try and determine which phospholipids were affected by the targeted decrease of MBOAT7/LPIAT1, the global phospholipid pool was analyzed.

No significant changes were found in any phospholipid molecular species. Although there were not changes in the global phospholipid pool, there could be changes in a very small subset of phospholipids, such as in the nuclear envelope, that are critical for the local machinery essential for the production of leukotrienes. In fact, subcellular location of PI molecular species has been studied with the observation that the nuclear PI pool is small compared to the ER or plasma membrane (254).

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CHAPTER VI

THE EFFECT OF ACYL‐COENZYME A SYNTHETASE LONG CHAIN 1 ON ACYL CHAIN

COMPOSITION OF PHOSPHOLIPIDS IN THE MOUSE HEART

Introduction–

Acyl coenzyme A synthetases (ACS) catalyze formation of fatty acyl CoA esters from fatty acids and coenzyme A (CoA). A large number of these enzymes have been reported with various substrate specificities as well as unique expression in different tissues and subcellular locations (255). The chemical reaction catalyzed by ACS involves a high‐energy two‐step mechanism (256). Fatty acyl CoA esters are critical in numerous cellular functions including, but not limited to, beta‐oxidation, fatty acid elongation, fatty acid desaturation, and as substrates for acyltransferases (255).

Of this large family of proteins, thirteen ACS isoforms have been characterized that use long chain fatty acids as substrates (161). These thirteen enzymes each have their own substrate preference; however, all isoforms will use any fatty acid from 16:0 to 22:6 (161).

The work in this chapter will focus on the isoform ACSL1. ACSL1 was previously characterized in vitro to prefer the fatty acid oleate (18:1, ω‐9) (257). The substrate preference was determined in an artificial system using one substrate at a time and measuring enzymatic parameters such as Km or Vmax. This type of assay allowed for the determination of the enzymatic preference of the enzyme; however, these data do not delve into the complexity of substrate availability that exists in biological systems and the potential of substrate competition.

ACSL1 expression has been found in many different cell types and tissues. A limited list of these includes monocytes, adipocytes, liver, lung, brain, and heart. Unlike other ACS proteins that are contained in specific subcellular compartments, in some tissues ACSL1 is found in all subcellular membrane fractions, however in the heart tissue it is

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predominately found in the mitochondrial membrane fraction (258). The specificity of each of the ACSLs most likely has more to do with the availability of substrates in each compartment. To further study the complexity surrounding substrate availability, Dr.

Rosalind Coleman’s laboratory (University of North Carolina) created a global tamoxifen inducible Cre recombinase mouse to decrease the expression of ACSL1 (ACSL1T‐/‐) in most tissue (259). After 10 weeks of the induced gene deletion by treatment with tamoxifen, many cells and tissues were examined for the degree of gene expression changes. It was observed in the heart there was almost complete loss of ACSL1 gene expression, ACSL activity, mitochondrial dysfunction, and left ventricle enlargement (259). Additional gene analysis showed that ACSL1 was the predominate ACSL isoform expressed in the mouse heart and that it was a key regulator of mitochondrial fatty acid oxidation in the heart

(259,260).

Cardiac myocytes are large consumers of oxygen, which contributes to the metabolism of fat in the heart to generate ATP by β‐oxidation. Fasted animals with healthy heart tissue meet 70‐90% of their energy needs by the oxidation of fatty acids (261).

However, as the heart begins to fail this dependence on fatty acids switches to an alternative ATP generating mechanism (261). A diminished use of fatty acids metabolism for energy was observed in the ACSL1T‐/‐ mouse, leading to the hypothesis that ACSL1 is critical in the formation of fatty acyl CoA ester for facilitating entry into the mitochondria for subsequent β‐oxidation. The mitochondria of these mice were no longer able to utilize fatty acids efficiently and had switched to a dependence on glucose and amino acid metabolism for ATP generation (259). Since ACSL1 forms fatty acyl CoA esters required for

β‐oxidation and ACSL1 depletion in the heart led to decreased use of fatty acids for energy production, it was thought that transport of lipids into the mitochondria may be impaired

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and potentially would lead to changes in the lipids available to form other mitochondrion lipid species.

Cardiolipin (CL) is a specific lipid that is found almost exclusively in mitochondrial membrane where it comprises approximately 13‐15% of the total phospholipid content

(262). Cardiolipin is a glycerophospholipid containing four acyl chains, two phosphates and three glycerol moieties. The most commonly found eukaryotic CL molecular species in heart is a tetra‐linoleoyl containing four 9,12‐octadecadienoyl acyl (18:2) chains (Figure 6‐

1) (263,264). The role of cardiolipin is not completely understood, although extremely conserved throughout prokaryotes and eukaryotes (265). There are many suggested roles for cardiolipin in the mitochondrion including interaction with membrane proteins, assembly of respiratory chain supercomplexes, as a proton trap for electron transport, and as a sink for radical oxygen species produced within the mitochondrion (264,266,267).

Insight into the importance of cardiolipin in humans has been observed in the loss of specific molecular species of cardiolipin described in Barth syndrome. Barth syndrome is an X‐linked disorder found primarily in males in which individuals inherit a mutation in the tafazzin gene (268). Although the enzymatic function of tafazzin has not been well characterized, it has been suggested to be a lysophospholipid transacylase, which has a critical role in the proper assembly of cardiolipin (269,270). Individuals who inherit mutated tafazzin are found to have multiple physical manifestations including cardiomyopathy, neutropenia, underdeveloped skeletal musculature and muscle weakness, exercise intolerance, and growth delay (271). The presence of these abnormalities supports the importance of cardiolipin in mitochondrial function in muscle.

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O O

O O O P O O P O O

O OH OH OH O O O O

Figure 6‐1: Tetra‐linoleoyl cardiolipin (18:2/18:2/18:2/18:2‐CL or m/z 1447.9) is the most abundant cardiolipin species in mammalian cardiac muscle.

Additional evidence in heart failure has supported the importance of cardiolipin in cardiac muscle. In healthy rat and human tissue, (18:2/18:2/18:2/18:2‐CL) is the predominant cardiolipin (264). When the heart begins to fail there is a remodeling of the acyl composition of cardiolipins (272). Previous experiments have shown that feeding rats high levels of dietary linoleic acid (safflower oil) reversed the defect in tetra‐linoleoyl cardiolipin and improved the function of the heart (273,274). These findings support the tetra‐linoleoyl species as an important cardiolipin in proper heart function. Considering the finding that ACSL1T‐/‐ mice undergo left ventricular enlargement and mitochondrial dysfunction, our laboratory collaborated with Dr. Coleman laboratory to analyze cardiolipin molecular species in the cardiac tissue of these animals to determine if there were changes due to the loss of ACSL1.

Experimental Procedure–

Sample Preparation for Mass Spectrometry– Total heart samples were homogenized on ice using a Dounce homogenizer in a buffer consisting of 50mM phosphate buffer pH 7.2, 0.1 M NaCl, 2 mM EDTA, 1 mM DTT, and 1 cOmplete Mini

Protease Tab (Roche, Indianapolis, IN). Total protein was determined using the

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instructions included with the bicinchoninic acid assay from Pierce Biotechnology

(Rockford, IL). Heart mitochondrial preparation (180 µg of total protein) or total heart

(300 µg of total protein) was diluted with 1x PBS to a total volume of 200 µl. An internal standard mixture was made in 100% methanol containing: 1‐dodecanoyl‐2‐tridecanoyl‐sn‐ glycero‐3‐phosphate (PA‐12:0/13:0), 1‐dodecanoyl‐2‐tridecanoyl‐sn‐glycero‐3‐ phosphocholine (PC‐12:0/13:0), 1‐dodecanoyl‐2‐tridecanoyl‐sn‐glycero‐3‐ phosphoethanolamine (PE‐12:0/13:0), 1‐dodecanoyl‐2‐tridecanoyl‐sn‐glycero‐3‐ phosphoglycerol (PG‐12:0/13:0), 1‐dodecanoyl‐2‐tridecanoyl‐sn‐glycero‐3‐ phosphoinositol (PI‐12:0/13:0), 1‐dodecanoyl‐2‐tridecanoyl‐sn‐glycero‐3‐phosphoserine

(PS‐12:0/13:0), and 1’ –[1,2‐di‐(9Z‐tetradecenoyl)‐sn‐glycero‐3‐phospho], 3’ –[1‐(9Z‐ tetradecenoyl), 2‐(10Z‐pentadecenoyl)‐sn‐glycero‐3‐phospho]‐sn‐glycero (CL‐ (14:1) x3/15:1). Then 750 µl of methanol: chloroform (2:1, v/v) and an internal standard mixture

(for mitochondrial preparations and total hearts on normal diets, 50 ng of each phospholipid class and 100 ng of CL; for total hearts on safflower oil, 25 ng of each phospholipid class and 100 ng of CL) was added and products were extracted by the Bligh and Dyer method (226). After the samples were dried under a stream of nitrogen, they were resuspended in 100 µl of 75 % solvent A (isopropanol: hexanes 4:3, v/v) and 25% solvent B (isopropanol: hexanes: water 4:3:0.7, v/v/v, containing 5 mM ammonium acetate). Samples were analyzed by liquid chromatography coupled to tandem mass spectrometry (LC‐MS/MS) as described below.

Liquid Chromatography/Mass Spectrometry– For normal phase separation, samples were injected onto an Ascentis‐Si HPLC column (150 x 2.1 mm, 5 µm; Supelco) at a flow rate of 0.2 ml/min at 25 % solvent B. Solvent B was maintained at 25 % for 5 min, increased to 60 % over 10 min, and then to 95 % over 5 min. The system was held at 95 %

B for 20 min prior to re‐equilibration at 25% for 14 min. Phospholipids were measured

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using an API3200 triple quadrupole mass spectrometer (AB Sciex, Framingham, MA).

Positive ion mode was used to detect PC and PE lipids with quadrupole 1 scanning a m/z range from 250 to 1100 in 0.1 Da increments over 2 sec. Negative ion mode was used to detected CL, PA, PG, PI, and PS with quadrupole 1 scanning a m/z range from 150 to 1600 in 0.1 Da increments over 4 sec. Quantitation was performed using AB Sciex MultiQuant software and using PA 12:0/13:0, PC 12:0/13:0, PE 12:0/13:0, PG 12:0/13:0, PI 12:0/13:0,

PS 12:0/13:0 and CL 14:1/14:1/14:1/15:1 internal standards. All quantitated data was corrected for isotope abundance. Fragmentation of endogenous lipids m/z 818.5 (PC),

846.5, 864.5 (PC), 890.5 (PC), 885.5 (PI), and 1447.9 (CL) was performed as described above except for the following details. In the MS/MS experiment, the parent ions listed above were selected in quadrupole 1, subjected to collision‐induced decomposition using

N2 gas, and quadrupole 2 was allowed to scan the product ions in the m/z range from 150 to 900 (m/z 818.5, 846.5, 864.5, 890.5, 885.5) or 150 to 1450 (m/z 1447.9). The number of acyl carbons and double bonds present were determined using tandem mass spectrometry and identification of the fragment molecule. From these data, the other phospholipids were converted from mass‐to‐charge to number of acyl carbons and double bonds.

Preparation and Analysis of Tissue for Matrix Assisted Laser Desorption

Ionization/Imaging Mass Spectrometry (MALDI‐IMS)– A modified optimal cutting medium

(mOCT) was made by heating a 10% solution of Mowiol 6‐98 in MilliQ H2O. Once in solution, 8% Polypropylene glycol average MW 2000 was added until mixed thoroughly

(Acros Organics, New Jersey). A heart from a control animal and an ACSL1T‐/‐ animal was placed in the mOCT mixture and stored overnight at ‐20°C. A control and an ACSL1T‐/‐ heart were sectioned at ‐17°C at 20 μm, placed on glass cover slips, and stored at ‐20°C until used. DHAP (2' 5'‐dihydroxyacetophenone) matrix (150 mg) was sublimated onto the tissue, as previously described (275). MALDI‐IMS data was collected using a method

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previously described (214). In brief detail, an AB‐Sciex qTOF XL using a solid state laser

(337 nm) at and energy of 7.6 μJ and a pulse rate of 500 Hz in negative ion mode was used to capture the MALDI/IMS data. The laser was moved in a horizontal pattern in a step‐size of 50 μm. A the negative ion spectrum from m/z 600 – 1700 was recoreded as the mass spectrum at each x‐y coordinate. The data set was analyzed using TissueView software

(AB‐Sciex).

Results–

In collaboration with Dr. Rosalind Coleman’s laboratory at the University of North

Carolina, we carried out a series of experiments to determine how the inducible global deletion of ACSL1 affected phospholipid molecular species composition. The most notable of all of these, was the dramatic change of phospholipid molecular species in the heart.

Data generated from that work is presented in this chapter.

Formation of Long‐Chain Fatty Acyl CoA Esters is Diminished in the Heart when

ACSL1 is Depleted– ACSL1 was targeted using a floxed ACSL1 allele under a global promoter to drive Cre recombinase expression. After 10 weeks of tamoxifen treatment, these animals had almost no detectable ACSL activity in the heart (Figure 6‐2). Notably, the most dramatic decrease was seen in the activity for the formation of 18:2‐CoA, which is thought to be the preferred fatty acyl CoA formed by heart ACSL1. In addition to the loss of activity, these animals displayed left ventricular enlargement, loss of mitochondrial beta‐ oxidation, and increased dependence upon glucose metabolism. There were notably more mitochondria found in the heart tissue of the ACSL1T‐/‐ animals. Additional validation of

ACSL1 depletion was performed and can be found in (276).

Total Quantity of Cardiolipin Does Not Change but is Remodeled– Depletion of

ACSL1 using Cre recombinase did not lead to a change in the relative amount of each

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phospholipid class normalized to tissue protein levels. Dramatic changes were observed for the distribution of molecular species within the cardiolipin class. (Figure 6‐2).

However, additional changes were observed with several of the major glycerophospholipids. Since the mitochondria seem to have the most pronounced phenotype in the ACSL1T‐/‐ mouse, experiments initially focused on the purified mitochondria. Cardiolipins being specific to mitochondria were initially investigated since they are suggested to be critical in the proper function of the mitochondria. When cardiolipin was analyzed by LC‐MS a dramatic change in the acyl composition was observed.

45" 120" Control" 40" Control"

" 100" 35" Acsl1T'/') Acsl1T'/') " 30" 80" 25" 60" 20" /mg"protein /mg"protein/min) ACSL"Ac0vity" 15" 40" nmol

nmol 10" "( 20" 5" 0" 0" 16:0" 18:0" 18:1" 18:2" 20:4" CL" PA" PE" PS" PI" PC"

Figure 6‐2: Loss of ACSL activity in ACSL1T‐/‐ but no change in total phospholipid measurement. A) ACSL activity with 5 different long chain fatty acids were measured in control (white bars) and ACSL1T‐/‐ (black bars). B) Total phosphorous content in six major phospholipid classes. Data collected at Coleman lab (UNC) and published in Grevengoed, TJ et al. 2015.

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b A 100% 1447.9 Negative MS! 1449.9 Control ACSL1 T-/- 1451.9 c a 1453.9 d e 1475.9 1477.9 1423.9 1447.9 1455.9 1499.8 1497.8 1421.9 1425.9 1473.9 1491.9 Relative Intensity(%) 1479.9 1427.9 1523.8 1429.9 1472.0 1495.9 1522.0 1525.8

1420 1440 1460 1480 1500 1520 1540 m/z, Da BC 279.4 100% 279.2 100% MS2 MS2 18:2 FA-H 18:2 FA-H [M-H]- of 1495.9 [M-H]- of 1447.9 415.1 18:2(x2)/20:4(x2)-CL [18:2/18:2-PA- 18:2 (x4)-CL [18:2/18:2-PA- ketene-sn2]- ketene-sn2]- 415.0 695.2 18:2/18:2-PA-H 18:2/18:2-PA-H 695.3 CyclicPhosphate 743.2 433.1 432.8 20:4/20:4-PA-H Relative Intensity(%) CyclicPhosphate Relative Intensity(%) 153 18:2- 18:2- LPA-H 153 LPA-H

200 400 600 800 1000 1200 1400 200 400 600 800 1000 1200 1400 m/z, Da m/z, Da

Figure 6‐3: The remodeling of acyl chain composition of cardiolipin in ACSL1T‐/‐ heart tissue. A) The molecular ion distribution of the singly charged cardiolipin species of control (blue trace) and ACSL1T‐/‐ (red trace). B & C) Product ion spectra of m/z 1447.9 and 1495.9 from mitochondrial extracts from a control animal.

As reviewed above, the most abundant cardiolipin is a tetra‐linoleoyl

(18:2/18:2/18:2/18:2‐CL) species in the heart. The tetra‐linoleoyl molecular species was determined to be the predominant cardiolipin isoform in the control animals from the parent ion and tandem mass spectrometry data of the m/z 1447.9 molecular ion (Figure 6‐

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3A&B). In the ACSL1 depleted heart tissue, there was approximately an 80% decrease in this specific molecular species as compared to the control. The mass of total cardiolipin was accounted for an increased abundance of alternative species (Figure 6‐3A, red trace).

The dramatic shift in the molecular ions is readily apparent upon comparison of the

1447.9 ion in the control (blue trace) and ACSL1T‐/‐ (red trace) in Figure 6‐3A, bundle b.

The two cardiolipin molecular ions that experienced the most dramatic loss in the ACSL1 T‐

/‐ heart tissues were m/z 1447.9 and 1495.9. To confirm the acyl composition of these two molecular ions, tandem mass spectrometry was used. The fragmentation pattern of the m/z 1447.9 ion was characteristic of the tetra‐linoleoyl cardiolipin, producing the following anions: 18:2 carboxylate, 18:2/18:2 phosphatidic acid, and 18:2 lyso PA (Figure

6‐3B).

Similar to the fragmentation of m/z 1447.9, the fragmentation of m/z 1495.9 yielded similar anions producing an 18:2 and 20:4 carboxylate, 18:2/18:2 or 20:4/20:4 phosphatidic acid, and 18:2 or 20:4 lyso PA (Figure 6‐3C). The data supported the species observed as m/z at 1447.9 as a (18:2/18:2/18:2/18:2) cardiolipin and m/z 1496

(18:2/18:2_20:4/20:4) cardiolipin (Figure 6‐3).

Cardiolipin Content is Remodeled Throughout the Right and Left Ventricle– To determine if the changes in cardiolipin were specific to the left ventricle or specific regions of the left ventricle, the technique of MALDI imaging mass spectrometry (MALDI‐IMS) was utilized. MALDI‐IMS technique permits determination of the distribution of different lipids across a tissue and semi‐quantitative measurements can be made based on relative signal intensity. Prior to this, no work had been performed to image cardiolipin distribution in mouse heart. Using negative ion mass spectrometry, images of the three most abundant groups of molecular ions of cardiolipin (Bundles b, c, and d in Figure 6‐3A) could be detected by IMS. The groups of molecular ions from bundle b are shown in Figure 6‐4.

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Control" ACSL1"T-/-" A E

1"mm" 1"mm" B F

C G 1452"(72:6-CL)" 1450"(72:7-CL)" 1448"(72:8-CL)"

DH

RV" RV"

H"&"E" LV" LV"

Figure 6‐4: MALDI imaging of cardiolipin in the ventricles from control and ACSL1T‐/‐ mice shows global changes in acyl composition. A,B,C,E,F,G) Each image of control and ACSL1T‐/‐ mouse heart was reconstructed for a specific ion derived from the cardiolipin molecular species involved. The intensity of the ion is indicated by color using the scale bar displayed in the image. All ions were normalized to the total ion current (scale bars are displayed on right). D&H) H&E staining was performed on the same sections that were used in MALDI imaging.

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As seen in the LC‐MS data, the control animals have the highest signal intensity for the m/z ion 1447.9 (shown by the brighter color intensity) with decreasing signal intensity for m/z 1449.9 followed by 1451.9 (Figure 6‐4 A, B, and C). In the ACSL1T‐/‐ heart tissue there is a significant reduction in the signal intensity of m/z 1447.9 (compared to the intensity in the controls as well as relative to the other ions) with increasing signal intensity for m/z 1449.9 and 1451.9 (Figure 6‐4 E, F, and G). However, these signal intensity changes appear to distribute globally throughout the ventricle, and were not specific to any unique anatomical region of the heart. The observations of relative intensity made by LC‐MS data in Figure 6‐3A was recapitulated in the MALDI‐IMS data for all molecular ions that were above the instrumental noise.

Loss of DHA Containing Phosphatidylcholines in the ACSL1T‐/‐– While performing the analysis on cardiolipin, data were collected for all of the phospholipids. During the initial analysis, it was noted that there were no changes in the total content of any of the phospholipids (Figure 6‐2). However, since cardiolipin also had no change in total amount, but rather in acyl chain composition, we pursued the analysis of all of the phospholipid classes at the molecular species level. No major changes could be noted in the phosphatidylglycerol or phosphatidic acid molecular species. The remainder of the major classes of glycerophospholipids had significant changes in the acyl chain composition.

Comparison of phosphatidylcholine molecular species of the control mice compared to ACSL1T‐/‐ revealed significant changes. The control mice had two high molecular weight PC species, m/z 806 and 834 [M+H]+ which decreased significantly in the

ACSL1T‐/‐ mice, but increased in lower molecular weight species was observed (Figure 6‐

5a). Specifically, there was a gain of m/z 760.6 and m/z 788.6 [M+H]+. The initial analysis of PC and PE was performed in positive ion mode, since phosphatidylcholine species readily form positive ions. However, to gain additional structural information about the

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acyl composition, negative ion mode had to be used. Phosphatidylcholines do not readily form negative ions, therefore solvents are enriched with ammonium acetate to facilitate the formation of an adduct. The [M+OAc]‐ m/z 864 is the same phospholipid molecular species as the [M+H]+ m/z 806 and the [M+OAc]‐ m/z 892 was the same molecular species as the [M+H]+ m/z 834. Tandem mass spectrometry was performed on these two molecular ions to determine their acyl composition as described next.

Two Major Molecular Species Lost with Depletion of ACSL1 in the Heart Contained

22:6 or DHA– The positive ion m/z 806 was determined to be a 16:0/22:6‐PC (38:6‐PC) and m/z 834 to be 18:0/20:4‐PC (40:6‐PC) (Figure 5‐5B‐C). The identification of the acyl composition of 16:0/22:6‐PC (38:6‐PC), was supported by the observation of both carboxylate anions (m/z 255 (16:0) and m/z 327 (22:6)), and the choline head group was supported by the loss of methyl from the quaternary amine (m/z 790).

The observation of m/z 283.2, in the fragmentation of [M+OAc]‐ m/z 864.6, did not correspond to the presence of octadecanoate ester, but rather to the loss of CO2 from the docosahexaenoyl carboxylate anion (228). Additional structural information was gained by the loss of the sn‐2 fatty acid as a ketene [M‐FA‐ketene]‐ (m/z 480). A characteristic fragmentation mechanism of phospholipids is to lose the sn‐2 group as a ketene neutral, thus providing structural insight into the positioning of the fatty acyl chains on the phospholipid. A change of shorthand notation from unknown position of the fatty acyl groups 16:0_22:6‐PC to known position on the glycerol backbone 16:0/22:6‐PC (228). A similar identification was made for the [M‐OAc]‐ m/z 892, confirming that this PC lipid was an 18:0/22:6‐PC.

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Posi2ve'MS' 100% 806.5 Control'' A 760.6 788.6 834.5 ACSL1'T-/-'' 808.5 786.6 758.7 810.6 836.5 784.5 782.6 812.7 832.5 734.6 732.7 790.5 830.6 838.5 856.5 722.6 756.7 780.7 828.6 848.5 862.6 Relative Intensity(%)

720 740 760 780 800 820 840 860 m/z, Da B C" 100%' 327' 100%' 327' 22:6' anion' !" 22:6' anion' 255' [M+OAc]

!" 22:6-CO2' 283' [M+OAc] !" [M-CH3] 864' !" [M-22:6-ketene] 892' 283' !" 790'

818' 16:0' [M-22:6-ketene] anion'

18:0' 480' anion' 508' !"

Rela2ve'Intensity' [M-CH ]

3 Rela2ve'Intensity'

200' 400' 600' 800' 200' 400' 600' 800' D m/z,'Da' E" m/z,'Da' 100%' 100%' 281' 281' 18:1'anion' 18:1' anion' 16:0' [M+OAc]-' anion' [M+OAc]-' 255' 283' 818' 772' 744' 18:0'' 846' [M-CH3]-'

Rela2ve'Intensity' Rela2ve'Intensity'' anion' [M-CH3]-'

200' 300' 400' 500' 600' 700' 800' 200' 300' 400' 500' 600' 700' 800' m/z,'Da' m/z,'Da' 100%' 100%' 18:2' 768' 770' F G" !" anion' [M-CH ]-' [M-CH3] 16:0' 3 18:1' anion' 18:2' 255' anion' anion' 18:1' anion' 18:0' 279' 20:3' [M-OAc]-' anion'

281' anion' Rela2ve'Intensity'

Rela2ve'Intensity' 279' [M-18:2-ketene]!" 480' 842' 281' 305' 504' 283' 508' 250' 400' 600' 800' 250' 400' 550' 700' 850' m/z,'Da' m/z,'Da'

Figure 6‐5: The remodeling of acyl chain composition of phosphatidylcholine in ACSL1T‐/‐ heart tissue by LC‐MS/MS.. A) The molecular ion distribution of the positively charged PC species of control (blue trace) and ACSL1T‐/‐ (red trace). B‐G) Negative product ion spectra of m/z 818.6, 846.6, 842.6, 844.6, 892.6, and 864.6 (nominal mass are displayed).

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To determine the PC species that increased with the depletion of ACSL1, a similar approach was taken. Again, the [M+OAc]‐ ion was used for structural identification of

[M+H]+ m/z 760.6 and 788.6, which shifted to m/z 818.6 and m/z 846.6, respectively. The molecular species gained in the depletion of ACSL1 in the heart, contained a fatty acyl group of 18:1 because of the presence of the m/z 281.2 carboxylate anion. The identification of the acyl composition was supported by the observation of both carboxylate anions, and the choline head group was supported by the loss of methyl from the quaternary amine [M‐CH3]‐ after collisional activation (Figure 6‐5D,E). The positive ion m/z 760.6 was determined to be a 16:0/18:1‐PC (34:1‐PC) and m/z 788.6 to be 18:0/18:1‐

PC (36:1‐PC).

Phosphatidylcholine Containing DHA is Lost Throughout the Right and Left

Ventricle in the ACSL1T‐/‐– Similar to cardiolipin, changes in the distribution of PC were analyzed using MALDI imaging mass spectrometry. The tissue was imaged in negative ion mode. PC does not readily form negative ions therefore identification of PC anions were made by looking for adducts that formed with the matrix, 2,5‐dihydroxyacetophenone. The four most abundant PC molecular species were identified and their tissue distribution is presented as images in Figure 6‐6. As seen in the LC‐MS data, the control animals have the highest signal intensity for the [M+DHAP]‐ phosphatidylcholine ions corresponding to

38:6‐PC and 40:6‐PC with decreasing signal intensity for the ions corresponding to 36:1‐PC and 38:1‐PC. In the ACSL1T‐/‐ heart tissue, the opposite is true with the signal intensity for the ions corresponding to 36:1‐PC and 38:1‐PC being much stronger. Similar to cardiolipin, these signal intensity appear to distribute increased or decreased globally throughout the ventricle, and were not specific to any unique anatomical feature of the heart. The observations made by LC‐MS data in Figure 6‐5A was recapitulated in the MALDI‐IMS data for all molecular ions that were above the instrumental noise.

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Control( ACSL1(T%/%( 40:6%PC( 38:6%PC( 36:1%PC( 34:1%PC(

Figure 6‐6: MALDI imaging of PC in the ventricles from control and ACSL1T‐/‐ mice shows global loss of 40:6 and 38:6‐PC and gain of 36:1 and 34:1‐PC. All ions were normalized to the total ion current presented as false color (scale bars are displayed on right). Intensity of the ion is displayed using pixel intensity.

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Phosphatidylinositol, Phosphatidylethanolamine, and Phosphatidylcholine

Experienced Acyl Chain Remodeling with the Depletion of ACSL1 in the Heart– Tafazzin, the protein that has been shown to assemble cardiolipin, has been characterized as a transacylase that transfers acyl chains from PC to cardiolipin (270). There were significant changes in the [M+H]+ m/z 784 (36:3‐PC) and m/z 786 (36:2‐PC), hence, these two ions were further investigated by liquid chromatography‐tandem mass spectrometry (Fig. 6‐

7a). It was determined that the 36:3 species was a mixture of 16:0_20:3‐PC and 18:1_18:2‐

PC because of the identification of m/z 255.2 (16:0), 279.2 (18:2), 281.2 (18:1), and 305.2

(205) corresponding to each carboxylate anion. The 36:2 species was a mixture of

18:0/18:2 and 18:1/18:1 as determined by the presence of m/z 279.2 (18:2), 281.2 (18:1), and 283.2 (18:0). No major shift in the ratio of any of these acyl compositions was observed in the two molecular species that make up each of these molecular anions in the

ACSL1T‐/‐ heart tissue. Since cardiolipin is believed to gain acyl chains from PC it would be feasible that changes in PC composition could change cardiolipin. There is an increase in the presence of 18:1 over 22:6, but also an increase in 18:2 containing PC’s, which would not account for loss of tetra‐linoleoyl cardiolipin.

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12" A * * * 10" * * * 8"

6" * * * * 4" * * * * * * *

Isotope'Corrected'Area'Ra/o' 2"

0" 32:1" 32:0" 34:3" 34:2" 34:1" 36:4" 36:3" 36:2" 36:1" 38:6" 38:5" 38:4" 38:3" 38:2" 40:9" 40:8" 40:7" 40:6" 40:5" 40:4" 42:6"

25" B * 20"

15" ** **** * 10" ** ** * 5" Isotope'Corrected'Area'Ra/o' * **

0" 34:2" 34:1" 36:5" 36:4" 36:3" 36:2" 36:1" 36:0" 38:6" 38:5" 38:4" 38:3" 38:1" 38:0" 40:8" 40:7" 40:6" 40:5" 42:10" 42:9" 42:8"

50" C * 40"

30"

20"

10" * * * * Isotope'Corrected'Area'Ra/o' * ** * * ** *

0" 34:2" 34:1" 36:4" 36:3" 36:2" 36:1" 38:6" 38:5" 38:4" 38:3" 40:7" 40:6" 40:5" 40:4"

Figure 6‐7: Semi‐quantitative changes in acyl composition of PC, PI, and PE in the heart tissue of the ACSL1T‐/‐ compared to control animals reveals major changes in acyl chain composition by LC‐MS. Changes in the molecular ions of A) PC, B) PE, and C) PI. The x –axis is labeled with the number of carbons: number of double bonds based on the head group assignment by chromatography. The y‐axis was determined by using correction factors made for isotopic distribution and corrected values were the normalized to an internal standard. N=3, Average ± SEM, Students T‐test, *= p<0.05

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Of the 21 PC molecular ions that were reproducibly detectable, 18 of them had significant changes in the ACSL1T‐/‐ mouse heart when compared to the control mice heart tissue (Figure 6‐7A). The most notable were the PC species that lost DHA (40:6 and 38:6) in the ACSL1T‐/‐ heart tissue and gained shorter, less unsaturated acyl compositions (34:1,

36:3, 36:2, and 36:1). The shift in acyl composition did not correlate well with the known activity of ACSL1, which was believed to preferentially catalyze the covalent linkage of coenzyme A with 18 carbon fatty acyl esters (277). Interestingly, there was a trend towards an increase in species that contained 18 carbon fatty acyl chains in the ACSL1T‐/‐ revealing that substrate availability to the enzymes may be more important than the preferred fatty acid preference of the isolated enzyme.

Similar trends were observed with the PE molecular species. Significant changes were observed with 16 of the 21 reproducibly detected PE molecular species (Figure 6‐

7B). Of these, the most dramatic losses in the ACSL1T‐/‐ were those of the DHA containing

PE species (38:6 and 40:6). Since there was no loss in the total quantity of PE, the loss of these two molecular species was accounted for by a gain in numerous PE species with mostly containing a total of 34 and 36 carbon and less than two double bonds. This observation again was not consistent with what would have been expected from the known activity preferences of ACSL1. One notable change that was not observed with the

PC molecular species was an increase in the 38:4 PE molecular species. This phospholipid was determined to be an 18:0/20:4‐PE by tandem mass spectrometric analysis. The

18:0/20:4‐PC but had no significant change. The data in Figure 6‐7 supported exchange between PC and PE, since many of the changes were similar, but also supported distinct regulation of the two phospholipid classes.

In most mammalian systems 38:4‐PI or 18:0/20:4‐PI is the overwhelming PI molecular species. However, in the ACSL1T‐/‐ mouse, there was approximately a 50%

152

reduction of this molecular ion, observed as m/z 885 in negative ion mode (Figure 6‐7C).

The m/z 885 molecular ion was confirmed to be the 18:0/20:4‐PI species by tandem mass spectrometry. Similar to the other glycerophospholipids, there was no reduction in the total mass of PI, therefore the other molecular species of PI increased to compensate for this loss of the 38:4‐PI. Of the PI species that could be readily detected, 13 of the 14 species had significant changes compared to the control and compensated to render the total PI mass unchanged.

Cardiolipin Content is Recovered with High Levels of Linoleic but No Recovery of

Mitochondrial Function– With the finding of major changes in cardiolipin molecular species the Coleman lab used dietary changes to try to regain the molecule species profile.

Previous work that was done in collaboration with our laboratory showed that rats that spontaneously develop heart failure have a similar loss of tetra‐linoleoyl cardiolipin. When these rats with heart failure were fed a high safflower oil diet it prolonged their life span and improved heart function (273). Safflower oil is 70‐80% esterified linoleic acid (18:2).

Those rats provided with this diet also had a reversal of the loss of tetra‐linoleoyl cardiolipin. After the depletion of ACSL1 for 10 weeks, the high safflower oil diet was fed to the ACSL1T‐/‐ mice for 4 weeks to determine if the remodeling of cardiolipin could be reversed and ultimately reverse the respiratory impairment of the heart.

Cardiolipin was again measured in control and ACSL1T‐/‐ animals, both of which were fed a safflower oil diet. These data were compared to the animals that were maintained on a normal chow diet. After four weeks on the safflower oil diet, there was a significant recovery of the tetra‐linoleoyl cardiolipin in the ACSL1T‐/‐ mice as compared to the mice on normal chow (Figure 6‐8A). Data from multiple animals were quantitated, and showed that the ACSL1T‐/‐ hearts from mice fed a high safflower oil diet had levels of tetra‐ linoleoyl comparable to the control animals on normal chow (Figure 6‐8B). The recovery of

153

tetra‐linoleoyl cardiolipin was not sufficient to regain normal mitochondrial function

(Figure 6‐12 or reference (276)).

Similar to the measurement of cardiolipin, the glycerophospholipids molecular species were analyzed for changes after mice were given a safflower oil diet (SO). The loss of DHA was not recovered with the diet shown by the graphs in Figure 6‐9 graphs labeled

40:6‐PC, 40:6‐PE, 38:6‐PC and 38:6‐PE. That the SO diet did not reverse the loss of DHA containing phospholipids was an expected finding since safflower oil is high in the fatty acid 18:2 [ω‐6] which cannot be converted to 22:6 [ω‐3] (DHA) in mammals. However, the

SO diet should be able to increase the 20:4, ω‐6 (AA) content, thus potentially increasing the 38:4 PI and 38:4 PE. When the PI species were compared to the normal chow animals

(NC), the ACSL1T‐/‐ showed no recovery of the 38:4 molecular species, and if anything there was a trend towards a decrease (Figure 6‐9). However, when it came to the 18:0/20:4‐PE there was an increase in both animals. The control animals showed a slight increase in the

38:4 PI species with the safflower oil diet (SO compared to NC). The data supports the concept that ACSL1 may be critical in the formation of 18:0/20:4‐PI, at least in the heart, but addition investigation is needed.

As for the molecular ions of specific phospholipids that increased with the depletion of ACSL1, these ions were either unchanged or increased with the safflower diet

(SO). Two examples, 34:1 and 36:1‐PC are shown in Figure 6‐9. It is difficult to gather from these data sets if any of the phospholipid changes are causing an effect, but it is interesting to note that even with almost complete depletion of ACSL activity in the heart, there is still incorporation of acyl chains from the diet being incorporated into the phospholipids in the heart.

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72:7 A 72:8 B 100% 10 72:6 Negative MS' Normal'Chow'ACSL1'T0/0' High'Safflower'Oil'ACSL1'T0/0' 8

72:5 74:8 6 74:9 74:7 76:10 76:11 4 Relative Intensity 74:10 76:9

70:5 72:4 74:6 Isotope'Corrected'Area'Ra?o' 70:6 78:12 76:8 70:4 78:13 78:11 2 78:10 0 1420 1440 1460 1480 1500 1520 1540 NC SO NC SO m/z, Da Control' ACSL1'T0/0'

Figure 6‐8: Cardiolipin shows significant changes in molecular species with a high safflower oil. A) The molecular ion distribution of the singly charged cardiolipin species of ACSL1T‐/‐ mice on normal chow (blue) and ACSL1T‐/‐ mice on safflower oil [SO]. B) Quantitation of the molecular ion m/z 1447.9 (tetra‐linoleoyl cardiolipin) the control (black bars) and ACSL1T‐/‐ (white bars) on normal chow (NC) and safflower oil (SO).

Bis (Monoacylglycerol) Phosphate is Increased in the ACSL1T‐/‐– After a global analysis of the most abundant glycerophospholipids, a more targeted approach was looking at a minor phospholipid component, bis (monoacylglycerol) phosphate (BMP).

These are a unique group of phospholipids that are thought to be localized to the late endosomes and lysosome (278). Their presence in this compartment is critical as the inside of the lysosome/endosome is filed with lipases that cleave normal glycerophospholipids. Their synthesis is thought to stem from transacylation from other phospholipids, predominately PI (279). BMPs have also been shown to be precursors for the release of AA in macrophages (280).

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40:6'PC' 38:6'PC' 25 25

20 20

15 15

10 10

5 5 Isotope'Corrected'Area'Ra/o' Isotope'Corrected'Area'Ra/o' 0 0 NC SO NC SO NC SO NC SO Control' ACSL1'T6/6' Control' ACSL1'T6/6'

40 34:1'PC' 20 36:1'PC'

30 15

20 10

10 5 Isotope'Corrected'Area'Ra/o' Isotope'Corrected'Area'Ra/o' 0 0 NC SO NC SO NC SO NC SO Control' ACSL1'T6/6' Control' ACSL1'T6/6'

8 38:6'PE' 20 40:6'PE'

6 15

4 10

2 5 Isotope'Corrected'Area'Ra/o' Isotope'Corrected'Area'Ra/o' 0 0 NC SO NC SO NC SO NC SO

Control' ACSL1'T6/6' Control' ACSL1'T6/6'

12 38:4'PE' 20 38:4'PI'

9 15

6 10

3 5 Isotope'Corrected'Area'Ra/o'

Isotope'Corrected'Area'Ra/o' 0 0 NC SO NC SO NC SO NC SO

Control' ACSL1'T6/6' Control' ACSL1'T6/6'

Figure 6‐9: Safflower oil does not repair most of the changes made to the phospholipid molecular species in the ACSL1T‐/‐. Control and ACSL1T‐/‐mice were fed normal chow (nc) or safflower oil diet (SO), and phospholipid content was measured (n=3, average ± SEM).

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BMP"in"Purified"Mitochondrion" Control" 0.30 *" ACSL1"T./." 0.25 0.20 *" 0.15 *" *" *" *" *" *" Area"Ra

0.05

0.00 769$ 771$ 773$ 793$ 795$ 797$ 817$ 819$ 821$ 841$ 843$ 845$ 865$ 867$ 869$ Figure 6‐10: Many BMP molecular ions are increased in the heart tissue of ACSL1T‐/‐. Control (white bars) and ACSL1T‐/‐ (black bars) were analyzed for changes in BMP molecular species. Molecular ion (m/z) is displayed on the x‐axis. Peak area was compared to a PG standard to determine the values displayed on the y‐axis (n=5, average ± SEM, Students t‐test *=p value <0.05).

Work done in the Coleman laboratory suggested that there was a higher level of autophagy occurring in the heart and liver tissue of these ACSL1T‐/‐ mice. Numerous species of BMPs were measured, and it was found that in the heart, 8 of 15 BMP species were increased (Figure 6‐10) and 2 of 8 that were measured in the liver increased significantly

(Figure 6‐11). The increase in BMP level would support the increase in autophagy since the autophagic vesicle fuses with the lysosomes.

Unlike the heart, the liver specific ACSL1 depleted animal experienced no significant changes in acyl composition of any of the major glycerophospholipid classes

(Figure 6‐11). The only changes observed in the liver of the ACSL1L‐/‐ were the 18:1/18:1 and 18:2/18:2 BMP which were both significantly increased. The corresponding PG molecular species were unchanged (Figure 6‐11). Initially normal phase liquid chromatography was used to separate the phospholipid molecular species then a precursor ion scan was performed to identify molecular ions that contained an 18:1 fatty acyl chain(s). Since this targeted analysis was performed there may be additional changes in the BMP molecular species that were not detected by this type of scanning method.

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Discussion–

The ACSL family is critical in the formation of long‐chain fatty acyl CoA esters.

Although most tissues contain several of the 13 enzymes that encode this function, the murine heart almost exclusively uses ACSL1 for this purpose, and in other mammalian hearts this is the predominant isoform (281). The role of ACSL1 in β‐oxidation in the cardiac tissue had previously been characterized, but its the effect on the phospholipid molecular species had never been measured. The work that was performed in this chapter shed light into the critical role that ACSL1 has in cardiac phospholipid acyl chain composition.

One of the most dramatic changes observed between the control and ACSL1T‐/‐ mouse was the phospholipid acyl composition of cardiolipin in cardiac tissue. Cardiolipin is thought to be critical for mitochondrial function. The importance of the acyl composition of cardiolipin has been controversial because changes in acyl composition do not always impact function of the mitochondria. Some acyl chain compositions of cardiolipin do appear to be necessary for proper function. However, the work performed here would suggest that the acyl chain composition of cardiolipin is not a major factor in left ventricular enlargement or mitochondrial dysfunction when ACSL1 is depleted. The interpretation is supported by the fact that cardiolipin molecular species are restored to predominately tetra‐linoleoyl cardiolipin by the safflower oil diet (Figure 6‐8), but there are no significant changes in basal respiration or mitochondrial function (Figure 6‐12). The diet was provided to the animals after the damage to the heart had already occurred. It is possible that the safflower oil diet could prevent the onset of left ventricular enlargement and mitochondrial dysfunction if provided in parallel with the tamoxifen treatment.

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70"

60"

50"

40" ACSL1""Control" ACSL1"C/C" 30"

Percent"of"Total" 20"

10" *" *"

0" BMP"like"1" BMP"like"3" BMP"like"5" BMP"like"2" BMP"like"4" 18:1/20:4"PI" 18:0/18:1"PS" 18:1/20:4"PS" 18:1/22:6"PS" 16:0/18:1"PE" 18:1/18:2"PE" 18:1/18:1"PE" 18:0/18:1"PE" 18:1/20:4"PE" 18:1/22:6"PE" 16:1/18:1"PA" 16:0/18:1"PA" 18:1/18:2"PA" 18:1/18:1"PA" 18:0/18:1"PA" 18:1/20:4"PA" 18:1/22:6"PA" 16:1/18:1"PC" 16:0/18:1"PC" 18:1/18:2"PC" 18:1/18:1"PC" 18:0/18:1"PC" 18:1/20:4"PC" 18:1/22:6"PC" 16:0/18:1"PG" 18:1/18:2"BMP" 18:1/18:1"BMP" 18:0/18:1"BMP" 18:1/20:5"BMP" 18:1/20:4"BMP" 18:1/20:3"BMP" 18:1/22:6"BMP" 18:1/22:5"BMP"

100% 9.11

PG" Control" ACSL1"C/C" 1345" 1346" 1347" 1348" 1349" 1350"

BMP" 6.26 Relative Intensity NL" PC" CL" PE/PI" 4.31 18.46 29.67 12.26 PA/PS"

2 4 6 8 10 12 14 16 18 20 22 24 26 28 30 32 Time, min

18:1/18:1"BMP" 18:1/18:2"BMP"

Control" Control" ACSL1"C/C" 6.40 ACSL1"C/C" 100% 6.40 100% 1345" 1346" BMP 1345" 1346" BMP 1347" 1348" 1347" 1348" 1350" 1349" 1350" 1349" Relative Intensity Relative Intensity 9.28 9.28 PG PG

5.0 7.0 9.0 11.0 13.0 5.0 7.0 9.0 11.0 13.0 Time, min Time, min Figure 6‐11: BMP molecular ions are also increased in the livers of ACSL1L‐/‐. Control (blue bars) and ACSL1L‐/‐ (red bars) were analyzed for changes in BMP molecular species. Molecular species are displayed on the x‐axis and were determined by precursor ion scanning. Total peak area was compared each individual species to determine the values displayed on the y‐axis (n=3, average ± SEM, Students t‐test p value <0.05). Total chromatographic traces of 18:1 containing phospholipids reveal no changes in molecular species. The 18:1/18:1 and 18:1/18:2 BMP change, but not in the corresponding PG molecular species show no changes. Three independent animals were tested. Individual traces for Control animals (1345,1347, 1349) and ACSL1L‐/‐ (1346, 1348, 1350) are shown in the chromatographic traces.

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A B Basal"RespiraAon" Mitochondrial"FuncAon" 100" 4.0" Low"Fat"Control" Control Low"Fat"Acsl1T'/') " 3.5" T'/' Safflower"Oil"Control" 80" Acsl1 3.0" Safflower"Oil"Acsl1T'/')

60" 2.5" 2.0" /min/µg"protein) 40" 1.5" pmol

OCR"RelaAve"to"Basal" 1.0" 20" OCR"( 0.5"

0" 0" Low"fat" Safflower"Oil" ADP" Oligomycin" FCCP" AnAmycin"A"

Figure 6‐12: High linoleate diet did not improve mitochondrial function in Acsl1T‐/‐ hearts. For 16 weeks after tamoxifen injections, male mice were fed a low‐fat (10% fat) control diet. Mice were then either maintained on the low‐fat diet or switched to a high safflower oil diet (Research Diets, D02062104, 45% kcal fat (75% linoleic acid)) for 4 additional weeks to increase dietary linoleate. A&B) Mitochondrial function was measured in isolated mitochondria using a Seahorse XF24 Analyzer (n=4‐7). (O2 consumption rate: OCR). * p‐ value ≤0.05 between genotypes within diet (276).

The acyl composition of cardiolipin was not the only glycerophospholipid to have dramatic acyl chain remodeling. Phospholipid molecular species in cells are strictly regulated, and often do not change despite loss of proteins essential in the production of that specific phospholipid molecular species. The ability to maintain these specific species is due to the redundant pathways that exist for the synthesis of critical cellular phospholipids. Many of the molecular species are thought to be critical for membrane fluidity, membrane protein function, and protein docking, as well as being precursors of bioactive lipids. The phospholipids (PC, PE, PS, and PI) also had significant changes in their acyl chain composition.

In both PC and PE, there was a significant decrease in the incorporation of DHA into the phospholipid molecular species. In addition to these, there was a dramatic decrease in the AA contain PI. Both DHA and AA can be synthesized from dietary precursors linoleic or linoleic acid, respectively. The conversion of both fatty acids requires a delta‐6 desaturase.

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Previous work in rats has shown that inhibition of delta‐6 desaturase led to improved outcomes after mechanical injury to the heart, thus suggesting that the more polyunsaturated fatty acyl chains (PUFAs) in phospholipids the worse the damage to the heart post‐injury (282). However, supplementation of fish oil attenuates cardiac damage in models of left ventricular hypertrophy and pressure overload (283,284). In the ACSL1T‐/‐ mice, there was a significant loss of PUFAs, which may have added to the enlargement of the left ventricle. ACSL1T‐/‐mice provided with a high safflower oil diet did not have recovery of most of the phospholipid molecular species. Previous work had shown that high fat diets block the effects of a diet rich in DHA in heart failure (285). Possible explanations are that species differences (between mice and rats) have a different demand for acyl composition or that mechanical injury has a much different pathophysiology compared to the left ventricular remodeling that occurs in the ACSL1T‐/‐animal. However, from the data presented here, and previously published data, the acyl composition of phospholipids in the heart is important for proper function.

A major caveat to this work is that ACSL1 is expressed in many tissues and is potentially depleted throughout the animal (281). One major question is: Does lipid synthesis and modification in the liver in turn affect the phospholipid molecular species in the heart? Previous work performed with a liver‐specific ACSL1L‐/‐ mouse showed that there was about a 50% decrease in the expression of ACSL1 and a 35% decrease in ACSL activity (286). The decrease in ACSL1 in the liver led to decreased triglyceride synthesis, beta‐oxidation, and a shift to 16:0 and 18:1 containing PC and PE. The shift to more 16:0 and 18:1 containing phospholipids was a similar finding in the hearts of the global ACSL1T‐

/‐animal. However because of the global deletion of ACSL1, it is unknown if the phospholipid remodeling in the heart is a result of phospholipid synthesis in the liver or the heart.

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In has been previously established that there are very high levels of diverse lysophospholipid and phospholipid molecular species along with the many fatty acids and neutral lipids found in lipidomic studies of human plasma (239). They suggest that there are approximately 300 nmol/ml of DHA containing PC and PE molecular species in plasma, and approximately 5 nmol/ml of DHA containing lyso‐PC and –PE (239). There is also approximately 11 nmol/ml of the 38:4‐PI found in plasma (239). Considering the large concentration of DHA and AA contain phospholipids, potentially global deletion of ACSL1 leads to changes in the levels of these phospholipids in the liver ultimately affecting the phospholipid species in plasma. Further work to determine the origin of the phospholipid defect would have to be performed to determine if the defect in phospholipid synthesis is taking place in the heart or the liver.

From data previously described in the liver, ACSL1 seems to be critical for β‐ oxidation of fatty acids (286). One of the increases observed was in the 18:2 acyl compositions of PC, PE, and PI. If ACSL1 were important in the elongation and desaturation of linoleic acid to arachidonic acid, there could be selective loss of lipids that contain arachidonoyl chains. Similarly, if ACSL1 were critical for the elongation of linolenic acid there could be alterations in acyl composition away from docosahexaenoyl acyl chains.

Thus, ACSL1 may not be critical for the fatty acyl‐CoA ester formation necessary for lysophospholipid acyltransferase, but rather is critical as a fatty acyl‐CoA ester for elongation and desaturation.

Another possible explanation for left ventricular enlargement could be explained by changes in the molecular species of PI. PI and its phosphorylated forms are critical in numerous cellular functions including vesicle fusion, protein docking, signaling cascades, and as precursors for bioactive lipids (208,287,288). Many kinases that phosphorylate the inositol head group, to form critical phosphoinositides, prefer to associate with the

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18:0/20:4‐PI molecular species (208). MBOAT7/LPIAT1, which was the focus of the previous chapter, preferentially assembles 18:0/20:4‐PI (191,192). Therefore loss of this specific acyl composition could severely compromise cellular processes that are dependent on certain phosphorylation states of PI. Phosphoinositides are fundamental in the calcium signaling cascades required for proper cellular signaling that control heart beating, but their role in heart failure is complicated and dependent on the pathway they affect (289).

Therefore, it would be difficult to pinpoint a single signaling pathway that is affected in the

ACSL1T‐/‐ hearts if it was having an effect on phosphoinositides.

The heart has a dependence on functional mitochondria. The dependence on functional mitochondria makes the process of mitophagy, autophagy or recycling of damaged mitochondria, a critical process that occurs in myocytes (290). During the process of mitophagy, defective mitochondria are sequestered into the autophagosome, which later fuses to the lysosome to degrade and recycle the contents in the cell. The BMPs, a minor group of phospholipid, were found and later discovered to have a critical role in lining the inner membrane of the lysosome. Increases in BMP molecular species were observed in the heart and liver of the ACSL1T‐/‐ and ACSL1L‐/‐ mice. In addition PIP‐3 is critical in the formation of the autophagosome and promotes macroautophagy (291). Since there was a significant decrease in the PI species that get phosphorylated, this supported another possible explanation for a build‐up of defective mitochondria in the ACSL1T‐/‐ heart tissue. The data presented in this chapter would support a possible explanation for alterations in autophagy and mitophagy in these mice, but additional experiments would need to be performed.

Taken together, the work presented in this chapter showed major changes in acyl composition of many glycerophospholipids in the hearts of mice with global tamoxifen inducible ACSL1T‐/‐. The most noteworthy of these included the loss of DHA containing PC

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and PE (with a dramatic shift to palmitate and oleate containing species), loss of

18:0/20:4‐PI, and loss of tetra‐linoleoyl cardiolipin. Although the acyl chain composition of cardiolipin was restored when ACSL1T‐/‐ animals were provided a safflower oil diet, there was no correction to the impairments observed in the heart nor were there changes in the phospholipid compositions that were described here. Although ACSL1 is important in assembling proper cardiolipin, it does not appear to be the reason that these animals are undergoing left ventricular enlargement. Additional studies to understand the role of other phospholipids may be critical to further understand how ACSL1 is leading to mitochondrial dysfunction in these mice.

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CHAPTER VII

CONCLUSIONS AND FUTURE DIRECTIONS

Summary of Major Findings–

Chapter III– Previous lysophospholipid acyltransferase assays used one lysophospholipid and one fatty acyl CoA ester to determine the substrate preference and activity of LPATs. The previous LPAT assays used less sensitive methods to detect the formation of products from the reaction, which required high concentration of substrates.,

A novel acyltransferase activity assay (the dual choice assay) was described in Chapter 3 that took advantage of the increased sensitivity and specificity of LC‐MS/MS to detect product formation. Prepared cellular microsomes were incubated with 6 non‐endogenous lysophospholipids and 8 fatty‐acyl CoA esters with the potential of producing 48 novel phospholipid products. Products of the reaction were identified by the chromatographic profile and a precursor‐product relationship. This dual choice assay had comparable results to the more traditional assays, and using only a single assay it was able to give more insight into the complex mixture of LPATs found on the microsomes.

Chapter IV– The activity of lysophosphatidylcholine acyltransferase 3 (LPCAT3) was previously shown to preferentially incorporate AA‐CoA into lyso PC, PE, and PS. The substrate preference of this enzyme suggested its importance in regulation of arachidonoyl levels in phospholipids. Previous experiments pretreating neutrophils with a mercury containing small molecule (thimerosal) led to robust increases in the formation of leukotrienes (236). To further characterize the role of LPCAT3, RAW 264.7 cells were used in a series of experiments. Treatment of the RAW 264.7 cells with thimerosal, followed by stimulation with ATP, did not have an affect on leukotriene production. Increased expression of MBOAT5/LPCAT3nwith a PPARγ agonist (pioglitazone) elicited a dose dependent decrease in the formation of leukotrienes after ATP stimulation. Targeted

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decrease of MBOAT5/LPCAT3 with shRNAs led to significant reduction in the activity of the enzyme compared to the non‐targeted shRNA control cells. These changes confirmed that the dual choice activity assay, described in Chapter 3, was sensitive enough to detect changes in a single acyltransferase within a complex mixture of LPATs. However, when the

MBOAT5/LPCAT3shRNA cells were challenged with ATP, there was no significant change in the production of leukotrienes compared to cells that had the non‐targeted shRNA.

Chapter V– Fatty acyl CoA esters are essential for beta‐oxidation, elongation/desaturation, and acylation reactions by acyltransferases. During the course of the thesis work, we became interested in the role of acyl‐CoA synthetase long‐chains in the reincorporation of AA in bone marrow derived cells. ACSL4 is embryonic lethal, and no known bone marrow specific deletion has been created to our knowledge, but Dr. Rosalind

Coleman had created an ACSL1 global tamoxifen inducible mouse and Dr. Karin Bornfeldt had created a bone marrow specific deletion of ACSL1. Experiments are currently on going to determine the role of this protein in the bone marrow. However, in collaboration with

Dr. Coleman, an essential role for ACSL1 in the regulation of phospholipid molecular species was found in the heart. There were no changes in the total amount of any phospholipid, but rather there was acyl chain remodeling in cardiolipin, phosphatidylcholine, phosphatidylethanolamine, phosphatidylinositol and phosphatidylserine. Previous studies showed impaired mitochondrial function in the cardiac tissue, which matched the phospholipid data in which cells had a significant loss of the tetra‐linoleoyl cardiolipin. Treatment of ACSL1T‐/‐ animals with a high safflower oil diet mostly recovered cardiolipin molecular species, but did not recover mitochondrial function, left ventricle enlargement, or other remodeled phospholipids.

Chapter VI– The formation of phosphatidylinositol lipids is essential for normal cellular function, specifically, PI species that are enriched with the acyl chain combination

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of 18:0/20:4. Several protein domains are dedicated to the recognition of the phosphorylated forms of the 18:0/20:4‐PI species (208). Lysophosphatidylinositol acyltransferase 1 (LPIAT1) was identified to preferentially assemble the 18:0/20:4‐PI molecular species of PI (191,192). To determine the role of MBOAT7/LPIAT1 in the production of leukotrienes in the RAW 264.7 cell, an shRNA approach was used to selectively decrease the expression at the mRNA level. The selective decrease of

MBOAT7/LPIAT1 led to significantly decreased enzymatic activity for the incorporation of

AA‐CoA into lysophosphatidylinositol. Measurement of the global phospholipids in the

MBOAT7/LPIAT1 targeted cells did not reveal significant changes in the phosphatidylinositol species compared to the non‐targeted shRNA control cells. However, when the cells were stimulated to produce eicosanoids, the MBOAT7/LPIAT1 targeted cells had a significant decrease in the formation of leukotrienes compared to the non‐target shRNA cells. Add back of exogenous AA was not able to recover the decrease in leukotriene production. The mechanism by which MBOAT7/LPIAT1 leads to the decrease in leukotriene formation is currently being addressed by experiments laid out in the following section.

Future Directions–

The Complex Role of Thimerosal– The data presented in this thesis both supports and refutes the previous experiments surrounding LPCAT3’s role in the reacylation of AA.

The experiments that were previously performed focused on pretreatment of neutrophils with thimerosal to inhibit MBOAT5/LPCAT3 activity. Thimerosal can covalent bind free sulfur groups found within proteins (that have solvent accessible and unbound cysteines) and small molecules throughout the cell. Most of the enzymes critical in the reacylation pathway utilize cysteine chemistry in their active site; therefore, they would be inhibited

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by thimerosal. Since all of the enzymes in the reacylation pathway could be inhibited at once, AA would not be reacylated and would accumulate. This inhibition would leave AA available for oxidation by enzymes such as 5‐LO and COX1/2, and would lead to an increase in the production of eicosanoids.

To further understand why the findings in the neutrophil differ from the RAW

264.7 cell, experiments could be performed to test how thimerosal differentially affects the activity of acyltransferases and acyl‐CoA synthetases between the two cell types. The phenotype in the RAW 264.7 cell compared to the human neutrophil may be due to how the sensitivity to thimerosal differs between the mouse and human enzymes. If the mouse isoforms were not as sensitive to inhibition by thimerosal, then this would explain why treatment of the cells with thimerosal did not increase eicosanoids in the RAW 264.7 cell.

To test the sensitivity of the different acyltransferase or acyl‐CoA synthetase enzymes to thimerosal, a series of enzymatic assays could be performed. For example, a dual choice assay for acyltransferase activity and an acyl CoA synthetase activity assay could be performed with and without thimerosal with human neutrophil and RAW 264.7 cell microsomes. The differences in activity could potentially shed light on the critical regulatory enzyme activities essential for the incorporation of AA in each cell type. This data may explain why thimerosal did not inhibit the reincorporation of free AA or lead to increases in the production of leukotrienes in the RAW 264.7 cell.

The Regulation of MBOAT5/LPCAT3 by RXR and the LPCAT3T‐/‐ Mouse– Since pioglitazone treatment increased MBOAT5/LPCAT3 expression and decreased leukotriene production in the RAW 264.7 cell, additional work needs to be performed to determine how leukotriene production is regulated by LPCAT3. To continue our studies on the role of

MBOAT5/LPCAT3 in monocytes, a collaboration has been established with Dr. Peter

Tontonoz’s laboratory at UCLA. The focus of their laboratory has been on RXR

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transcriptional regulation, which has in turn led them to pursue experiments with LPCAT3.

Their laboratory has engineered a mouse with a global inducible Cre recombinase system to decrease MBOAT5/LPCAT3 expression. They are currently isolating bone‐marrow monocytes and treating them with tamoxifen to induce the MBOAT5/LPCAT3 depletion.

The goal of the study is to determine how both acute and chronic depletion of

MBOAT5/LPCAT3 in harvested monocytes affects eicosanoid production. This model of isolating the cells from animals and keeping them briefly in culture to induce the

MBOAT5/LPCAT3 depletion is more comparable to the human neutrophils that are isolated from the blood and stimulated within a few hours of removal compared to the

RAW 264.7 cells that have been kept in culture.

Previous studies performed with liver specific depletion of MBOAT5/LPCAT3 led to a decrease in the AA level esterified in PC (292). If deletion of MBOAT5/LPCAT3 in monocytes also leads to a decrease in the levels of AA esterified in PC, there would be less

AA containing phospholipid substrate for cPLA2α to hydrolyze. Decreased AA release should lead to a decrease in the formation of eicosanoids because there is less substrate available to the enzymes.

Alternatively, decreases in MBOAT5/LPCAT3 in monocytes could lead to an increase in the formation of eicosanoids. MBOAT5/LPCAT3 has been shown to be a critical enzyme in the esterification of AA into PC, PE, and PS. Once cPLA2α has hydrolyzed AA from phospholipids, the AA can be oxidized or recycled via the formation of a fatty acyl ester by acyl‐CoA synthetase long chains followed by esterification by an acyltransferase. Inhibiting reacylation of AA‐CoA can revert the ester back to the free fatty acid form through hydrolysis, therefore increasing the amount of free AA in the cell. This free AA can, in turn, be utilized by 5‐LO and COX1/2. Thereby, decreases in MBOAT5/LPCAT3 in monocytes would lead to an increase in eicosanoids.

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ACSL4 in the Regulation of AA in the RAW 264.7 Cell– Previously published data has suggested ACSL4 as a critical regulator of eicosanoids production, a shuttle of AA‐CoA into PI, and partitioning of fatty acids into diacylglycerol and triacylglycerol (246,293,294).

With all of these suggested roles, experiments were performed to determine how targeted decreases of ACSL4 affected the production of eicosanoids in RAW 264.7 cells. From the data presented in Chapter 6, there was a significant increase in the amount of free AA, but this did not lead to an increased production of leukotrienes. Previously published studies suggest that ACSL4 is located in the mitochondrial‐associated membrane in liver (258).

The enzymes to produce leukotrienes localize to the nuclear envelope. Therefore, build‐up of AA, because of the decrease in ACSL4, would be in the wrong cellular location for the production of leukotrienes.

The human neutrophil incorporates much of the free AA into diacylglycerol and triacylglycerol. If thimerosal inhibited the esterification of AA to CoA through covalently binding to ACSL4, there would be a significant increase in the production of leukotrienes.

Since the RAW 264.7 recycles free AA into phospholipids, potentially another ACSL isoform is critical the esterification of AA to CoA in the RAW 264.7 cell. Preliminary experiments with shRNAs targeted against ACSL1 were performed that showed a decrease in the production of leukotrienes, however those studies have not been followed up on. The expression of the other ACSL isoforms (2, 3, and 6) has not been tested in the RAW 264.7 cell. However, one of these three enzymes may be critical in controlling the availability of

AA to the 5‐LO/FLAP complex.

MBOAT7/LPIAT1 in the Regulation of AA Contain Phosphatidylinositols in the

RAW 264.7 Cell– Experiments are currently ongoing to determine why depletion of

MBOAT7/LPIAT1 in these cells leads to a decrease in leukotriene formation. A current hypothesis is that a specific pool of AA‐PI that contributes AA for leukotriene formation has

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decreased. However, the global measurements of PI do not change because levels of AA containing PI in membranes that do not contribute to the production of eicosanoids stay the same. To determine if there are specific changes in phospholipid molecular species, subcellular fraction by differential centrifugation is being carried out as previously performed in the RAW 264.7 cell (254). Data from this published paper suggested that PI at the nucleus was a very small percentage of the total PI in the cell. Therefore, small changes of the PI pool at the nuclear envelope could lead to dramatic changes in eicosanoid production while undetectable by measuring global phospholipids. Also, their data supported the role of PI in cellular stimulation. When RAW 264.7 cells were stimulated with the TLR4 agonist Kdo2‐lipid A for 24 hours there were decreases in the 18:0/20:4‐PI molecular species at the ER and the PM (254). It is difficult to draw comparisons since the stimulation of ATP is only 1‐60 minutes verse 24 hours, however their findings that stimulating the RAW 264.7 cells with a TLR agonist leads to decreases in an AA containing

PI. This data supported the role for PI in the activation of the RAW 264.7 cell. The goal of this study is to detect changes in the nuclear pool of AA containing PI to support the hypothesis that decreases in 18:0/20:4‐PI specific lipid will lead to changes in leukotriene production.

MBOAT7/LPIAT1in the Regulation of Mead Acid in the RAW 264.7 Cell– An observation was made during the global phospholipid analysis of the RAW 264.7 cell that the major phosphatidylinositol was not m/z 885.6, which corresponds to a 38:4

(18:0/20:4‐PI). Rather m/z 887.6, which corresponds to a 38:3 (18:0/20:3‐PI), was the major PI species. The increased prevalence of 38:3‐PI was confirmed by a prior data set that was collected to elucidate the lipidome of the RAW 264.7 cell. They measured the levels of 38:4‐PI at 25 pmol/μg of DNA to the levels of 38:3‐PI at 27 pmol/μg of DNA (230).

Whereas, glycerophospholipid composition of macrophages isolated from the peritoneal

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cavity of mice had a significantly higher level of 38:4‐PI (average of 30 pmol/μg of DNA) compared to that of 38:3‐PI (average of 5 pmol/μg of DNA) (295). Supporting that cells in culture have higher ratio of 20:3 verses 20:4 fatty acyl chain compared to cells isolated from mammals.

Groups have examined the role of 20:3 (5Z, 8Z, 11Z‐ eicosatrienoic acid, Mead acid) metabolism by 5‐lipoxygenase and found that potent bioactive compounds similar to those synthesized from AA are formed, such as 5‐oxo‐ETrE, leukotriene B3, and leukotriene C3

(296‐300). Little work has been performed in regards to these molecules other than their initial characterization because most mammalian systems are not depleted of the ω‐6 essential fatty acids. However, under the current conditions in the RAW 264.7 cell, the formation of 5‐LO products and the reacylation of Mead acid needs to be further examined since it appears to be a more abundant fatty acyl chain in phospholipids than AA. However, because the levels of the ion m/z 887.6 is not changing, it is unlikely that the presence of mead acid is causing changes in the production of leukotrienes. Rather that the formation of products from the oxidation of mead acid may be a better measurement of the affect of the depletion of acyltransferase activity in stimulated cells in culture.

MBOAT7/LPIAT1 in the Regulation of Phosphoinositides in the RAW 264.7 Cell–

Previous work has shown that deletion of MBOAT7/LPIAT1in the mouse led to decreased levels of 18:0/20:4‐PI and 4,5‐PIP2 (247,248). Phosphatidylinositols are critical in numerous cellular functions, however these lipids have been difficult to measure because of their low abundance. Recent methods have been developed using mass spectrometry to quantitate levels of the phosphorylated PI species (207). The method (derivatization) involves the addition of small molecules to the phosphates on the inositol ring. This modification of the phosphates allows for increased extraction efficiency of these low

172

abundant lipids, but also allows for targeted mass spectrometry experiments that take advantage of the loss of the derivative.

To determine if the levels of phosphoinositides change with the depletion of

LPIAT1, phospholipid extracts from non‐targeted shRNA cells and MBOAT7/LPIAT1 shRNA cells could be isolated, modified, and quantitated by the previously established method. Based on previously published data, it would be expected that there would be a decrease in some of the phosphoinositides with the depletion of MBOAT7/LPIAT1 in the

RAW264.7 cells. If there are changes in these lipids, it could potentially explain why there are decreases in leukotrienes compared to the non‐targeting shRNA control cells. However, it will be difficult to determine the mechanism(s) by which these phosphoinositides are leading to a decrease in leukotriene formation because of their multifunctional role in the cell.

The activity of cPLA2α was not tested and compared between the cells that had a non‐targeted shRNA and a targeted shRNA for LPIAT1. It is possible that reduction of phosphoinositides would lead to decreased docking time of cPLA2α to membranes. If there is a decrease in the phosphoinositides, the activity measurements of cPLA2α could be tested in the three cell different cell lines to determine if the decrease in leukotrienes is due to decreased cPLA2α activity.

Concluding remarks–

Taken together, the data presented in this dissertation further supports the role of the reacylation pathway, specifically as it pertains to PI, in the regulation of free AA and leukotriene formation. Many additional experiments need to be performed to determine how the complex regulation of AA is controlled. Identification of the key enzymes in the reacylation pathways of AA could allow for alternative modulation of leukotriene production. Through the development of small molecule inhibitors, specifically against

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certain isoforms of the LPATs and ACSL proteins, an increased understanding of how acute inhibition verse chronic depletion in the reacylation pathway leads to changes in eicosanoid production. The regulation of leukotrienes through direct inhibition of 5‐LO and the receptors for 5‐LO products have only had modest success in the treatment of disease.

Modulating the levels of eicosanoids, instead of completely inhibiting their production, would yield a modulated response and be a more successful strategy to treat diseases where eicosanoids are implemented.

REFERENCES

174

1. Hartley, P. (1907) On the Nature of the Fat Contained in the liver, kidney, and heart J. Physiol. 36, 17‐26

2. Hartley, P. (1909) On the Nature of the Fat Contained in the Liver, Kidney, and Heart. Part II. J. Physiol. 38, 353‐369

3. Roaf, H. E., and Edie, E. S. (1905) A Simple Method for the Preparation and Determination of Lecithin, Williams & Norgate, London

4. Lewkowitsch, J. (1913) Chemical Technology and Analysis of Oils, Fats and Waxes, Etc, 5th ed., London, Macmillan, London

5. Levene, P. A., and Rolf, I. P. (1926) Bromolecithins. II. Bromolecithins of the Liver and Egg Yolk. J Biol Chem 67, 659‐666

6. Klenk, E., and von Schoenebeck, O. (1932). Z. Physiol. Chem. 209, 112

7. Wesson, L. G. (1924) The Isolation of Arachidonc Acid from Brain Tissue. J Biol Chem 60, 183‐187

8. Levene, P. A., and Simms, H. S. (1921) The Liver Lecithin. J Biol Chem 48, 185‐196

9. Cartland, G. F., and Hart, M. C. (1925) The Chemical Investigations of Corpus Luteum: IV. The Acetone‐Soluble Fat. J Biol Chem 66, 619‐637

10. Eckstein, H. C. (1925) The Fatty Acids in the Subcutaneous Fat of Man. J Biol Chem 64, 797‐806

11. Bloor, W. R. (1926) Distribution of Unsaturated Fatty Acids in Tissues. I. Beef Heart Muscle. J Biol Chem 68, 33‐56

12. Toyama, Y., and Tsuchiya, T. (1935) The highly unsaturated acids in sardine oil. V. The constutution of eicosatetraenoic acid C20H32O2. Bull. Chem. Soc. Japan 10, 296‐300

13. Bull, H. B. (1937) The Biochemistry of the Lipid, J. Wiley & sons inc, New York, NY

14. Shinowara, G. Y., and Brown, J. B. (1940) Sudies on the Chemistry of the Fatty Acids: VI. The Application of Crystallization Methods to the Isolation of Arachidonic Acid, with a Comparision of the Properties of the Acid Prepared by Crystallization and by Debromination, Observations on the Structure of Arachidonic Acid. J Biol Chem 134, 331‐340

15. Dolby, D. E., Nunn, L. C. A., and Smedley‐Maclean, I. (1940) The Constitution of Arachidonic Acid (Preliminary Communication). Biochem. J. 34, 1422‐1426

16. Mowry, D. T., Brode, W. R., and Brown, J. B. (1942) Studies on the Chemistry of the Fatty Acids: IX A Spectroscopic Study of Methyl Arachidonate Purified by

175

Crystallization and Distillation and its Alkali Isomerization Product. J Biol Chem 142, 671‐678

17. Mowry, D. T., Brode, W. R., and Brown, J. B. (1942) Studies on the Chemistry of the Fatty Acids: X. The Structure of Arachidonic Acid as Evidenced by Oxidative Degradation and Selective Hydrogenation. J Biol Chem 142, 679‐691

18. Holman, R. T. (1951) Metabolism of isomers of linoleic and linolenic acids. Proc Soc Exp Biol Med 76, 100‐102

19. Shreve, O. D., Heether, M. R., Knight, H. B., and Swern, D. (1950) Infrated Absorption Spectra Some Long‐Chain Fatty Acids, Esters, and Alcohols. Anal Chem 22, 1498‐ 1501

20. McCutcheon, J. W., Crawford, M. F., and Welsh, H. L. (1941) The isomeric structure of the C18 unsaturated acids from their raman and infra red spectra. Oil&Soap 18, 9‐11

21. Sinclair, R. G., McKay, A. F., Myers, G. S., and Jones, R. N. (1952) The Infrared Absorption Spectra of Unsaturated Fatty Acids and Esters. J Am Chem Soc 74, 2578‐ 2585

22. Holman, R. T., and Edmondson, P. R. (1956) Near‐Infrared Spectra of Fatty Acids and Some Related Substances. Anal Chem 28, 1533‐1538

23. Ege, S. N., Wolovsky, R., and Gensler, W. J. (1961) Synthesis of Methyl 5,8,11,14‐ Eicosatetraenoate (Methyl Arachidonate). J. Am. Chem. Soc. 83, 3080‐3085

24. Wesson, L. G. (1925) On a Possible Relationship of Arachidonic Acid to the Saturated Fatty Acids in Fatty Acid Metabolism. J Biol Chem 65, 235‐250

25. Ellis, N. R., and Hankin, O. G. (1925) Soft Pork Studies: I. Formation of Fat in the Pig on a Ration Moderately Low in Fat. J Biol Chem 66, 101‐122

26. Ellis, N. R., and Isbell, H. S. (1926) Soft Pork Studies: II. The Influence of the Character of the Ration Upon the Composition of the Body Fat of Hogs. J Biol Chem 69, 219‐238

27. Ellis, N. R., and Isbell, H. S. (1926) Soft Pork Studies: III. The Effect of Food Fat Upon Body Fat, As Shown by the Separation of the Individual Fatty Acids of the Body Fat. J Biol Chem 69, 239‐248

28. Ellis, N. R., and Zeller, J. H. (1930) Soft Pork Studies: IV. The Influence of a Ration Low in Fat Upon the Composition of the Body Fat of Hogs. J Biol Chem 89, 185‐197

29. Spadola, J. M., and Ellis, N. R. (1936) The Effect of the Ingestion of Cottonseed Oil Before and After Hydrogenation on the Composition of the Body Fat of the Rat. J Biol Chem 113, 205‐218

30. Burr, G. O., and Burr, M. M. (1930) On the Nautre of Role of the Fatty Acids Essential in Nutrition. J Biol Chem 86, 587‐621

176

31. Burr, G. O., Burr, M. M., and Miller, E. S. (1932) On the nature and role of the fatty acids essential in nutrition. J Biol Chem 97, 1‐9

32. Nunn, L. C., and Smedley‐Maclean, I. (1938) The nature of the fatty acids stored by the liver in the fat‐deficiency disease of rats. Biochem J 32, 2178‐2184

33. Van Dorp, D. A., Beerthuis, R. K., Nugteren, D. H., and Vonkeman, H. (1964) The Biosynthesis of Prostaglandins. Biochim Biophys Acta 90, 204‐207

34. Bergstroem, S., Danielsson, H., Klenberg, D., and Samuelsson, B. (1964) The Enzymatic Conversion of Essential Fatty Acids into Prostaglandins. J Biol Chem 239, PC4006‐4008

35. Bergstroem, S., Danielsson, H., and Samuelsson, B. (1964) The Enzymatic Formation of Prostaglandin E2 from Arachidonic Acid Prostaglandins and Related Factors 32. Biochim Biophys Acta 90, 207‐210

36. Van Dorp, D. A., Beerthuis, R. K., Nugteren, D. H., and Vonkeman, H. (1964) Enzymatic Conversion of All‐Cis‐Polyunsaturated Fatty Acids into Prostaglandins. Nature 203, 839‐841

37. Samuelsson, B., Granstrom, E., Green, K., Hamberg, M., and Hammarstrom, S. (1975) Prostaglandins. Annu Rev Biochem 44, 669‐695

38. Borgeat, P., Hamberg, M., and Samuelsson, B. (1976) Transformation of arachidonic acid and homo‐gamma‐linolenic acid by rabbit polymorphonuclear leukocytes. Monohydroxy acids from novel lipoxygenases. J Biol Chem 251, 7816‐7820

39. Murphy, R. C., Hammarstrom, S., and Samuelsson, B. (1979) Leukotriene C: a slow‐ reacting substance from murine mastocytoma cells. Proc Natl Acad Sci U S A 76, 4275‐4279

40. Borgeat, P., and Samuelsson, B. (1979) Transformation of arachidonic acid by rabbit polymorphonuclear leukocytes. Formation of a novel dihydroxyeicosatetraenoic acid. J Biol Chem 254, 2643‐2646

41. Hirata, F., Corcoran, B. A., Venkatasubramanian, K., Schiffmann, E., and Axelrod, J. (1979) Chemoattractants stimulate degradation of methylated phospholipids and release of arachidonic acid in rabbit leukocytes. Proc Natl Acad Sci U S A 76, 2640‐ 2643

42. Stenson, W. F., and Parker, C. W. (1979) Metabolism of arachidonic acid in ionophore‐stimulated neutrophils. Esterification of a hydroxylated metabolite into phospholipids. J Clin Invest 64, 1457‐1465

43. Waite, M., DeChatelet, L. R., King, L., and Shirley, P. S. (1979) Phagocytosis‐induced release of arachidonic acid from human neutrophils. Biochem Biophys Res Commun 90, 984‐992

44. Nalefski, E. A., Sultzman, L. A., Martin, D. M., Kriz, R. W., Towler, P. S., Knopf, J. L., and Clark, J. D. (1994) Delineation of two functionally distinct domains of cytosolic

177

phospholipase A2, a regulatory Ca(2+)‐dependent lipid‐binding domain and a Ca(2+)‐independent catalytic domain. J Biol Chem 269, 18239‐18249

45. Leslie, C. C. (1990) Macrophage phospholipase A2 specific for sn‐2‐arachidonic acid. Methods Enzymol 187, 216‐225

46. de Carvalho, M. G., Garritano, J., and Leslie, C. C. (1995) Regulation of lysophospholipase activity of the 85‐kDa phospholipase A2 and activation in mouse peritoneal macrophages. J Biol Chem 270, 20439‐20446

47. Ghosh, M., Loper, R., Ghomashchi, F., Tucker, D. E., Bonventre, J. V., Gelb, M. H., and Leslie, C. C. (2007) Function, activity, and membrane targeting of cytosolic phospholipase A(2)zeta in mouse lung fibroblasts. J Biol Chem 282, 11676‐11686

48. Glover, S., de Carvalho, M. S., Bayburt, T., Jonas, M., Chi, E., Leslie, C. C., and Gelb, M. H. (1995) Translocation of the 85‐kDa phospholipase A2 from cytosol to the nuclear envelope in rat basophilic leukemia cells stimulated with calcium ionophore or IgE/antigen. J Biol Chem 270, 15359‐15367

49. Goldblatt, M. W. (1935) Properties of human seminal plasma. J Physiol 84, 208‐218

50. van Euler, U. S. (1935) Über die Spezifische Blutdrucksenkende Substanz des Menschlichen Prostata‐ und Samenblasensekretes. Wien Klin Wochenschr 14, 1182‐ 1183

51. Bergstroem, S., and Sjovall, J. (1960) The isolation of prostaglandin E from sheep prostate glands. Acta. Chem. Scand. 14, 1701‐1705

52. Bergstroem, S., and Sjovall, J. (1960) The isolation of prostaglandin F from sheep prostate glands. Acta. Chem. Scand. 14, 1693‐1700

53. Rome, L. H., and Lands, M. E. (1975) Properties of a partially‐purified preparation of the prostaglandin‐forming oxygenase from sheep vesicular gland. Prostaglandins 10, 813‐824

54. Hemler, M., and Lands, W. E. (1976) Purification of the cyclooxygenase that forms prostaglandins. Demonstration of two forms of iron in the holoenzyme. J Biol Chem 251, 5575‐5579

55. DeWitt, D. L., and Smith, W. L. (1988) Primary structure of prostaglandin G/H synthase from sheep vesicular gland determined from the complementary DNA sequence. Proc Natl Acad Sci U S A 85, 1412‐1416

56. Merlie, J. P., Fagan, D., Mudd, J., and Needleman, P. (1988) Isolation and characterization of the complementary DNA for sheep seminal vesicle prostaglandin endoperoxide synthase (cyclooxygenase). J Biol Chem 263, 3550‐ 3553

57. Yokoyama, C., Takai, T., and Tanabe, T. (1988) Primary structure of sheep prostaglandin endoperoxide synthase deduced from cDNA sequence. FEBS Lett 231, 347‐351

178

58. Xie, W. L., Chipman, J. G., Robertson, D. L., Erikson, R. L., and Simmons, D. L. (1991) Expression of a mitogen‐responsive gene encoding prostaglandin synthase is regulated by mRNA splicing. Proc Natl Acad Sci U S A 88, 2692‐2696

59. Yuan, C., and Smith, W. L. (2015) A cyclooxygenase‐2‐dependent prostaglandin E2 biosynthetic system in the Golgi apparatus. J Biol Chem 290, 5606‐5620

60. Kang, Y. J., Mbonye, U. R., DeLong, C. J., Wada, M., and Smith, W. L. (2007) Regulation of intracellular cyclooxygenase levels by gene transcription and protein degradation. Prog Lipid Res 46, 108‐125

61. Dong, L., Vecchio, A. J., Sharma, N. P., Jurban, B. J., Malkowski, M. G., and Smith, W. L. (2011) Human cyclooxygenase‐2 is a sequence homodimer that functions as a conformational heterodimer. J Biol Chem 286, 19035‐19046

62. Kulmacz, R. J., and Lands, W. E. (1984) Prostaglandin H synthase. Stoichiometry of heme cofactor. J Biol Chem 259, 6358‐6363

63. Kulmacz, R. J., and Lands, W. E. (1985) Stoichiometry and kinetics of the interaction of prostaglandin H synthase with anti‐inflammatory agents. J Biol Chem 260, 12572‐12578

64. Yuan, C., Rieke, C. J., Rimon, G., Wingerd, B. A., and Smith, W. L. (2006) Partnering between monomers of cyclooxygenase‐2 homodimers. Proc Natl Acad Sci U S A 103, 6142‐6147

65. Zou, H., Yuan, C., Dong, L., Sidhu, R. S., Hong, Y. H., Kuklev, D. V., and Smith, W. L. (2012) Human cyclooxygenase‐1 activity and its responses to COX inhibitors are allosterically regulated by nonsubstrate fatty acids. J Lipid Res 53, 1336‐1347

66. Smith, W. L., Urade, Y., and Jakobsson, P. J. (2011) Enzymes of the cyclooxygenase pathways of prostanoid biosynthesis. Chem Rev 111, 5821‐5865

67. Smith, W. L., DeWitt, D. L., and Garavito, R. M. (2000) Cyclooxygenases: structural, cellular, and molecular biology. Annu Rev Biochem 69, 145‐182

68. Smith, W. L., and Murphy, R. C. (2008) Cyclooxygenase, lipoxygenase, and epoxygenase pathways. in Biochemistry of Lipids, Lipoproteins and Membranes (Vance, D. E., and Vance, J. E. eds.), 5th Ed., Elsevier. pp 331‐362

69. Sampey, A. V., Monrad, S., and Crofford, L. J. (2005) Microsomal prostaglandin E synthase‐1: the inducible synthase for prostaglandin E2. Arthritis Res Ther 7, 114‐ 117

70. Tanioka, T., Nakatani, Y., Semmyo, N., Murakami, M., and Kudo, I. (2000) Molecular identification of cytosolic prostaglandin E2 synthase that is functionally coupled with cyclooxygenase‐1 in immediate prostaglandin E2 biosynthesis. J Biol Chem 275, 32775‐32782

71. Kanaoka, Y., and Urade, Y. (2003) Hematopoietic prostaglandin D synthase. Prostaglandins Leukot Essent Fatty Acids 69, 163‐167

179

72. White, D. M., Mikol, D. D., Espinosa, R., Weimer, B., Le Beau, M. M., and Stefansson, K. (1992) Structure and chromosomal localization of the human gene for a brain form of prostaglandin D2 synthase. J Biol Chem 267, 23202‐23208

73. Komoto, J., Yamada, T., Watanabe, K., and Takusagawa, F. (2004) Crystal structure of human prostaglandin F synthase (AKR1C3). Biochemistry 43, 2188‐2198

74. Watanabe, K., Yoshida, R., Shimizu, T., and Hayaishi, O. (1985) Enzymatic formation of prostaglandin F2 alpha from prostaglandin H2 and D2. Purification and properties of prostaglandin F synthetase from bovine lung. J Biol Chem 260, 7035‐ 7041

75. Yokoyama, C., Yabuki, T., Inoue, H., Tone, Y., Hara, S., Hatae, T., Nagata, M., Takahashi, E. I., and Tanabe, T. (1996) Human gene encoding prostacyclin synthase (PTGIS): genomic organization, chromosomal localization, and promoter activity. Genomics 36, 296‐304

76. Chiang, C. W., Yeh, H. C., Wang, L. H., and Chan, N. L. (2006) Crystal structure of the human prostacyclin synthase. J Mol Biol 364, 266‐274

77. Needleman, P., Moncada, S., Bunting, S., Vane, J. R., Hamberg, M., and Samuelsson, B. (1976) Identification of an enzyme in platelet microsomes which generates thromboxane A2 from prostaglandin endoperoxides. Nature 261, 558‐560

78. Ohashi, K., Ruan, K. H., Kulmacz, R. J., Wu, K. K., and Wang, L. H. (1992) Primary structure of human thromboxane synthase determined from the cDNA sequence. J Biol Chem 267, 789‐793

79. Wang, L. H., and Kulmacz, R. J. (2002) Thromboxane synthase: structure and function of protein and gene. Prostaglandins Other Lipid Mediat 68‐69, 409‐422

80. Ricciotti, E., and FitzGerald, G. A. (2011) Prostaglandins and inflammation. Arterioscler Thromb Vasc Biol 31, 986‐1000

81. Sugimoto, Y., and Narumiya, S. (2007) Prostaglandin E receptors. J Biol Chem 282, 11613‐11617

82. Aoki, T., and Narumiya, S. (2012) Prostaglandins and chronic inflammation. Trends Pharmacol Sci 33, 304‐311

83. Murakami, M., and Kudo, I. (2004) Recent advances in molecular biology and physiology of the prostaglandin E2‐biosynthetic pathway. Prog Lipid Res 43, 3‐35

84. Saito, S., Tsuda, H., and Michimata, T. (2002) Prostaglandin D2 and reproduction. Am J Reprod Immunol 47, 295‐302

85. Embrey, M. P., and Morrison, D. L. (1968) The effect of prostaglandins on human pregnant myometrium in vitro. J Obstet Gynaecol Br Commonw 75, 829‐832

180

86. Hillier, K., and Karim, S. M. (1968) Effects of prostaglandins E1, E2, F1‐alpha, F2‐ alpha on isolated human umbilical and placental blood vessels. J Obstet Gynaecol Br Commonw 75, 667‐673

87. Hyman, A. L. (1969) The active responses of pulmonary veins in intact dogs to prostaglandins F2 alpha and E1. J Pharmacol Exp Ther 165, 267‐273

88. Hamberg, M., Svensson, J., and Samuelsson, B. (1975) Thromboxanes: a new group of biologically active compounds derived from prostaglandin endoperoxides. Proc Natl Acad Sci U S A 72, 2994‐2998

89. Moncada, S., Gryglewski, R., Bunting, S., and Vane, J. R. (1976) An enzyme isolated from arteries transforms prostaglandin endoperoxides to an unstable substance that inhibits platelet aggregation. Nature 263, 663‐665

90. Whittaker, N., Bunting, S., Salmon, J., Moncada, S., Vane, J. R., Johnson, R. A., Morton, D. R., Kinner, J. H., Gorman, R. R., McGuire, J. C., and Sun, F. F. (1976) The chemical structure of prostaglandin X (prostacyclin). Prostaglandins 12, 915‐928

91. Kellaway, C. H., and Trethewie, E. R. (1940) The Liberation of a Slow‐Reacting Smooth Muscle‐Stimulating Substance in Anaphylaxis. Experimental Physiology 30, 121‐145

92. Brocklehurst, W. E. (1960) The release of histamine and formation of a slow‐ reacting substance (SRS‐A) during anaphylactic shock. J Physiol 151, 416‐435

93. Wasserman, S. I. (1989) Slow Reacting Substance of Anaphylaxis, Springer‐Verlag, New York, NY

94. Matsumoto, T., Funk, C. D., Radmark, O., Hoog, J. O., Jornvall, H., and Samuelsson, B. (1988) Molecular cloning and amino acid sequence of human 5‐lipoxygenase. Proc Natl Acad Sci U S A 85, 26‐30

95. Bonventre, J. V., Huang, Z., Taheri, M. R., O'Leary, E., Li, E., Moskowitz, M. A., and Sapirstein, A. (1997) Reduced fertility and postischaemic brain injury in mice deficient in cytosolic phospholipase A2. Nature 390, 622‐625

96. Dixon, R. A., Diehl, R. E., Opas, E., Rands, E., Vickers, P. J., Evans, J. F., Gillard, J. W., and Miller, D. K. (1990) Requirement of a 5‐lipoxygenase‐activating protein for leukotriene synthesis. Nature 343, 282‐284

97. Miller, D. K., Gillard, J. W., Vickers, P. J., Sadowski, S., Leveille, C., Mancini, J. A., Charleson, P., Dixon, R. A., Ford‐Hutchinson, A. W., Fortin, R., and et al. (1990) Identification and isolation of a membrane protein necessary for leukotriene production. Nature 343, 278‐281

98. Noguchi, M., Miyano, M., Matsumoto, T., and Noma, M. (1994) Human 5‐ lipoxygenase associates with phosphatidylcholine liposomes and modulates LTA4 synthetase activity. Biochim Biophys Acta 1215, 300‐306

181

99. Hammarberg, T., Provost, P., Persson, B., and Radmark, O. (2000) The N‐terminal domain of 5‐lipoxygenase binds calcium and mediates calcium stimulation of enzyme activity. J Biol Chem 275, 38787‐38793

100. Channon, J. Y., and Leslie, C. C. (1990) A calcium‐dependent mechanism for associating a soluble arachidonoyl‐hydrolyzing phospholipase A2 with membrane in the macrophage cell line RAW 264.7. J Biol Chem 265, 5409‐5413

101. Puustinen, T., Scheffer, M. M., and Samuelsson, B. (1988) Regulation of the human leukocyte 5‐lipoxygenase: stimulation by micromolar Ca2+ levels and phosphatidylcholine vesicles. Biochim Biophys Acta 960, 261‐267

102. Nguyen, T., Falgueyret, J. P., Abramovitz, M., and Riendeau, D. (1991) Evaluation of the role of conserved His and Met residues among lipoxygenases by site‐directed mutagenesis of recombinant human 5‐lipoxygenase. J Biol Chem 266, 22057‐22062

103. Ishii, S., Noguchi, M., Miyano, M., Matsumoto, T., and Noma, M. (1992) Mutagenesis studies on the amino acid residues involved in the iron‐binding and the activity of human 5‐lipoxygenase. Biochem Biophys Res Commun 182, 1482‐1490

104. Gilbert, N. C., Bartlett, S. G., Waight, M. T., Neau, D. B., Boeglin, W. E., Brash, A. R., and Newcomer, M. E. (2011) The structure of human 5‐lipoxygenase. Science 331, 217‐ 219

105. Zhang, Y. Y., Radmark, O., and Samuelsson, B. (1992) Mutagenesis of some conserved residues in human 5‐lipoxygenase: effects on enzyme activity. Proc Natl Acad Sci U S A 89, 485‐489

106. Haeggstrom, J. Z., and Funk, C. D. (2011) Lipoxygenase and leukotriene pathways: biochemistry, biology, and roles in disease. Chem Rev 111, 5866‐5898

107. Rouzer, C. A., Ford‐Hutchinson, A. W., Morton, H. E., and Gillard, J. W. (1990) MK886, a potent and specific leukotriene biosynthesis inhibitor blocks and reverses the membrane association of 5‐lipoxygenase in ionophore‐challenged leukocytes. J Biol Chem 265, 1436‐1442

108. Abramovitz, M., Wong, E., Cox, M. E., Richardson, C. D., Li, C., and Vickers, P. J. (1993) 5‐lipoxygenase‐activating protein stimulates the utilization of arachidonic acid by 5‐lipoxygenase. Eur J Biochem 215, 105‐111

109. Rinaldo‐Matthis, A., Wetterholm, A., Martinez Molina, D., Holm, J., Niegowski, D., Ohlson, E., Nordlund, P., Morgenstern, R., and Haeggstrom, J. Z. (2010) Arginine 104 is a key catalytic residue in leukotriene C4 synthase. J Biol Chem 285, 40771‐40776

110. Mancini, J. A., Abramovitz, M., Cox, M. E., Wong, E., Charleson, S., Perrier, H., Wang, Z., Prasit, P., and Vickers, P. J. (1993) 5‐lipoxygenase‐activating protein is an arachidonate binding protein. FEBS Lett 318, 277‐281

111. Samuelsson, B., and Funk, C. D. (1989) Enzymes involved in the biosynthesis of leukotriene B4. J Biol Chem 264, 19469‐19472

182

112. Haeggstrom, J. Z., and Wetterholm, A. (2002) Enzymes and receptors in the leukotriene cascade. Cell Mol Life Sci 59, 742‐753

113. Radmark, O., Shimizu, T., Jornvall, H., and Samuelsson, B. (1984) Leukotriene A4 hydrolase in human leukocytes. Purification and properties. J Biol Chem 259, 12339‐12345

114. Ohishi, N., Izumi, T., Minami, M., Kitamura, S., Seyama, Y., Ohkawa, S., Terao, S., Yotsumoto, H., Takaku, F., and Shimizu, T. (1987) Leukotriene A4 hydrolase in the human lung. Inactivation of the enzyme with leukotriene A4 isomers. J Biol Chem 262, 10200‐10205

115. Lam, B. K., Penrose, J. F., Freeman, G. J., and Austen, K. F. (1994) Expression cloning of a cDNA for human leukotriene C4 synthase, an integral membrane protein conjugating reduced glutathione to leukotriene A4. Proc Natl Acad Sci U S A 91, 7663‐7667

116. Ago, H., Kanaoka, Y., Irikura, D., Lam, B. K., Shimamura, T., Austen, K. F., and Miyano, M. (2007) Crystal structure of a human membrane protein involved in cysteinyl leukotriene biosynthesis. Nature 448, 609‐612

117. Murphy, R. C., and Gijon, M. A. (2007) Biosynthesis and metabolism of leukotrienes. Biochem J 405, 379‐395

118. Penrose, J. F., Spector, J., Lam, B. K., Friend, D. S., Xu, K., Jack, R. M., and Austen, K. F. (1995) Purification of human lung leukotriene C4 synthase and preparation of a polyclonal antibody. Am J Respir Crit Care Med 152, 283‐289

119. Lam, B. K., and Austen, K. F. (2002) Leukotriene C4 synthase: a pivotal enzyme in cellular biosynthesis of the cysteinyl leukotrienes. Prostaglandins Other Lipid Mediat 68‐69, 511‐520

120. Gijon, M. A., Zarini, S., and Murphy, R. C. (2007) Biosynthesis of eicosanoids and transcellular metabolism of leukotrienes in murine bone marrow cells. J Lipid Res 48, 716‐725

121. Lam, B. K., Gagnon, L., Austen, K. F., and Soberman, R. J. (1990) The mechanism of leukotriene B4 export from human polymorphonuclear leukocytes. J Biol Chem 265, 13438‐13441

122. Leier, I., Jedlitschky, G., Buchholz, U., Cole, S. P., Deeley, R. G., and Keppler, D. (1994) The MRP gene encodes an ATP‐dependent export pump for leukotriene C4 and structurally related conjugates. J Biol Chem 269, 27807‐27810

123. Rius, M., Hummel‐Eisenbeiss, J., and Keppler, D. (2008) ATP‐dependent transport of leukotrienes B4 and C4 by the multidrug resistance protein ABCC4 (MRP4). J Pharmacol Exp Ther 324, 86‐94

124. Owman, C., Nilsson, C., and Lolait, S. J. (1996) Cloning of cDNA encoding a putative chemoattractant receptor. Genomics 37, 187‐194

183

125. Yokomizo, T., Izumi, T., Chang, K., Takuwa, Y., and Shimizu, T. (1997) A G‐protein‐ coupled receptor for leukotriene B4 that mediates chemotaxis. Nature 387, 620‐ 624

126. Yokomizo, T., Kato, K., Hagiya, H., Izumi, T., and Shimizu, T. (2001) Hydroxyeicosanoids bind to and activate the low affinity leukotriene B4 receptor, BLT2. J Biol Chem 276, 12454‐12459

127. Haribabu, B., Verghese, M. W., Steeber, D. A., Sellars, D. D., Bock, C. B., and Snyderman, R. (2000) Targeted disruption of the leukotriene B(4) receptor in mice reveals its role in inflammation and platelet‐activating factor‐induced anaphylaxis. J Exp Med 192, 433‐438

128. Tager, A. M., Dufour, J. H., Goodarzi, K., Bercury, S. D., von Andrian, U. H., and Luster, A. D. (2000) BLTR mediates leukotriene B(4)‐induced chemotaxis and adhesion and plays a dominant role in eosinophil accumulation in a murine model of peritonitis. J Exp Med 192, 439‐446

129. Tager, A. M., and Luster, A. D. (2003) BLT1 and BLT2: the leukotriene B(4) receptors. Prostaglandins Leukot Essent Fatty Acids 69, 123‐134

130. Iizuka, Y., Okuno, T., Saeki, K., Uozaki, H., Okada, S., Misaka, T., Sato, T., Toh, H., Fukayama, M., Takeda, N., Kita, Y., Shimizu, T., Nakamura, M., and Yokomizo, T. (2010) Protective role of the leukotriene B4 receptor BLT2 in murine inflammatory colitis. FASEB J 24, 4678‐4690

131. Yokomizo, T. (2011) Leukotriene B4 receptors: novel roles in immunological regulations. Adv Enzyme Regul 51, 59‐64

132. Lynch, K. R., O'Neill, G. P., Liu, Q., Im, D. S., Sawyer, N., Metters, K. M., Coulombe, N., Abramovitz, M., Figueroa, D. J., Zeng, Z., Connolly, B. M., Bai, C., Austin, C. P., Chateauneuf, A., Stocco, R., Greig, G. M., Kargman, S., Hooks, S. B., Hosfield, E., Williams, D. L., Jr., Ford‐Hutchinson, A. W., Caskey, C. T., and Evans, J. F. (1999) Characterization of the human cysteinyl leukotriene CysLT1 receptor. Nature 399, 789‐793

133. Jones, T. R., Zamboni, R., Belley, M., Champion, E., Charette, L., Ford‐Hutchinson, A. W., Frenette, R., Gauthier, J. Y., Leger, S., Masson, P., and et al. (1989) Pharmacology of L‐660,711 (MK‐571): a novel potent and selective leukotriene D4 receptor antagonist. Can J Physiol Pharmacol 67, 17‐28

134. Heise, C. E., O'Dowd, B. F., Figueroa, D. J., Sawyer, N., Nguyen, T., Im, D. S., Stocco, R., Bellefeuille, J. N., Abramovitz, M., Cheng, R., Williams, D. L., Jr., Zeng, Z., Liu, Q., Ma, L., Clements, M. K., Coulombe, N., Liu, Y., Austin, C. P., George, S. R., O'Neill, G. P., Metters, K. M., Lynch, K. R., and Evans, J. F. (2000) Characterization of the human cysteinyl leukotriene 2 receptor. J Biol Chem 275, 30531‐30536

135. Dahlen, S. E., Hedqvist, P., Hammarstrom, S., and Samuelsson, B. (1980) Leukotrienes are potent constrictors of human bronchi. Nature 288, 484‐486

184

136. Radmark, O., and Samuelsson, B. (2007) 5‐lipoxygenase: regulation and possible involvement in atherosclerosis. Prostaglandins Other Lipid Mediat 83, 162‐174

137. Farias, S. E., Heidenreich, K. A., Wohlauer, M. V., Murphy, R. C., and Moore, E. E. (2011) Lipid mediators in cerebral spinal fluid of traumatic brain injured patients. J Trauma 71, 1211‐1218

138. Knab, L. M., Grippo, P. J., and Bentrem, D. J. (2014) Involvement of eicosanoids in the pathogenesis of pancreatic cancer: the roles of cyclooxygenase‐2 and 5‐ lipoxygenase. World J Gastroenterol 20, 10729‐10739

139. Jack, D. B. (1997) One hundred years of aspirin. Lancet 350, 437‐439

140. Piper, P. J., and Vane, J. R. (1969) Release of additional factors in anaphylaxis and its antagonism by anti‐inflammatory drugs. Nature 223, 29‐35

141. Vane, J. R. (1971) Inhibition of prostaglandin synthesis as a mechanism of action for aspirin‐like drugs. Nat New Biol 231, 232‐235

142. Ferreira, S. H., Moncada, S., and Vane, J. R. (1971) Indomethacin and aspirin abolish prostaglandin release from the spleen. Nat New Biol 231, 237‐239

143. Drazen, J. M., Israel, E., and O'Byrne, P. M. (1999) Treatment of asthma with drugs modifying the leukotriene pathway. N Engl J Med 340, 197‐206

144. Carter, G. W., Young, P. R., Albert, D. H., Bouska, J., Dyer, R., Bell, R. L., Summers, J. B., and Brooks, D. W. (1991) 5‐lipoxygenase inhibitory activity of zileuton. J Pharmacol Exp Ther 256, 929‐937

145. Watkins, P. A., Maiguel, D., Jia, Z., and Pevsner, J. (2007) Evidence for 26 distinct acyl‐coenzyme A synthetase genes in the human genome. J Lipid Res 48, 2736‐2750

146. FDA. (2009) Updated Information on Leukotriene Inhibitors: Montelukast (marketed as Singulair), Zafirlukast (marketed as Accolate), and Zileuton (marketed as Zyflo and Zyflo CR). http://www.fda.gov/Drugs/DrugSafety/PostmarketDrugSafetyInformationf orPatientsandProviders/DrugSafetyInformationforHeathcareProfessionals/ ucm165489.htm

147. Gillard, J., Ford‐Hutchinson, A. W., Chan, C., Charleson, S., Denis, D., Foster, A., Fortin, R., Leger, S., McFarlane, C. S., Morton, H., and et al. (1989) L‐663,536 (MK‐ 886) (3‐[1‐(4‐chlorobenzyl)‐3‐t‐butyl‐thio‐5‐isopropylindol‐2‐yl]‐2,2 ‐ dimethylpropanoic acid), a novel, orally active leukotriene biosynthesis inhibitor. Can J Physiol Pharmacol 67, 456‐464

148. Hutchinson, J. H., Li, Y., Arruda, J. M., Baccei, C., Bain, G., Chapman, C., Correa, L., Darlington, J., King, C. D., Lee, C., Lorrain, D., Prodanovich, P., Rong, H., Santini, A., Stock, N., Prasit, P., and Evans, J. F. (2009) 5‐lipoxygenase‐activating protein inhibitors: development of 3‐[3‐tert‐butylsulfanyl‐1‐[4‐(6‐methoxy‐pyridin‐3‐yl)‐

185

benzyl]‐5‐(pyridin‐2‐ylmeth oxy)‐1H‐indol‐2‐yl]‐2,2‐dimethyl‐propionic acid (AM103). J Med Chem 52, 5803‐5815

149. Stock, N. S., Bain, G., Zunic, J., Li, Y., Ziff, J., Roppe, J., Santini, A., Darlington, J., Prodanovich, P., King, C. D., Baccei, C., Lee, C., Rong, H., Chapman, C., Broadhead, A., Lorrain, D., Correa, L., Hutchinson, J. H., Evans, J. F., and Prasit, P. (2011) 5‐ Lipoxygenase‐activating protein (FLAP) inhibitors. Part 4: development of 3‐[3‐ tert‐butylsulfanyl‐1‐[4‐(6‐ethoxypyridin‐3‐yl)benzyl]‐5‐(5‐methylpyridin‐2‐y lmethoxy)‐1H‐indol‐2‐yl]‐2,2‐dimethylpropionic acid (AM803), a potent, oral, once daily FLAP inhibitor. J Med Chem 54, 8013‐8029

150. Kent, S. E., Bentley, J. H., Miller, D., Sterling, R., Menendez, R., Tarpay, M., Pearlman, D. S., and Norris, V. (2014) The effect of GSK2190915, a 5‐lipoxygenase‐activating protein inhibitor, on exercise‐induced bronchoconstriction. Allergy Asthma Proc 35, 126‐133

151. Barchuk, W., Lambert, J., Fuhr, R., Jiang, J. Z., Bertelsen, K., Fourie, A., Liu, X., Silkoff, P. E., Barnathan, E. S., and Thurmond, R. (2014) Effects of JNJ‐40929837, a leukotriene A4 hydrolase inhibitor, in a bronchial allergen challenge model of asthma. Pulm Pharmacol Ther 29, 15‐23

152. Gobley, T. N. (1847) Recherches Chimiques sur le Jaune d'Oeuf. J. Pharm. Chim. 11, 409‐417

153. Gobley, N. T. (1874) Sur la lécithine et la cérébrine. J de Parmacie et de Chimie, 346‐ 354

154. Kennedy, E. P. (1956) The biological synthesis of phospholipids. Can J Biochem Physiol 34, 334‐348

155. Kennedy, E. P., and Weiss, S. B. (1956) The function of cytidine coenzymes in the biosynthesis of phospholipides. J Biol Chem 222, 193‐214

156. Mitchell, J. R., and Saggerson, E. D. (1994) A study of glycerolphosphate acyltransferase in rat liver mitochondria‐submitochondrial localization and some properties of the solubilized enzyme. Int J Biochem 26, 181‐187

157. Cao, J., Li, J. L., Li, D., Tobin, J. F., and Gimeno, R. E. (2006) Molecular identification of microsomal acyl‐CoA:glycerol‐3‐phosphate acyltransferase, a key enzyme in de novo triacylglycerol synthesis. Proc Natl Acad Sci U S A 103, 19695‐19700

158. Lehner, R., and Kuksis, A. (1996) Biosynthesis of triacylglycerols. Prog Lipid Res 35, 169‐201

159. Monroy, G., Kelker, H. C., and Pullman, M. E. (1973) Partial purification and properties of an acyl coenzyme A:sn‐glycerol 3‐phosphate acyltransferase from rat liver mitochondria. J Biol Chem 248, 2845‐2852

160. Wang, S., Lee, D. P., Gong, N., Schwerbrock, N. M., Mashek, D. G., Gonzalez‐Baro, M. R., Stapleton, C., Li, L. O., Lewin, T. M., and Coleman, R. A. (2007) Cloning and

186

functional characterization of a novel mitochondrial N‐ethylmaleimide‐sensitive glycerol‐3‐phosphate acyltransferase (GPAT2). Arch Biochem Biophys 465, 347‐358

161. Grevengoed, T. J., Klett, E. L., and Coleman, R. A. (2014) Acyl‐CoA metabolism and partitioning. Annu Rev Nutr 34, 1‐30

162. Shindou, H., and Shimizu, T. (2009) Acyl‐CoA:lysophospholipid acyltransferases. J Biol Chem 284, 1‐5

163. Gibellini, F., and Smith, T. K. (2010) The Kennedy pathway‐‐De novo synthesis of phosphatidylethanolamine and phosphatidylcholine. IUBMB Life 62, 414‐428

164. Kennedy, E. P., and Weiss, S. B. (1955) Cytidine diphosphate choline: a new intermediate in lecithin biosynthesis. J. Am. Chem. Soc. 77, 250‐251

165. McMaster, C. R., and Bell, R. M. (1997) CDP‐choline:1,2‐diacylglycerol cholinephosphotransferase. Biochim Biophys Acta 1348, 100‐110

166. Vance, J. E., and Steenbergen, R. (2005) Metabolism and functions of phosphatidylserine. Prog Lipid Res 44, 207‐234

167. Ohsawa, T., Nishijima, M., and Kuge, O. (2004) Functional analysis of Chinese hamster phosphatidylserine synthase 1 through systematic alanine mutagenesis. Biochem J 381, 853‐859

168. Satre, M., and Kennedy, E. P. (1978) Identification of bound pyruvate essential for the activity of phosphatidylserine decarboxylase of Escherichia coli. J Biol Chem 253, 479‐483

169. Vance, D. E., and Vance, J. E. (2008) Biochemistry of Lipids, Lipoproteins and Membranes, 5th ed., Elsevier

170. Noga, A. A., Zhao, Y., and Vance, D. E. (2002) An unexpected requirement for phosphatidylethanolamine N‐methyltransferase in the secretion of very low density lipoproteins. J Biol Chem 277, 42358‐42365

171. Li, Z., Agellon, L. B., and Vance, D. E. (2005) Phosphatidylcholine homeostasis and liver failure. J Biol Chem 280, 37798‐37802

172. Bremer, J., Figard, P. H., and Greenberg, D. M. (1960) The Biosynthesis of Choline and its Relation to Phospholipid Metabolism. Biochemica et Biophysica Acta 43, 477‐488

173. Anderson, R. J. (1930) The chemistry of the lipoids of tubercle bacilli. XIV. The occurrence of inosite in the phosphatide from human tubercle bacilli. J. Am. Chem. Soc. 52, 1607‐1608

174. Klenk, E., and Sakai, R. (1939) Inositmonophosphorsaure, ein Spaltprodukt der Sojabohnenphosphatide. Z. f. physiol. Chemie. 258, 33

187

175. Folch, J., and Woolley, D. (1942) Inositol, A constituent of Brain Phosphatide. J Biol Chem 142, 963‐964

176. Nikawa, J., Kodaki, T., and Yamashita, S. (1987) Primary structure and disruption of the phosphatidylinositol synthase gene of Saccharomyces cerevisiae. J Biol Chem 262, 4876‐4881

177. D'Souza, K., and Epand, R. M. (2015) The phosphatidylinositol synthase‐catalyzed formation of phosphatidylinositol does not exhibit acyl chain specificity. Biochemistry 54, 1151‐1153

178. Pangborn, M. C. (1942) Isolation and Purification of a Serologically Active Phospholipid from Beef Heart. J Biol Chem 143, 247‐256

179. de Haas, G. H., and van Deenen, L. L. (1965) Chemical structure and serological activity of synthetic and natural cardiolipin. Nature 206, 935

180. Benson, A. A., and Maruo, B. (1958) Piant phospholipids. I. Identification of the phosphatidyl glycerols. Biochim Biophys Acta 27, 189‐195

181. Haverkate, F., and van Deenen, L. L. (1965) Isolation and chemical characterization of phosphatidyl glycerol from spinach leaves. Biochim Biophys Acta 106, 78‐92

182. Kiyasu, J. Y., Pieringer, R. A., Paulus, H., and Kennedy, E. P. (1963) The biosynthesis of phosphatidylglycerol. J Biol Chem 238, 2293‐2298

183. Hostetler, K. Y., Van den Bosch, H., and Van Deenen, L. L. (1971) Biosynthesis of cardiolipin in liver mitochondria. Biochim Biophys Acta 239, 113‐119

184. Shindou, H., Hishikawa, D., Harayama, T., Yuki, K., and Shimizu, T. (2009) Recent progress on acyl CoA: lysophospholipid acyltransferase research. J Lipid Res 50 Suppl, S46‐51

185. Lands, W. E. (1960) Metabolism of glycerolipids. 2. The enzymatic acylation of lysolecithin. J Biol Chem 235, 2233‐2237

186. Yamashita, A., Hayashi, Y., Nemoto‐Sasaki, Y., Ito, M., Oka, S., Tanikawa, T., Waku, K., and Sugiura, T. (2014) Acyltransferases and transacylases that determine the fatty acid composition of glycerolipids and the metabolism of bioactive lipid mediators in mammalian cells and model organisms. Prog Lipid Res 53, 18‐81

187. Jain, S., Zhang, X., Khandelwal, P. J., Saunders, A. J., Cummings, B. S., and Oelkers, P. (2009) Characterization of human lysophospholipid acyltransferase 3. J Lipid Res 50, 1563‐1570

188. Nakanishi, H., Shindou, H., Hishikawa, D., Harayama, T., Ogasawara, R., Suwabe, A., Taguchi, R., and Shimizu, T. (2006) Cloning and characterization of mouse lung‐ type acyl‐CoA:lysophosphatidylcholine acyltransferase 1 (LPCAT1). Expression in alveolar type II cells and possible involvement in surfactant production. J Biol Chem 281, 20140‐20147

188

189. Zhao, Y., Chen, Y. Q., Bonacci, T. M., Bredt, D. S., Li, S., Bensch, W. R., Moller, D. E., Kowala, M., Konrad, R. J., and Cao, G. (2008) Identification and characterization of a major liver lysophosphatidylcholine acyltransferase. J Biol Chem 283, 8258‐8265

190. Harayama, T., Shindou, H., and Shimizu, T. (2009) Biosynthesis of phosphatidylcholine by human lysophosphatidylcholine acyltransferase 1. J Lipid Res 50, 1824‐1831

191. Lee, H. C., Inoue, T., Imae, R., Kono, N., Shirae, S., Matsuda, S., Gengyo‐Ando, K., Mitani, S., and Arai, H. (2008) Caenorhabditis elegans mboa‐7, a member of the MBOAT family, is required for selective incorporation of polyunsaturated fatty acids into phosphatidylinositol. Mol Biol Cell 19, 1174‐1184

192. Gijon, M. A., Riekhof, W. R., Zarini, S., Murphy, R. C., and Voelker, D. R. (2008) Lysophospholipid acyltransferases and arachidonate recycling in human neutrophils. J Biol Chem 283, 30235‐30245

193. Kazachkov, M., Chen, Q., Wang, L., and Zou, J. (2008) Substrate preferences of a lysophosphatidylcholine acyltransferase highlight its role in phospholipid remodeling. Lipids 43, 895‐902

194. Cao, J., Shan, D., Revett, T., Li, D., Wu, L., Liu, W., Tobin, J. F., and Gimeno, R. E. (2008) Molecular identification of a novel mammalian brain isoform of acyl‐ CoA:lysophospholipid acyltransferase with prominent ethanolamine lysophospholipid acylating activity, LPEAT2. J Biol Chem 283, 19049‐19057

195. Yang, Y., Cao, J., and Shi, Y. (2004) Identification and characterization of a gene encoding human LPGAT1, an endoplasmic reticulum‐associated lysophosphatidylglycerol acyltransferase. J Biol Chem 279, 55866‐55874

196. Shindou, H., Eto, M., Morimoto, R., and Shimizu, T. (2009) Identification of membrane O‐acyltransferase family motifs. Biochem Biophys Res Commun 383, 320‐325

197. Chen, X., Hyatt, B. A., Mucenski, M. L., Mason, R. J., and Shannon, J. M. (2006) Identification and characterization of a lysophosphatidylcholine acyltransferase in alveolar type II cells. Proc Natl Acad Sci U S A 103, 11724‐11729

198. van Meer, G., Voelker, D. R., and Feigenson, G. W. (2008) Membrane lipids: where they are and how they behave. Nat Rev Mol Cell Biol 9, 112‐124

199. Shindou, H., Hishikawa, D., Nakanishi, H., Harayama, T., Ishii, S., Taguchi, R., and Shimizu, T. (2007) A single enzyme catalyzes both platelet‐activating factor production and membrane biogenesis of inflammatory cells. Cloning and characterization of acetyl‐CoA:LYSO‐PAF acetyltransferase. J Biol Chem 282, 6532‐ 6539

200. Aguado, B., and Campbell, R. D. (1998) Characterization of a human lysophosphatidic acid acyltransferase that is encoded by a gene located in the class III region of the human major histocompatibility complex. J Biol Chem 273, 4096‐ 4105

189

201. Hollenback, D., Bonham, L., Law, L., Rossnagle, E., Romero, L., Carew, H., Tompkins, C. K., Leung, D. W., Singer, J. W., and White, T. (2006) Substrate specificity of lysophosphatidic acid acyltransferase beta ‐‐ evidence from membrane and whole cell assays. J Lipid Res 47, 593‐604

202. Yuki, K., Shindou, H., Hishikawa, D., and Shimizu, T. (2009) Characterization of mouse lysophosphatidic acid acyltransferase 3: an enzyme with dual functions in the testis. J Lipid Res 50, 860‐869

203. Burke, J. E., and Dennis, E. A. (2009) Phospholipase A2 structure/function, mechanism, and signaling. J Lipid Res 50 Suppl, S237‐242

204. Maffucci, T. (2012) Chapter 1: An Introduction to Phosphoinositides,

205. Kutateladze, T. G. (2012) Molecular analysis of protein‐phosphoinositide interactions. Curr Top Microbiol Immunol 362, 111‐126

206. Kutateladze, T. G. (2010) Translation of the phosphoinositide code by PI effectors. Nat Chem Biol 6, 507‐513

207. Wakelam, M. J., and Clark, J. (2011) Methods for analyzing phosphoinositides using mass spectrometry. Biochim Biophys Acta 1811, 758‐762

208. D'Souza, K., and Epand, R. M. (2014) Enrichment of phosphatidylinositols with specific acyl chains. Biochim Biophys Acta 1838, 1501‐1508

209. Griffiths, J. (2008) A brief history of mass spectrometry. Anal Chem 80, 5678‐5683

210. Fenn, J. B., Mann, M., Meng, C. K., Wong, S. F., and Whitehouse, C. M. (1989) Electrospray ionization for mass spectrometry of large biomolecules. Science 246, 64‐71

211. Karas, M., Bachmann, D., and Hillenkamp, F. (1985) Influence of the wavelength in high‐irradiance ultraviolet laser desorption mass spectrometry of organic molecules. Analytical Chemistry 57, 2953‐2959

212. Zenobi, R., and Knochenmuss, R. (1998) Ion Formation in MALDI Mass Spectrometry. Mass Spectrometry Reviews 17, 337‐366

213. Murphy, R. C., Hankin, J. A., and Barkley, R. M. (2009) Imaging of lipid species by MALDI mass spectrometry. J Lipid Res 50 Suppl, S317‐322

214. Zemski Berry, K. A., Gordon, W. C., Murphy, R. C., and Bazan, N. G. (2014) Spatial organization of lipids in the human retina and optic nerve by MALDI imaging mass spectrometry. J Lipid Res 55, 504‐515

215. Kebarle, P., and Verkerk, U. H. (2009) Electrospray: from ions in solution to ions in the gas phase, what we know now. Mass Spectrom Rev 28, 898‐917

216. Konermann, L., Ahadi, E., Rodriguez, A. D., and Vahidi, S. (2013) Unraveling the mechanism of electrospray ionization. Anal Chem 85, 2‐9

190

217. Wu, X., Sadeghi, M., and Vertes, A. (1998) Molecular Dynamics of Matrix‐Assisted Laser Desoption of Leucine Enkephalin Guest Molecules from Nicotinic Acid Host Crystals. J Phys Chem B 102, 4770‐4778

218. Kero, F. A., Yost, R. A., and Pedder, R. E. (2005) Quadrupole mass analyzers: theoretical and practical considerations. Core Methodologies, 1‐9

219. Mathieu, E. (1886) Mémoire sur le mouvement vibratoire d’une membrane de forme elliptique. J. Math. Pures Appl. 13, 137‐203

220. Wolff, M. M., and Stephens, W. E. (1953) A Pulsed Mass Spectrometer with Time Dispersion. Review of Scientific Instruments 24, 616‐617

221. Cornett, D. S., Reyzer, M. L., Chaurand, P., and Caprioli, R. M. (2007) MALDI imaging mass spectrometry: molecular snapshots of biochemical systems. Nat Methods 4, 828‐833

222. Berry, K. A., Hankin, J. A., Barkley, R. M., Spraggins, J. M., Caprioli, R. M., and Murphy, R. C. (2011) MALDI imaging of lipid biochemistry in tissues by mass spectrometry. Chem Rev 111, 6491‐6512

223. Kerkhoff, C., Habben, K., Gehring, L., Resch, K., and Kaever, V. (1998) Substrate specificity of acyl‐CoA:Lysophospholipid acyltransferase (LAT) from pig spleen. Arch Biochem Biophys 351, 220‐226

224. Koeberle, A., Shindou, H., Harayama, T., Yuki, K., and Shimizu, T. (2012) Polyunsaturated fatty acids are incorporated into maturating male mouse germ cells by lysophosphatidic acid acyltransferase 3. FASEB J 26, 169‐180

225. Martin, S. A., Gijon, M. A., Voelker, D. R., and Murphy, R. C. (2014) Measurement of lysophospholipid acyltransferase activities using substrate competition. J Lipid Res 55, 782‐791

226. Bligh, E. G., and Dyer, W. J. (1959) A rapid method of total lipid extraction and purification. Can J Biochem Physiol 37, 911‐917

227. Hall, L. M., and Murphy, R. C. (1998) Electrospray mass spectrometric analysis of 5‐ hydroperoxy and 5‐hydroxyeicosatetraenoic acids generated by lipid peroxidation of red blood cell ghost phospholipids. J Am Soc Mass Spectrom 9, 527‐532

228. Murphy, R. C. (2013) Tandem Mass Spectrometry of Lipids, Royal Society of Chemistry, Cambridge, UK

229. Stafford, R. E., Fanni, T., and Dennis, E. A. (1989) Interfacial properties and critical micelle concentration of lysophospholipids. Biochemistry 28, 5113‐5120

230. Dennis, E. A., Deems, R. A., Harkewicz, R., Quehenberger, O., Brown, H. A., Milne, S. B., Myers, D. S., Glass, C. K., Hardiman, G., Reichart, D., Merrill, A. H., Jr., Sullards, M. C., Wang, E., Murphy, R. C., Raetz, C. R., Garrett, T. A., Guan, Z., Ryan, A. C., Russell, D. W., McDonald, J. G., Thompson, B. M., Shaw, W. A., Sud, M., Zhao, Y., Gupta, S.,

191

Maurya, M. R., Fahy, E., and Subramaniam, S. (2010) A mouse macrophage lipidome. J Biol Chem 285, 39976‐39985

231. Chilton, F. H., and Murphy, R. C. (1986) Remodeling of arachidonate‐containing phosphoglycerides within the human neutrophil. J Biol Chem 261, 7771‐7777

232. Hofmann, K. (2000) A superfamily of membrane‐bound O‐acyltransferases with implications for wnt signaling. Trends Biochem Sci 25, 111‐112

233. Riekhof, W. R., Wu, J., Jones, J. L., and Voelker, D. R. (2007) Identification and characterization of the major lysophosphatidylethanolamine acyltransferase in Saccharomyces cerevisiae. J Biol Chem 282, 28344‐28352

234. Hishikawa, D., Shindou, H., Kobayashi, S., Nakanishi, H., Taguchi, R., and Shimizu, T. (2007) Discovery of a lysophospholipid acyltransferase family essential for membrane asymmetry and diversity. PNAS 105, 2830‐2835

235. Clarkson, T. W., and Magos, L. (2006) The toxicology of mercury and its chemical compounds. Crit Rev Toxicol 36, 609‐662

236. Zarini, S., Gijon, M. A., Folco, G., and Murphy, R. C. (2006) Effect of arachidonic acid reacylation on leukotriene biosynthesis in human neutrophils stimulated with granulocyte‐macrophage colony‐stimulating factor and formyl‐methionyl‐leucyl‐ phenylalanine. J Biol Chem 281, 10134‐10142

237. Demeure, O., Lecerf, F., Duby, C., Desert, C., Ducheix, S., Guillou, H., and Lagarrigue, S. (2011) Regulation of LPCAT3 by LXR. Gene 470, 7‐11

238. Matsuda, S., Inoue, T., Lee, H. C., Kono, N., Tanaka, F., Gengyo‐Ando, K., Mitani, S., and Arai, H. (2008) Member of the membrane‐bound O‐acyltransferase (MBOAT) family encodes a lysophospholipid acyltransferase with broad substrate specificity. Genes Cells 13, 879‐888

239. Quehenberger, O., Armando, A. M., Brown, A. H., Milne, S. B., Myers, D. S., Merrill, A. H., Bandyopadhyay, S., Jones, K. N., Kelly, S., Shaner, R. L., Sullards, C. M., Wang, E., Murphy, R. C., Barkley, R. M., Leiker, T. J., Raetz, C. R., Guan, Z., Laird, G. M., Six, D. A., Russell, D. W., McDonald, J. G., Subramaniam, S., Fahy, E., and Dennis, E. A. (2010) Lipidomics reveals a remarkable diversity of lipids in human plasma. J Lipid Res 51, 3299‐3305

240. Ichi, I., Kono, N., Arita, Y., Haga, S., Arisawa, K., Yamano, M., Nagase, M., Fujiwara, Y., and Arai, H. (2014) Identification of genes and pathways involved in the synthesis of Mead acid (20:3n‐9), an indicator of essential fatty acid deficiency. Biochim Biophys Acta 1841, 204‐213

241. Bell, R. L., Kennerly, D. A., Stanford, N., and Majerus, P. W. (1979) Diglyceride lipase: a pathway for arachidonate release from human platelets. Proc Natl Acad Sci U S A 76, 3238‐3241

242. Bell, R. L., and Majerus, P. W. (1980) Thrombin‐induced hydrolysis of phosphatidylinositol in human platelets. J Biol Chem 255, 1790‐1792

192

243. Takenawa, T., Homma, Y., and Nagai, Y. (1983) Role of Ca2+ in phosphotidylinositol response and arachidonic acid release in formylated tripeptide‐or Ca2+ ionophore A23187‐stimulated guinea pig neutrophils. J Immunol 130, 2849‐2855

244. Prescott, S. M., and Majerus, P. W. (1981) The fatty acid composition of phosphatidylinositol from thrombin‐stimulated human platelets. J Biol Chem 256, 579‐582

245. Balsinde, J., Balboa, M. A., Li, W. H., Llopis, J., and Dennis, E. A. (2000) Cellular regulation of cytosolic group IV phospholipase A2 by phosphatidylinositol bisphosphate levels. J Immunol 164, 5398‐5402

246. Golej, D. L., Askari, B., Kramer, F., Barnhart, S., Vivekanandan‐Giri, A., Pennathur, S., and Bornfeldt, K. E. (2011) Long‐chain acyl‐CoA synthetase 4 modulates prostaglandin E(2) release from human arterial smooth muscle cells. J Lipid Res 52, 782‐793

247. Lee, H. C., Inoue, T., Sasaki, J., Kubo, T., Matsuda, S., Nakasaki, Y., Hattori, M., Tanaka, F., Udagawa, O., Kono, N., Itoh, T., Ogiso, H., Taguchi, R., Arita, M., Sasaki, T., and Arai, H. (2012) LPIAT1 regulates arachidonic acid content in phosphatidylinositol and is required for cortical lamination in mice. Mol Biol Cell 23, 4689‐4700

248. Anderson, K. E., Kielkowska, A., Durrant, T. N., Juvin, V., Clark, J., Stephens, L. R., and Hawkins, P. T. (2013) Lysophosphatidylinositol‐acyltransferase‐1 (LPIAT1) is required to maintain physiological levels of PtdIns and PtdInsP(2) in the mouse. PLoS One 8, e58425

249. Gijon, M. A., and Leslie, C. C. (1999) Regulation of arachidonic acid release and cytosolic phospholipase A2 activation. J Leukoc Biol 65, 330‐336

250. Reisenberg, M., Singh, P. K., Williams, G., and Doherty, P. (2012) The diacylglycerol lipases: structure, regulation and roles in and beyond endocannabinoid signalling. Philos Trans R Soc Lond B Biol Sci 367, 3264‐3275

251. Hammond, V. J., and O'Donnell, V. B. (2012) Esterified eicosanoids: generation, characterization and function. Biochim Biophys Acta 1818, 2403‐2412

252. Junger, W. G. (2011) Immune cell regulation by autocrine purinergic signalling. Nat Rev Immunol 11, 201‐212

253. Subramanian, P., Vora, M., Gentile, L. B., Stahelin, R. V., and Chalfant, C. E. (2007) Anionic lipids activate group IVA cytosolic phospholipase A2 via distinct and separate mechanisms. J Lipid Res 48, 2701‐2708

254. Andreyev, A. Y., Fahy, E., Guan, Z., Kelly, S., Li, X., McDonald, J. G., Milne, S., Myers, D., Park, H., Ryan, A., Thompson, B. M., Wang, E., Zhao, Y., Brown, H. A., Merrill, A. H., Raetz, C. R., Russell, D. W., Subramaniam, S., and Dennis, E. A. (2010) Subcellular organelle lipidomics in TLR‐4‐activated macrophages. J Lipid Res 51, 2785‐2797

193

255. Coleman, R. A., Lewin, T. M., Van Horn, C. G., and Gonzalez‐Baro, M. R. (2002) Do long‐chain acyl‐CoA synthetases regulate fatty acid entry into synthetic versus degradative pathways? J Nutr 132, 2123‐2126

256. Watkins, P. A. (2008) Very‐long‐chain acyl‐CoA synthetases. J Biol Chem 283, 1773‐ 1777

257. Tong, F., Black, P. N., Coleman, R. A., and DiRusso, C. C. (2006) Fatty acid transport by vectorial acylation in mammals: roles played by different isoforms of rat long‐ chain acyl‐CoA synthetases. Arch Biochem Biophys 447, 46‐52

258. Lewin, T. M., Kim, J. H., Granger, D. A., Vance, J. E., and Coleman, R. A. (2001) Acyl‐ CoA synthetase isoforms 1, 4, and 5 are present in different subcellular membranes in rat liver and can be inhibited independently. J Biol Chem 276, 24674‐24679

259. Ellis, J. M., Mentock, S. M., Depetrillo, M. A., Koves, T. R., Sen, S., Watkins, S. M., Muoio, D. M., Cline, G. W., Taegtmeyer, H., Shulman, G. I., Willis, M. S., and Coleman, R. A. (2011) Mouse cardiac acyl coenzyme a synthetase 1 deficiency impairs Fatty Acid oxidation and induces cardiac hypertrophy. Mol Cell Biol 31, 1252‐1262

260. Durgan, D. J., Smith, J. K., Hotze, M. A., Egbejimi, O., Cuthbert, K. D., Zaha, V. G., Dyck, J. R., Abel, E. D., and Young, M. E. (2006) Distinct transcriptional regulation of long‐ chain acyl‐CoA synthetase isoforms and cytosolic thioesterase 1 in the rodent heart by fatty acids and insulin. Am J Physiol Heart Circ Physiol 290, H2480‐2497

261. Doenst, T., Nguyen, T. D., and Abel, E. D. (2013) Cardiac metabolism in heart failure: implications beyond ATP production. Circ Res 113, 709‐724

262. Daum, G., and Vance, J. E. (1997) Import of lipids into mitochondria. Prog Lipid Res 36, 103‐130

263. Smaal, E. B., Romijn, D., Geurts van Kessel, W. S., de Kruijff, B., and de Gier, J. (1985) Isolation and purification of cardiolipin from beef heart. J Lipid Res 26, 634‐637

264. Schlame, M., Brody, S., and Hostetler, K. Y. (1993) Mitochondrial cardiolipin in diverse eukaryotes. Comparison of biosynthetic reactions and molecular acyl species. Eur J Biochem 212, 727‐735

265. Schlame, M., and Hostetler, K. Y. (1997) Cardiolipin synthase from mammalian mitochondria. Biochim Biophys Acta 1348, 207‐213

266. Haines, T. H., and Dencher, N. A. (2002) Cardiolipin: a proton trap for oxidative phosphorylation. FEBS Lett 528, 35‐39

267. Paradies, G., Ruggiero, F. M., Petrosillo, G., and Quagliariello, E. (1998) Peroxidative damage to cardiac mitochondria: cytochrome oxidase and cardiolipin alterations. FEBS Lett 424, 155‐158

268. Bione, S., D'Adamo, P., Maestrini, E., Gedeon, A. K., Bolhuis, P. A., and Toniolo, D. (1996) A novel X‐linked gene, G4.5. is responsible for Barth syndrome. Nat Genet 12, 385‐389

194

269. Vreken, P., Valianpour, F., Nijtmans, L. G., Grivell, L. A., Plecko, B., Wanders, R. J., and Barth, P. G. (2000) Defective remodeling of cardiolipin and phosphatidylglycerol in Barth syndrome. Biochem Biophys Res Commun 279, 378‐382

270. Xu, Y., Malhotra, A., Ren, M., and Schlame, M. (2006) The enzymatic function of tafazzin. J Biol Chem 281, 39217‐39224

271. Barth, P. G., Scholte, H. R., Berden, J. A., Van der Klei‐Van Moorsel, J. M., Luyt‐ Houwen, I. E., Van 't Veer‐Korthof, E. T., Van der Harten, J. J., and Sobotka‐Plojhar, M. A. (1983) An X‐linked mitochondrial disease affecting cardiac muscle, skeletal muscle and neutrophil leucocytes. J Neurol Sci 62, 327‐355

272. Saini‐Chohan, H. K., Holmes, M. G., Chicco, A. J., Taylor, W. A., Moore, R. L., McCune, S. A., Hickson‐Bick, D. L., Hatch, G. M., and Sparagna, G. C. (2009) Cardiolipin biosynthesis and remodeling enzymes are altered during development of heart failure. J Lipid Res 50, 1600‐1608

273. Chicco, A. J., Sparagna, G. C., McCune, S. A., Johnson, C. A., Murphy, R. C., Bolden, D. A., Rees, M. L., Gardner, R. T., and Moore, R. L. (2008) Linoleate‐rich high‐fat diet decreases mortality in hypertensive heart failure rats compared with lard and low‐ fat diets. Hypertension 52, 549‐555

274. Mulligan, C. M., Sparagna, G. C., Le, C. H., De Mooy, A. B., Routh, M. A., Holmes, M. G., Hickson‐Bick, D. L., Zarini, S., Murphy, R. C., Xu, F. Y., Hatch, G. M., McCune, S. A., Moore, R. L., and Chicco, A. J. (2012) Dietary linoleate preserves cardiolipin and attenuates mitochondrial dysfunction in the failing rat heart. Cardiovasc Res 94, 460‐468

275. Berry, K. A., Li, B., Reynolds, S. D., Barkley, R. M., Gijon, M. A., Hankin, J. A., Henson, P. M., and Murphy, R. C. (2011) MALDI imaging MS of phospholipids in the mouse lung. J Lipid Res 52, 1551‐1560

276. Grevengoed, T. J., Martin, S. A., Katunga, L., Cooper, D. E., Anderson, E. J., Murphy, R. C., and Coleman, R. A. (2015) Acyl‐CoA synthetase 1 deficiency alters cardiolipin species and impairs mitochondrial function. J Lipid Res 56, 1572‐1582

277. Iijima, H., Fujino, T., Minekura, H., Suzuki, H., Kang, M. J., and Yamamoto, T. (1996) Biochemical studies of two rat acyl‐CoA synthetases, ACS1 and ACS2. Eur J Biochem 242, 186‐190

278. Wherrett, J. R., and Huterer, S. (1972) Enrichment of bis‐(monoacylglyceryl) phosphate in lysosomes from rat liver. J Biol Chem 247, 4114‐4120

279. Matsuzawa, Y., Poorthuis, B. J., and Hostetler, K. Y. (1978) Mechanism of phosphatidylinostiol stimulation of lysosomal bis (monoacylglyceryl)phosphate synthesis. J Biol Chem 253, 6650‐6653

280. Cochran, F. R., Connor, J. R., Roddick, V. L., and Waite, B. M. (1985) Lyso(bis)phosphatidic acid: a novel source of arachidonic acid for oxidative metabolism by rabbit alveolar macrophages. Biochem Biophys Res Commun 130, 800‐806

195

281. Mashek, D. G., Li, L. O., and Coleman, R. A. (2006) Rat long‐chain acyl‐CoA synthetase mRNA, protein, and activity vary in tissue distribution and in response to diet. J Lipid Res 47, 2004‐2010

282. Le, C. H., Mulligan, C. M., Routh, M. A., Bouma, G. J., Frye, M. A., Jeckel, K. M., Sparagna, G. C., Lynch, J. M., Moore, R. L., McCune, S. A., Bristow, M., Zarini, S., Murphy, R. C., and Chicco, A. J. (2014) Delta‐6‐desaturase links polyunsaturated fatty acid metabolism with phospholipid remodeling and disease progression in heart failure. Circ Heart Fail 7, 172‐183

283. Duda, M. K., O'Shea, K. M., Lei, B., Barrows, B. R., Azimzadeh, A. M., McElfresh, T. E., Hoit, B. D., Kop, W. J., and Stanley, W. C. (2007) Dietary supplementation with omega‐3 PUFA increases adiponectin and attenuates ventricular remodeling and dysfunction with pressure overload. Cardiovasc Res 76, 303‐310

284. Duda, M. K., O'Shea, K. M., Tintinu, A., Xu, W., Khairallah, R. J., Barrows, B. R., Chess, D. J., Azimzadeh, A. M., Harris, W. S., Sharov, V. G., Sabbah, H. N., and Stanley, W. C. (2009) Fish oil, but not flaxseed oil, decreases inflammation and prevents pressure overload‐induced cardiac dysfunction. Cardiovasc Res 81, 319‐327

285. Shah, K. B., Duda, M. K., O'Shea, K. M., Sparagna, G. C., Chess, D. J., Khairallah, R. J., Robillard‐Frayne, I., Xu, W., Murphy, R. C., Des Rosiers, C., and Stanley, W. C. (2009) The cardioprotective effects of fish oil during pressure overload are blocked by high fat intake: role of cardiac phospholipid remodeling. Hypertension 54, 605‐611

286. Li, L. O., Ellis, J. M., Paich, H. A., Wang, S., Gong, N., Altshuller, G., Thresher, R. J., Koves, T. R., Watkins, S. M., Muoio, D. M., Cline, G. W., Shulman, G. I., and Coleman, R. A. (2009) Liver‐specific loss of long chain acyl‐CoA synthetase‐1 decreases triacylglycerol synthesis and beta‐oxidation and alters phospholipid fatty acid composition. J Biol Chem 284, 27816‐27826

287. Majerus, P. W., Ross, T. S., Cunningham, T. W., Caldwell, K. K., Jefferson, A. B., and Bansal, V. S. (1990) Recent insights in phosphatidylinositol signaling. Cell 63, 459‐ 465

288. Hille, B. (2012) Diversity of phosphoinositide signaling. Biochemistry (Moscow) Supplement Series A: Membrane and Cell Biology 6, 113‐119

289. Somlyo, A. P., and Somlyo, A. V. (1994) Signal transduction and regulation in smooth muscle. Nature 372, 231‐236

290. Thomas, R. L., and Gustafsson, A. B. (2013) Mitochondrial autophagy‐‐an essential quality control mechanism for myocardial homeostasis. Circ J 77, 2449‐2454

291. Petiot, A., Ogier‐Denis, E., Blommaart, E. F., Meijer, A. J., and Codogno, P. (2000) Distinct classes of phosphatidylinositol 3'‐kinases are involved in signaling pathways that control macroautophagy in HT‐29 cells. J Biol Chem 275, 992‐998

292. Rong, X., Wang, B., Dunham, M. M., Hedde, P. N., Wong, J. S., Gratton, E., Young, S. G., Ford, D. A., and Tontonoz, P. (2015) Lpcat3‐dependent production of arachidonoyl phospholipids is a key determinant of triglyceride secretion. Elife 4

196

293. Kuch, E. M., Vellaramkalayil, R., Zhang, I., Lehnen, D., Brugger, B., Sreemmel, W., Ehehalt, R., Poppelreuther, M., and Fullekrug, J. (2014) Differentially localized acyl‐ CoA synthetase 4 isoenzymes mediate the metabolic channeling of fatty acids towards phosphatidylinositol. Biochim Biophys Acta 1841, 227‐239

294. Askari, B., Kanter, J. E., Sherrid, A. M., Golej, D. L., Bender, A. T., Liu, J., Hsueh, W. A., Beavo, J. A., Coleman, R. A., and Bornfeldt, K. E. (2007) Rosiglitazone inhibits acyl‐ CoA synthetase activity and fatty acid partitioning to diacylglycerol and triacylglycerol via a peroxisome proliferator‐activated receptor‐gamma‐ independent mechanism in human arterial smooth muscle cells and macrophages. Diabetes 56, 1143‐1152

295. Raetz, C. R., Garrett, T. A., Reynolds, C. M., Shaw, W. A., Moore, J. D., Smith, D. C., Jr., Ribeiro, A. A., Murphy, R. C., Ulevitch, R. J., Fearns, C., Reichart, D., Glass, C. K., Benner, C., Subramaniam, S., Harkewicz, R., Bowers‐Gentry, R. C., Buczynski, M. W., Cooper, J. A., Deems, R. A., and Dennis, E. A. (2006) Kdo2‐Lipid A of Escherichia coli, a defined endotoxin that activates macrophages via TLR‐4. J Lipid Res 47, 1097‐ 1111

296. Evans, J., Zamboni, R., Nathaniel, D., Leveille, C., and Ford‐Hutchinson, A. W. (1985) Characterization of biological properties of synthetic and biological leukotriene B3. Prostaglandins 30, 981‐988

297. Evans, J. F., Nathaniel, D. J., Zamboni, R. J., and Ford‐Hutchinson, A. W. (1985) Leukotriene A3. A poor substrate but a potent inhibitor of rat and human neutrophil leukotriene A4 hydrolase. J Biol Chem 260, 10966‐10970

298. Lee, T. H., Sethi, T., Crea, A. E., Peters, W., Arm, J. P., Horton, C. E., Walport, M. J., and Spur, B. W. (1988) Characterization of leukotriene B3: comparison of its biological activities with leukotriene B4 and leukotriene B5 in complement receptor enhancement, lysozyme release and chemotaxis of human neutrophils. Clin Sci (Lond) 74, 467‐475

299. Orning, L., Bernstrom, K., and Hammarstrom, S. (1981) Formation of leukotrienes E3, E4 and E5 in rat basophilic leukemia cells. Eur J Biochem 120, 41‐45

300. Hammarstrom, S. (1981) Conversion of 5,8,11‐eicosatrienoic acid to leukotrienes C3 and D3. J Biol Chem 256, 2275‐2279

197