Phospholipase D: Key Player in Macrophage- Mediated Inflammation and Resolution

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PHOSPHOLIPASE D: KEY PLAYER IN MACROPHAGE- MEDIATED INFLAMMATION AND RESOLUTION

A dissertation submitted in partial fulfillment of the requirements for the degree of Doctor of Philosophy

RAMYA GANESAN B.Tech., Anna University, 2011 M.S., Wright State University, 2014

2017 Wright State University

i WRIGHT STATE UNIVERSITY GRADUATE SCHOOL

Dec 11, 2017

I HEREBY RECOMMEND THAT THE THESIS PREPARED UNDER MY SUPERVISION BY Ramya Ganesan ENTITLED Phospholipase D: Key Player in Macrophage-mediated Inflammation and Resolution BE ACCEPTED IN PARTIAL FULFILLMENT OF THE REQUIREMENTS FOR THE DEGREE OF Doctor of Philosophy.

Committee on Final Examination

Julian G. Cambronero, Ph.D. Professor Julian G. Cambronero, Ph.D. Thesis Director

Nancy J. Bigley, Ph.D. Professor

Juliusz A. Kozak, Ph.D. Associate Professor Mill W. Miller, Ph.D. Director, Biomedical Sciences Ph.D. Program Michael P. Markey, Ph.D. Research Associate Professor

Yong-jie Xu, Ph.D. Associate Professor Barry Milligan, Ph.D. Interim Dean of the Graduate Gerald M. Alter, Ph.D. School Professor

iii ABSTRACT

Ganesan, Ramya Ph.D., Biomedical Sciences Ph.D. program, Wright State University, 2017. Phospholipase D: Key Player in Macrophage-mediated Inflammation and Resolution

Macrophages are central to the inflammatory response and its ability to resolve effectively. They are complex cells that adopt a range of subtypes depending on the tissue type and stimulus. This flexibility allows them to play multiple, sometimes opposing, roles in inflammation and tissue repair. Their central role in the inflammatory process is reflected in macrophage dysfunction being implicated in chronic inflammation and poorly healing wounds. Acute inflammatory response induces an increase in prostaglandins and leukotrienes and leads to chronic inflammation, which is inhibited by resolvins. Additionally, resolvins play a crucial role in wound healing. During inflammation, leukocytes release cytokines, exacerbate inflammation and damage tissues, while neutrophils produce oxygen radicals that worsen the initial inflammation.

Atherosclerosis is an inflammatory disease caused by accumulation of foam cells derived from macrophages on blood vessel walls. It is a significant health problem and a major contributor to cardiovascular disease (CVD), which accounts for one in three deaths in the U.S. (Mozaffarian, Benjamin et al. 2015), and continues to rise globally (Field,

Hazinski et al. 2010, Townsend, Nichols et al. 2015). Deleterious inflammation is a primary feature of breast cancer. Accumulating evidence demonstrates that macrophages, the most abundant leukocyte population in mammary tumors, have a critical role at each stage of cancer progression.

iv Phospholipase D (PLD) is a cell membrane remodeling and signaling protein implicated in the pathology of chronic inflammation. As PLD is also central to macrophage cell migration, we investigated the molecular basis of PLD’s involvement and regulation in macrophage-initiated inflammation (atherosclerosis) and resolution. We have found that PLD is associated with signaling proteins and positively affects cell movement, phagocytosis and NADPH-initiated release of Reactive Oxygen Species

(ROS). We found that PLD2 but not PLD1 is important for foam cell formation that causes atherogenesis. We have also found a novel way of inducing macrophage (MØ) class-switch (polarization) by PLD overexpression. PLD induces a macrophage M1 to

M2 class-switch that accelerates resolution of inflammation and limits damage to blood vessels and affected tissues during atherosclerosis and other inflammatory conditions. We also investigated a new molecular pathway for MØ class-switch (M1-to-M2) by overexpressed PLD resulting in inflammation by bacterial phagocytosis or resolution of inflammation by efferocytosis. In order to understand the physiological relevance of

PLD’s role in inflammation and resolution, we studied the effect of resolvins, a class of specialized proresolving lipid mediators (SPMs), on PLD expression and activity in the different macrophage populations taking into consideration the time course of inflammation and resolution. We found that RvD5 upregulates PLD activity and expression in M2 macrophages confirming a molecular mechanism for PLD’s role in resolution of inflammation.

TABLE OF CONTENTS

INTRODUCTION ...... 1

Phospholipase D...... 1

Inflammation ...... 8

Resolution of Inflammation ...... 12

Resolvins ...... 16

Role of Macrophages in Inflammation ...... 20

Macrophage Phenotypes ...... 21

Atherosclerosis ...... 24

Inflammation in Atherosclerosis ...... 27

Role of PLD2 in Atherosclerosis ...... 28

Inflammation and Resolution Involving Macrophages ...... 29

MATERIALS AND METHODS ...... 31

Materials ...... 31

Animals ...... 31

Ischemia-reperfusion-induced second-organ injury ...... 33

Macrophage and PMN phagocytosis and efferocytosis ...... 33

PLD activity assay ...... 35

Real-time (quantitative) Reverse Transcription-PCR ...... 35

SDS-PAGE and Western blot analyses ...... 36

Myeloperoxidase (MPO) Assay ...... 37

Isolation of bone marrow-derived monocytes ...... 38

vi Differentiation of bone marrow cells into bone marrow-derived macrophages (BMDM)

...... 39

Preparation of aggregated, oxidized LDL particles (Agg-Ox-LDL) ...... 39

Phagocytosis of Agg-Ox-LDL by macrophage foam cells ...... 40

Phagocytosis of Zymosan ...... 41

Immunofluorescence microscopy of phagocytosis and key proteins ...... 41

Human artery specimens ...... 41

Computational transcriptomic analysis ...... 42

PLD inhibitors ...... 43

Co-immunoprecipitation ...... 43

Statistical Analysis ...... 45

RESULTS ...... 46

Chapter I: To study the regulation of Phospholipase D by resolvins in

macrophage mediated inflammation-resolution ...... 46

1.1. PLD activity and expression are altered in anti- and pro-inflammatory

macrophages ...... 46

1.2. Effect of resolvins on PLD and S6 Kinase gene expression ...... 49

1.3. Effect of resolvins on anti- and pro-inflammatory macrophage PLD activity .. 56

1.4. Effect of resolvins on PLD-mediated inflammation resolution ...... 58

1.5. Effect of PLD and resolvins on macrophage functions ...... 61

DISCUSSION ...... 65

Chapter II: To study the role of Phospholipase D in macrophage polarization ... 68

2.1. Effect of PLD on macrophage function ...... 68

vii 2.2. Macrophage polarization in inflammation ...... 71

2.3. PLD affects human M0 macrophage polarization ...... 73

2.4. M1 to M2 macrophage class-switch induced by PLD ...... 75

DISCUSSION ...... 79

Chapter III: To study the role of PLD in macrophage foam cell formation ...... 82

3.1. Macrophage phagocytosis of Agg-Ox-LDL is reduced in the absence of PLD2

activity ...... 82

3.2. PLD inhibitors reduce Agg-Ox-LDL phagocytosis ...... 85

3.4. The heterotrimeric protein complex of PLD2-Grb2-WASp ...... 88

3.5. Arterial gene expression during atherosclerosis ...... 92

3.6. PLD2 but not PLD1 is upregulated in diseased artery tissues ...... 94

3.7. Upregulation of Wiskott-Aldrich Syndrome protein (WASp) and Grb2 in

diseased artery tissues ...... 94

3.8. Validation of PLD2, WASp and Grb2 in diseased human artery samples ...... 97

DISCUSSION ...... 99

CONCLUSIONS ...... 104

LIST OF ABBREVIATIONS ...... 106

REFERENCES ...... 110

viii LIST OF FIGURES

Figure 1. Enzymatic reaction of Phospholipase D (PLD) ...... 3

Figure 2. Classification of PLD ...... 4

Figure 3. PLD signaling in cells ...... 6

Figure 4. Cardinal signs of inflammation ...... 9

Figure 5. Process of inflammation ...... 11

Figure 6. Acute inflammatory response ...... 13

Figure 7. Pillars of resolution of inflammation ...... 15

Figure 8. Structure of Resolvins ...... 18

Figure 9. Macrophage polarization ...... 23

Figure 10. Atherosclerosis in artery ...... 26

Figure 11. PLD activity and expression are altered in anti- and pro-inflammatory

macrophages ...... 48

Figure 12. Effect of Resolvins on PLD gene expression ...... 51

Figure 13. Effect of Resolvins on S6K gene expression ...... 53

Figure 14. Effect of Resolvins on non-classical PLD gene expression ...... 55

Figure 15. Effect of Resolvins on anti- and pro-inflammatory macrophage PLD activity57

ix Figure 16. Resolvin-PLD in Hind-limb ischemia reperfusion injury ...... 60

Figure 17. Effect of PLD and Resolvins on macrophage functions ...... 62

Figure 18. Resolvins affect PLD protein expression in M1 and M2 macrophages ...... 64

Figure 19. Model for Resolvin-PLD signaling in macrophages ...... 67

Figure 20. Cytokine stimulation affects PLD activity and expression ...... 70

Figure 21. Macrophage polarization in inflammation ...... 72

Figure 22. PLD affects human M0 macrophage polarization ...... 74

Figure 23. M1 to 2 macrophage class-switch induced by PLD ...... 76

Figure 24. Effect of PLD on M1 and M2 macrophage functions ...... 78

Figure 25. Model for PLD induced macrophage polarization ...... 81

Figure 26. Macrophage phagocytosis of oxidized LDL is PLD2-dependent ...... 84

Figure 27. Inhibition of PLD reduces LDL phagocytosis ...... 87

Figure 28. The adapter protein Grb2 links PLD2 to Wiskott-Aldrich Syndrome protein

(WASp) ...... 89

Figure 29. PLD2 interacts with Wiskott-Aldrich Syndrome protein (WASp) and actin .. 91

Figure 30. Bioinformatic analysis of PLD and its signaling proteins in human atheroma

plaque tissues ...... 93

Figure 31. PLD2, Grb2 and phospho-WASp are upregulated in diseased artery tissues . 96

x Figure 32. PLD2, WASp and Grb2 are overexpressed in diseased human artery samples

...... 98

Figure 33. Model of PLD2-mediated foam cell atherogenesis ...... 103

Figure 34. Model of PLD2-mediated inflammation and resolution in macrophages ..... 105

xi LIST OF TABLES

Table 1. Specialized Proresolving Lipid Mediators (SPMs) ...... 19

xii ACKNOWLEDGMENTS

I would like to sincerely thank my mentor Dr. Julian Gomez-Cambronero for providing incredible help, guidance and encouragement. I am very grateful for his patience and for his immense support through my journey as a Ph.D. candidate. I am especially thankful to my committee members: Dr. Nancy Bigley, Dr. Gerald Alter, Dr.

Ashot Kozak, Dr. Yong-jie Xu and Dr. Michael Markey for their ideas, feedback and advice during the committee meetings.

I would like to thank the past and present members of Cambronero lab, especially

Karen Henkels (“the lab mom”) for training me, guiding and helping me a lot, both in the lab and outside of work. She is my go-to person. I would also like to thank my previous colleagues and my friends, Dr. Kristen Fite and Taylor Miller for their constant support, intellectual conversations, coffee breaks and fun-times together. I would like to thank my best friend Nagarjuna Reddy Cheemarla for his support, valuable suggestions and guidance during my Ph.D. I would like to extend my gratitude to Dr. Charles Serhan and his lab members at Brigham and Women’s Hospital for their help and freedom to utilize their laboratory resources. I would like to thank Dr. Cambronero and Dr. Alter for giving me the opportunity to join the BMS Ph.D. program.

I am thankful to the BMB department for the support and wonderful research environment. Thanks to Dr. Mill Miller, Dr. Gerald Alter and Karen Luchin, who have worked greatly towards the success of BMS program. Thanks to the BMS program for their travel funds and continuous support that helped me a great deal in learning, gaining experience and networking with other researchers at scientific society meetings.

xiii Earnest thanks to my parents Mr. S. Ganesan and Mrs. M. Jayavijaya for their unconditional love and support. They have always encouraged me to pursue what I like. I am very grateful for their faith in me and I couldn’t have accomplished any of this without their love, care and support. I am very grateful for the tremendous support provided by my husband Srini Viswanathan. I would also like to thank my brother, sister- in-law and my mother-in-law. My warmest thanks to my friends for always being there for me.

Finally, I thank God for giving me the strength and showering me with blessings to strive hard to succeed in life.

xiv INTRODUCTION

Phospholipase D

Phoshpholipase D (PLD) is a lipase enzyme that catalyzes the conversion of

Phosphatidylcholine (PC) into Phosphatidic acid (PA) and Choline. This enzymatic action involves breaking the phospholipid phosphodiester bond (Figure 1). PLD in the presence of butanol converts phosphatidylcholine into phosphatidylbutanol by a process termed transphosphatidylation. This enables measuring PLD activity in vitro using a radioactive assay involving radioactive 3H-butanol to yield 3H-phosphatidylbutanol, which can then be measured in a scintillation counter after extraction of lipids and separation by chromatography.

Phospholipase D belongs to the HKD superfamily, as it possesses a

“HxxKxxxDx” motif in its catalytic domain (Rudolph, Stuckey et al. 1999). The two major mammalian isoforms of PLD are PLD1 and PLD2, that have phosphoinositide binding (PX) and pleckstrin homology (PH) domains with 2 HKD motifs, while the non- classical isoforms of PLD such as PLD3, 4 and 5 do not possess the HKD motif and hence lack the lipase activity (Figure 2).

PLD1 gene is on chromosome 3q.26 (human), which encodes a protein of

120KDa. The gene encoding PLD2 is on chromosome 17p.13, encoding a protein of

105KDa. PLD1 protein is bigger than PLD2 because of the activation loop. The subcellular localizations of PLD1 and PLD2 are very different. PLD1 is present mainly in

1 the ER and golgi, while PLD2 is predominantly found in the plasma membrane and cytosol.

Phospholipase D (PLD) has a pivotal role in cellular activities such as chemotaxis, phagocytosis, neurite extension and retraction and cell growth based on their ability to regulate actin cytoskeletal reorganization, cell cycle progression and gene transcription. Phospholipase D2 (PLD2) is known for its role in signaling due to its lipase or guanine-nucleotide exchange factor (GEF) activities and protein-protein interactions

(Mahankali, Peng et al. 2011, Mahankali, Henkels et al. 2012, Mahankali, Henkels et al.

2013, Gomez-Cambronero 2014, Gomez-Cambronero and Kantonen 2014, Gomez-

Cambronero 2016).

Figure 1. Enzymatic reaction of Phospholipase D (PLD) This is a schematic of the catalytic conversion of phosphatidylcholine (PC) into phosphatidic acid (PA) and choline by PLD.

Figure 2. Classification of PLD PLD is classified into classical and non-classical PLD based on HKD motif and regulatory domains (PX and PH).

4 Phosphatidic acid (PA) produced by PLD, is an important second messenger and cell signaling molecule (Kooijman and Burger 2009). PA initiates signal cascade by binding to S6 kinase (S6K) leading to actin polymerization and chemotaxis or cell migration. PA also interacts with other proteins such as mTOR, Sos (Zhao, Du et al.

2007), Rac and Ras (Peng, Henkels et al. 2011) resulting in actin polymerization and thus cell migration in both normal and cancer cells. PA also activates p70S6 kinase independent of mTOR, resulting in phosphorylation of ribosomal S6 protein resulting in protein synthesis (Fang, Vilella-Bach et al. 2001). Both PLD1 and PLD2 regulate macrophage phagocytosis important for innate immune functions (Iyer, Barton et al.

2004, Iyer, Agrawal et al. 2006) (Figure 3).

Additionally, aberrant PLD expression and activity is implicated in inflammation

(Iyer, Barton et al. 2004, Iyer, Agrawal et al. 2006, Park, Lee et al. 2013, Schonberger,

Jurgens et al. 2014, Mancini and Ciervo 2015) such as in atherosclerosis, infectious diseases, autoimmune disease and ischemia/reperfusion injury (I/RI). PLD is important for the phagocytic activity of macrophages and other cellular functions, as well as during inflammation (Gomez-Cambronero, Di Fulvio et al. 2007, Gomez-Cambronero 2014).

PLD is also actively involved in pro-inflammatory cytokine recruitment, reactive oxygen species (ROS) generation, chemotaxis and cell invasion (Koch, Seifert et al. 2009,

Usatyuk, Gorshkova et al. 2009, Usatyuk, Kotha et al. 2013).

Figure 3. PLD signaling in cells This is a schematic showing PLD and its product PA initiating signaling cascade within a cell for various physiological functions. Adapted from (Gomez-Cambronero 2011).

6 Studies have recently shown that PLD is activated by both tyrosine kinase receptors and

G-protein coupled receptors. As such, PLD is essential for FPR activity and a peptide that activates FPR during inflammation results in PLD activation and phagocytic activity in dendritic cells (Lee, Kang et al. 2004). Another class of specialized pro-resolving lipid mediators, called lipoxins, inhibits PLD via hindering presqualene diphosphate (Levy,

Fokin et al. 1999).

Second organ injury refers to the damage of distant organs, such as the lungs, liver, spleen and kidneys, as a result of ischemia/reperfusion injury (I/RI), which occurs as a result of release of reactive oxygen species (ROS) and other signaling molecules from the ischemic tissue. PLD activation following H2O2 exposure results in NADPH oxidase activation (Madesh, Ibrahim et al. 1997). Cell survival is affected when the H2O2 attack continues unchecked, which then contributes to accumulation of intracellular phosphatidic acid (PA), the product of the PLD reaction. During ischemia, depletion of

ATP and high intracellular Ca2+ results in damage of the electron transport chain and high mitochondrial reactive oxygen species (ROS) synthesis; once oxygen is restored, cell death and release of harmful ROS into the circulation is further elevated (Murphy and

Steenbergen 2008). Studies have also shown an isoform dependent role of PLD in inflammatory response to retinal pigment epithelium by promoting ERK1/2 and COX activation (Mateos, Kamerbeek et al. 2014). It has been shown that PLD1 is a major mediator of peritonitis via TNF-α activation (Sethu, Pushparaj et al. 2010) and that it is a peripheral blood marker for pancreatitis (Bluth, Lin et al. 2008). PLD1 plays a crucial role in pro-inflammatory cytokine IL-1β mediated activation of synoviocyte and chronic inflammatory arthritis (Kang, Park et al. 2013).

7 Inflammation

Inflammation is a response that is initiated by harmful stimuli, including infection and tissue injury. Infection is initially recognized by tissue-resident macrophages and mast cells, leading to the production and release of several inflammatory mediators, including chemokines, cytokines, vasoactive amines, eicosanoids and products of proteolytic cascades (Medzhitov 2008). The cardinal signs of inflammation are heat, swelling, redness, pain and loss of function (Lawrence, Willoughby et al. 2002) (Figure

4). An immediate effect of these mediators is to elicit an inflammatory exudate locally, such as plasma proteins and leukocytes (mainly neutrophils and monocytes) that are restricted to the blood vessels now access the extravascular tissues at the site of infection

(or injury), through the postcapillary venules (Perkins, Nathani et al. 2007). Neutrophils and macrophages are the first line of defense. Neutrophils are activated by direct contact with inflammatory mediators at the site of inflammation or by several pro-inflammatory cytokines that travel in bloodstream. These activated neutrophils then secrete several inflammatory molecules like cytokines, reactive oxygen species, reactive nitrogen species and elastase (Nathan 2006). Macrophages are key players in inflammatory diseases and are the phagocytic cells involved in the clearance of bacteria during the inflammatory phase and later phagocytose apoptotic neutrophils after an injury (“resolution phase”).

Macrophages undergo a change in their lipid profile from pro-inflammatory to anti- inflammatory/pro-resolution following differentiation from Mzero to M1 and to M2.

(Serhan, Chiang et al. 2008).

Figure 4. Cardinal signs of inflammation Adapted from (Lawrence, Willoughby et al. 2002).

9 There are seven major mediators and effectors including vasoactive amines

(serotonin and histamine), vasoactive peptides (substance P, fibrinogen and fibrin), complement fragments (C3a, C4a, C5a), lipid mediators (prostaglandins and leukotrienes), inflammatory cytokines (TNF-α, IL-1, IL-6, etc.), chemokines (IL-8) and proteolytic enzymes (elastin, MMPs and cathepsins) (Higgs, Moncada et al. 1984, Barton

2008, Medzhitov 2008).

Inflammation is classified as acute and chronic. Initially after an inflammatory insult, local cytokine release, recruitment of leukocytes and platelets, followed by lowered proinflammatory mediators and increased antagonists resulting in homeostasis.

This phase of inflammatory response is known as acute inflammation.

If homeostasis is not restored, there occurs a systemic reaction with uncontrolled cytokine release, activation of humoral cascades and loss of circulatory integrity leading to organ dysfunction. This systemic response is called chronic inflammation.

The sequence of events in an inflammatory response are as follows:

1. cytokine release

2. margination

3. diapedesis

4. chemotaxis

5. phagocytosis (Figure 5); and

6. efferocytosis

Figure 5. Process of inflammation This image shows the sequence of events that occurs in an inflammatory response. ©

Tata McGraw-Hill Companies Inc.

11 Resolution of Inflammation

A successful acute inflammatory response results in the elimination of the infectious agents followed by a resolution and repair phase, which is mediated mainly by tissue-resident and recruited macrophages (Serhan and Savill 2005). Resolution, the return to normal inflammatory conditions, does not merely consist of catabolism of inflammatory mediators and abrogation of inflammatory processes. Instead, resolution is an active and coordinated anti-inflammatory program aimed at restoration of tissue homeostasis, integrity and function (Ortega-Gomez, Perretti et al. 2013). During an acute inflammatory response, neutrophils are recruited releasing pro-inflammatory mediators, which then recruit monocytes and differentiate them to pro-inflammatory macrophages in the tissue. This results in increased pro-inflammatory mediator release including prostaglandins and leukotrienes. Later in inflammation, there occurs a class-switch of not only the cytokines and macrophages, but also the lipid mediators. This lipid mediator class-switch results in increased specialized pro-resolving mediators such as lipoxins, resolvins, protectins and maresins (Serhan 2014) (Figure 6). These SPMs are derived from omega 3 or omega 6 polyunsaturated fatty acids such as arachidonic acid (AA), docosahexanoic acid (DHA) and eicosapentanoic acid (EPA). Resolvins and protectins, which constitute a class of lipid mediators, as well as transforming growth factor-β and growth factors produced by macrophages, also have a crucial role in the resolution of inflammation, including the initiation of tissue repair (Serhan and Savill 2005, Serhan,

Brain et al. 2007).

Figure 6. Acute inflammatory response This image shows the scheme of events that happen during an acute inflammatory response. Highlights the lipid mediator class-switch from pro-inflammatory to pro- resolving facilitating inflammation resolution. Adapted from (Serhan 2014).

13 During the short or acute inflammatory response period (0-4 h), SPMs promote phagocytosis via activation of S6Kinase that phosphorylates ribosomal S6 protein resulting in phagosome formation and hence phagocytosis (Ohira, Arita et al. 2010).

SPMs regulate the immune system by controlling functions of specific cell types.

Resolvins and protectins have been shown to stimulate innate killing mechanisms to manage bacterial loads and stimulate clearance of bacteria (Serhan, Chiang et al. 2008).

Similar to the five cardinal signs of inflammation, there are five cardinal signs of resolution such as removal of microbes and dead cells, restoration of vascular integrity, regeneration of tissue, remission of fever and relief of pain (Basil and Levy 2016) (Figure

7).

Figure 7. Pillars of resolution of inflammation An image showing the five cardinal signs or steps in resolution of inflammation. Adapted from (Basil and Levy 2016).

15 Resolvins

Resolvins are specialized pro-resolving lipid mediators that are derived from omega-3 and omega-6 essential fatty acids docosahexanoic acid (DHA) and eicosapentanoic acid (EPA) (Serhan, Clish et al. 2000, Serhan, Hong et al. 2002) (Figure

8). Resolvins are known to have both anti-inflammatory and pro-resolution activities.

Resolvins have been found to act via specific GPCR in resolving inflammation and preventing infiltration of monocytes and neutrophils to the site of inflammation

(Devchand, Arita et al. 2003, Krishnamoorthy, Recchiuti et al. 2012). Also another group of lipid mediators called lipoxins are known to act by inhibiting release of prostaglandins and leukotrienes (Serhan, Hamberg et al. 1984, Serhan, Hamberg et al. 1984). Resolvins are divided into two classes based on their precursor fatty acids as D-series resolvins denoted RvD1-5- derived from DHA; and E-series resolvins denoted RvE1-2- derived from EPA (Buckley, Gilroy et al. 2014) (Table 1).

Resolvins like the other class of specialized proresolving lipid mediators called lipoxins act via G-protein coupled receptors namely ALX/FPR2 and GPR32 (Devchand,

Arita et al. 2003, Krishnamoorthy, Recchiuti et al. 2012). Resolvins exhibit a time dependent response to inflammation resolution. During the short or acute inflammatory response period (0-4h), resolvins promote phagocytosis via activation of S6Kinase that phosphorylates ribosomal S6 protein resulting in phagosome formation and hence phagocytosis (Ohira, Arita et al. 2010).

Resolvins regulate the immune system by controlling functions of specific cell types. For instance, RvD1 differentially modulates primary human macrophage responses

16 to lipopolysaccharides, depending on the context in which this molecule is presented to the macrophage (Palmer, Mancuso et al. 2011). Resolvins and protectins have been shown to stimulate innate killing mechanisms to manage bacterial loads and stimulate clearance of bacteria (Serhan, Chiang et al. 2008). RvE1 is a potent inhibitor of leukocyte infiltration, dendritic cell migration, IL-12 production and PMN transendothelial migration (Arita, Clish et al. 2005, Campbell, Louis et al. 2007). Furthermore, RvE1 was found to negatively regulate the development of an allergic inflammation in vivo (Aoki,

Hisada et al. 2008). Other studies demonstrated that RvE2 stimulates host-protective actions throughout initiation and resolution of the innate immune responses (Oh, Dona et al. 2012). Additionally, RvE3 has proven to be a potent inhibitor of PMN chemotaxis in vitro and in vivo (Isobe, Arita et al. 2012). Recently, it was demonstrated that in E. coli infections, the combination of RvD1, RvD5 and protectin D1 (a dihydroxy product formed in inflammatory exudates), together with antibiotics, increased antimicrobial responses in mouse peritoneum (Chiang, Fredman et al. 2012). RvD1 promotes tissue repair in salivary glands and helps in restoring gland dysfunction (Odusanwo,

Chinthamani et al. 2012). RvD5 and LxB4 were identified in arterial walls in rabbit arterial angioplasty and found to be important for smooth muscle proliferation and leukocyte recruitment in arteries (Miyahara, Runge et al. 2013). RvD1 promotes macrophage phagocytosis and reduces reactive oxygen species (ROS) production (Titos,

Rius et al. 2011). RvD1promotes wound healing and also increased M2 macrophage markers, important for anti-inflammation and resolution (Hellmann, Tang et al. 2011,

Tang, Zhang et al. 2013). RvD3 and AT-RvD3 act via GPR32 in promoting mibrocial uptake by macrophages (Dalli, Winkler et al. 2013).

Figure 8. Structure of Resolvins This figure shows the structure of E-series and D-series reolvins. Adapted from (Serhan

2017).

Table 1. Specialized Proresolving Lipid Mediators (SPMs) This table shows the families if SPMs, their precursors, target cells and specific action.

Adapted from (Buckley, Gilroy et al. 2014).

19 The studies stated above show that resolvins prevent undue inflammatory reactions and promote resolution of inflammation by the following:

(i) inhibiting cytokine production;

(ii) impeding neutrophil transendothelial migration; and

(iii) increasing macrophage activity resulting in the efferocytosis of apoptotic cells and cellular debris from inflamed regions.

Role of Macrophages in Inflammation

Macrophages are phagocytic leukocytes critical to immune system function and play a critical role in maintaining tissue homeostasis (Gordon and Taylor 2005, Wynn and Barron 2010, Bashir, Sharma et al. 2016). Macrophages originate from bone marrow- derived monocytes and are long-lived cells that reside in tissues (Bashir, Sharma et al.

2016). Macrophages respond to numerous stimuli representing tissue stress, including infection, hypoxia, and metabolic stress (Lecca, Trincavelli et al. 2008, Rausch, Weisberg et al. 2008, Zhou, Huang et al. 2014, Tan, Wang et al. 2016) or may also be self- stimulated (Biswas and Mantovani 2010, Tan, Wang et al. 2016). Macrophages are the first line of defense in the human body immediately after an inflammatory insult. Their diverse functions, including cytokine production and phagocytosis, place macrophages at the balance of inflammation and resolution of inflammation, both necessary components of the healing process (Wynn and Barron 2010, Bashir, Sharma et al. 2016). As such, macrophages are important to inflammation in response to tissue injury or infection, as well as the removal of debris from damaged tissue.

20 Macrophage Phenotypes

Macrophages have multiple phenotypes that mediate different stages of tissue healing (Gordon and Taylor 2005, Wynn and Barron 2010). Mzero macrophages are mature cells that reside in tissues normally. Macrophages may polarize into M1 or M2 phenotypes (Johnson, Milner et al. 2012) (Figure 9). M1 cells are regarded as generally pro-inflammatory and have a high microbicidal capacity and secrete pro-inflammatory cytokines, such as interleukin-1β (IL-1β), interleukin-12 (IL-12) and tumor necrosis factor α (TNF-α) (Tan, Wang et al. 2016). In contrast, M2 cells favor resolution of inflammation by secreting interleukin-10 (IL-10) and transforming growth factor β (TGF-

β) (Tan, Wang et al. 2016). While both cell populations exist normally during homeostasis in healthy tissues, disruption of this balance results in various pathologies

(Wynn and Barron 2010, Braga, Agudelo et al. 2015, Tan, Wang et al. 2016). Pro- inflammatory signals, such as toll-like receptor (TLR) ligands and interferon γ (IFNγ), induce polarization to the M1 phenotype, while anti-inflammatory signals, such as interleukin-4 (IL-4) and IL-10, induce polarization to the M2 phenotype (Zhou, Huang et al. 2014). Thus, macrophage class switch is critical to proper macrophage function and tissue homeostasis and disruption of the class switch results in further damage to tissues.

Examples of macrophages in pathology include ischemia/reperfusion injury, cancer tumor microenvironment, obesity and conditions of chronic inflammation (Dandekar,

Kingaonkar et al. 2011, Tan, Wang et al. 2016).

Phospholipase D (PLD) is a ubiquitously expressed lipase that hydrolyzes phosphatidylcholine (PC) into free choline and phosphatidic acid (PA) (Foster, Salloum et al. 2014, Gomez-Cambronero 2014, Nelson and Frohman 2015). PLD has been shown

21 to be a stress response protein and is upregulated in response to various cell stressors, such as hypoxia and nutrient starvation (Schonberger, Jurgens et al. 2014, Fite,

Elkhadragy et al. 2016). Moreover, PLD2-/- mice produced less pro-inflammatory cytokines in a sepsis model (Lee, Kim et al. 2015). The product of the PLD reaction, PA, is itself a mitogen and critical secondary messenger that activates many downstream pathways leading to cell growth and proliferation, vesicle trafficking and cell migration

(Foster, Salloum et al. 2014, Gomez-Cambronero 2014, Nelson and Frohman 2015).

Additionally, PA is conical in shape and carries a negative charge, meaning its accumulation in the membrane results in membrane curvature. Our lab and others have implicated PLD and PA in cell migration in various cell types, including leukocytes

(Schonberger, Jurgens et al. 2014, Speranza, Mahankali et al. 2014) and cancer cells

(Henkels, Boivin et al. 2013, Fite and Gomez-Cambronero 2016, Xiao, Sun et al. 2016).

Additionally, it has been demonstrated that PLD/PA plays a role in leukocyte adhesion

(Gobel, Schuhmann et al. 2014, Speranza, Mahankali et al. 2014). PLD and its catalytic product PA play a role in inflammation. It would be novel to determine if PLD has a role in macrophage polarization.

Figure 9. Macrophage polarization This figure shows M0 macrophage polarization by Th1 and Th2 cytokine stimulation into either M1 (pro-inflammatory) or M2 (anti-inflammatory) macrophages. Adapted from

(Johnson, Milner et al. 2012).

Atherosclerosis

Atherosclerosis is an inflammatory disease of the vasculature that leads to the development of atheromatous plaques within vessel walls (Paquissi 2016). A key mechanism in plaque development involves migration and proliferation of vascular smooth muscle cells and macrophages. Phagocytosis, most commonly associated with cells of the innate immune system such as macrophages, is used to engulf and eliminate pathogens and damaged cells. Exaggerated macrophage phagocytosis of accumulating lipids in arterial walls leads to the formation and accumulation of foam cells (Figure 10).

The phagocytosis proceeds through both receptor-dependent and -independent uptake of low density lipoprotein (LDL) (Johnson and Newby 2009). LDL-laden foam cell formation in atheromatous plaques narrows arterial lumens, which can partially or completely restrict blood flow to the heart, brain (stroke) and limbs (peripheral arterial disease (PAD)). Ruptured plaques can also trigger strokes through the delivery of cellular-debris emboli to the brain.

Several cells are involved in atherosclerotic plaque formation including endothelial cells, macrophages, foam cells, T lymphocytes, platelets and some neutrophils (Libby, Sukhova et al. 1997). These cells release pro-inflammatory cytokines and growth factors promoting VSMC proliferation and plaque fibrous cap stability

(Libby, Sukhova et al. 1997). Atherosclerotic plaques comprise a variety of cells, including foam cells, macrophages, endothelial cells, platelets and proliferating or apoptotic VSMCs. Fibrous cap stability is important as cap break-off results in plaque rupture, which loosely circulates blocking blood vessels and results in accumulated

24 platelets that cause stroke or myocardial infarction (Badimon, Padro et al. 2012). Hence, promoting proliferation of VSMCs is key to atherosclerosis.

Oxidatively-modified LDL or oxidized LDL (ox-LDL), consists of protein components that have been modified by aldehyde products that create net negative charges that render the ox-LDL very attractive to macrophages in terms of uptake

(phagocytosis) specially when aggregated in large particles as Agg-Ox-LDL

(Parthasarathy, Raghavamenon et al. 2010) and are prevalent in atheromatous plaques

(Aviram, Maor et al. 1995, Maor, Hayek et al. 2000, Zhao, Li et al. 2006, Waldo, Li et al.

2008). Monocytes differentiated into macrophages with M-CSF phagocytose aggregated, oxidized LDL (Agg-Ox-LDL) in large quantities (Aviram, Maor et al. 1995, Maor,

Hayek et al. 2000, Zhao, Li et al. 2006). This process is more prevalent in the arterial wall due to macrophage-secreted proteoglycan (Schrijvers, De Meyer et al. 2007).

Macrophage phagocytosis requires Agg-Ox-LDL and neutralization of the CD36 receptor, which is a distinct process from that of CD36 receptor-mediated uptake of LDL

(Podrez, Febbraio et al. 2000). The molecular and genetic mechanisms that govern macrophage phagocytosis of LDL and hasten inflammation are still unknown in terms of vascular biology. Until we understand these processes, therapeutic or surgical approaches used to disrupt/remove plaque will have suboptimal long-term outcomes, as the process of macrophage/foam cell formation will restart.

Figure 10. Atherosclerosis in artery This is a figure showing a normal artery (left) compared to atherosclerotic plaque containing artery (right). The atherosclerotic plaque consists of various cells, damaged endothelium, lipids, calcium, debris and fibrous cap.

©Encyclopedia Britannica 2007.

26 Inflammation in Atherosclerosis

An underlying, chronic inflammation is responsible for the development and continued progression of atherosclerosis (Nabel and Braunwald 2012), which is a significant health problem and a major contributor to cardiovascular disease (CVD).

Atherosclerosis accounts for one in three deaths in the U.S. (Mozaffarian, Benjamin et al.

2015), continues to rise globally (Field, Hazinski et al. 2010, Townsend, Nichols et al.

2015, WHO 2016), and is a major contributor to current healthcare costs (Mozaffarian,

Benjamin et al. 2015). Thus, it is very important to limit or terminate inflammation and start vascular healing early on. Plaque growth and swelling in a diseased artery involves migration of immune cells and the conversion of macrophages into foam cells (Ross

1999). Accumulation of ox-LDL in macrophage-derived foam cells directly affects foam cell migration and increases focal adhesion kinases (FAK), which drives foam cell accumulation in the arterial intima leading to atheromatous plaque formation (Park,

Febbraio et al. 2009).

FAK signaling pathways heavily implicated in inflammation, cell migratory processes, and macrophage podosome and phagocytic cup formation include

Phospholipase D (PLD), Wiskott-Aldrich Syndrome protein (WASp) and actin polymerization (Serrels, Serrels et al. 2007, Murphy and Courtneidge 2011, Eleniste and

Bruzzaniti 2012, Schober and Siess 2012, Tang, Li et al. 2013, Mahankali, Henkels et al.

2015). PLD catalyzes the breakdown of the phospholipid phosphatidylcholine into phosphatidic acid (PA) and choline (McDermott, Wakelam et al. 2004, Gomez-

Cambronero 2010). PLD plays a key role in neutrophil and macrophage-initiated inflammation in many disease states, including chronic inflammation and cardiovascular

27 disease, due to its involvement in signaling pathways promoting cell growth, proliferation and migration (Bruntz, Lindsley et al. 2014, Gomez-Cambronero and Kantonen 2014,

Frohman 2015).

Studies using PLD knockout (KO) mice or chemical inhibitors have reported that suppressing PLD is “protective” during ischemic stroke (Stegner, Thielmann et al. 2013), hypertension (Olala, Seremwe et al. 2013), thrombosis (Noh, Lim et al. 2010, Thielmann,

Stegner et al. 2012) and myocardial infarction (Elvers, Stegner et al. 2010). WASp has a poly-proline rich region, a polybasic region, and a GTPase Binding Domain and is a key regulator in the formation of phagocytic cups (Kim, Kakalis et al. 2000, Rohatgi, Ho et al. 2000, Thrasher, Burns et al. 2000, Marchand, Kaiser et al. 2001, Thrasher 2002,

Panchal, Kaiser et al. 2003, Tomasevic, Jia et al. 2007). WASp binds to cofilin and the

Arp2/3 complex through its C-terminal activity (Verprolin-Cofilin-Acidic, VCA) region to polymerize actin monomers (G-actin) into F-actin (Zhang, Shi et al. 2002). Previously, our laboratory showed that the Phospholipase D2 isoform (PLD2) has a signaling role in

WASp-mediated macrophage phagocytosis and phagocytic cup formation (Kantonen,

Hatton et al. 2011). Defining a direct protein-protein interaction between PLD2-WASp in foam cells within the plaque is a novel approach that could potentially help to explain excessive phagocytosis of ox-LDL.

Role of PLD2 in Atherosclerosis

Phospholipase D is a lipase enzyme that catalyzes the conversion of phosphatidylcholine (PC) into phosphatidic acid (PA) and choline. Phospholipase D and its product PA are involved in several cellular functions including cell migration, DNA synthesis, cell proliferation, membrane ruffling and phagocytosis (Gomez-Cambronero

28 2014). Studies have shown that PLD is involved in superoxide anion generation

(Usatyuk, Kotha et al. 2013), which is known to be involved in ROS generation and endothelial dysfunction. A potential role of PLD in atherogenesis has been implicated wherein PLD pathway is upregulated and activated by oxidixed LDL (ox-LDL) (Gomez-

Munoz, Martens et al. 2000). Another study showed PLD activation in smooth muscle cells in the presence of ox-LDL (Natarajan, Scribner et al. 1995).

Inflammation and Resolution Involving Macrophages

The most numerous and fastest responders to an initial inflammatory insult are the innate immune cells. The initial neutrophil response that infiltrates the distressed myocardium with 30 min after injury quickly wanes and is replaced by a sustained infiltration of inflammatory monocytes for up to 4 days post-ischemia (Nahrendorf,

Swirski et al. 2007, Jung, Kim et al. 2013, Heidt, Courties et al. 2014). Inflammatory monocytes, which differentiate into M1-like macrophages, pursue proteolysis of pre- existing extracellular matrix and phagocytosis of dying cells. After 4 days, there is a transition to a second, less inflammatory phase (Nahrendorf, Swirski et al. 2007), during which low numbers of patrolling Ly6Clow monocytes are also recruited, and Ly6Chigh monocytes give rise to macrophages with fewer inflammatory functions (Hilgendorf,

Gerhardt et al. 2014). Macrophages now support tissue rebuilding, while crosstalk between stromal cells via vascular endothelial growth factor (VEGF) and transforming growth factor β (TGFβ) supports angiogenesis and production of new extracellular matrix, respectively, in human blood (Tsujioka, Imanishi et al. 2009) and heart (van der

Laan, Ter Horst et al. 2014). If either of these phases is compromised, then heart failure occurs (Panizzi, Swirski et al. 2010). The association between leukocytes and

29 inflammation underscores the putative role of innate immune cells as a therapeutic target

(Weirather, Hofmann et al. 2014), while B cells regulate the migratory patterns of myeloid cells (Zouggari, Ait-Oufella et al. 2013).

Resolution of inflammation is an active interaction between innate and adaptive immunity and macrophages play an important role in this cross-talk to facilitate resolution.

30 MATERIALS AND METHODS

Materials

RAW264.7 mouse macrophages (cat. # TIB-71) and DMEM (cat. # 30-2002) were obtained from ATCC (Manassas, VA, USA). RPMI 1640 with L-glutamine and 25 mM

HEPES (cat. # SH30255.01) and ECL reagent (cat. # RPN2106) were from GE

Healthcare Life Sciences (Logan, UT, USA). Fetal bovine serum (heat inactivated) (cat. #

900-108) and Penicillin/Streptomycin (10,000 units penicillin/10,000 mg/ml streptomycin) (cat. # 400-109) were from Gemini Bio-Products (West Sacramento, CA,

USA). Oxidized LDL (cat. # L34357) was from Life technologies (Carlsbad, CA).

Sterile-filtered Histopaque 1077 (cat. # 10771), sterile-filtered Histopaque 1119 (cat.

#11191) and Oil Red O stain (1-(2,5-dimethyl-4-(2,5-dimethylphenyl) phenyldiazenyl) azonapthalen-2-ol) (cat. # O0625) were from Sigma-Aldrich (St. Louis, MO, USA). 0.5

M EDTA, pH 8.0 (cat. # 15575-038) was from Life Techologies (Carlsbad, CA, USA).

Recombinant mouse M-CSF (cat. # 315-02) was from PeproTech (Rocky Hill, NK,

USA). CD36 blocking antibody (cat. # ab23680) was obtained from Abcam (Cambridge,

MA). Mouse isotype control antibody (cat. # 553476) was obtained from BD Biosciences

(San Diego, CA).

Animals

Bone marrow-derived macrophages (BMDMs) were obtained from male or female c57Bl/6 wild-type (Charles River Laboratories, Charleston, SC, USA). PLD1-/- were generated at Dr. Yasunori Kanaho’s laboratory, University of Tsukuba, Tennodai, Japan

(Chen, Hongu et al. 2012). These PLD1-KO c57BL/6 mice had initially PLD2 in ES with

31 exons 13 removed (Chen, Hongu et al. 2012). PLD2-/- were generated at Dr. Gilbert Di

Paolo’s laboratory, Columbia University (Oliveira, Chan et al. 2010). These PLD2-KO c57BL/6 mice had initially PLD2 in ES with exons 13-15 removed (Oliveira, Chan et al.

2010). Wild type mice were also in the C57Bl/6 background at 6-8 wks of age (weighing

20-25 g) comparable to the KOs. The mice were provided a temperature- and light- controlled environment with unrestricted access to food (laboratory standard rodent diet

5001 (Laboratory Diet, St. Louis, MO, USA)) and water. The mice had veterinary care, were checked ever day, and experiments were performed in accordance with the Wright

State University (WSU) Institutional Animal Care and Use Committee (IACUC) guidelines. Experiments for this manuscript have also followed the National Institutes of

Health guide for the care and use of Laboratory animals (NIH Publications No. 8023, revised 1978).

For hind-limb ischemia reperfusion injury experiments, animals were provided a temperature and light-controlled environment with unrestricted access to food (laboratory standard rodent diet 5001 (Lab Diet, St. Louis, MO, USA)) and water. Experiments were conducted in accordance with the Harvard Medical School Standing Committee on

Animals guidelines for animal care (Protocol 02570).

Transfection

Human macrophages were transfected with 2 µg plasmid DNA, HA-PLD1 or myc-PLD2 or 300 ng shRNA, shPLD1 (cat. # sc-44000-SH) or shPLD2 (cat. # sc-44001-SH)

(Santacruz Biotechnology, Dallas, TX) using JetPEI macrophage transfection reagent

(cat. # 103-05N, Polyplus transfection, NY). RAW 264.7 macrophages were transfected

32 with 2 µg plasmid DNA, HA-PLD1 or myc-PLD2 using Amaxa® Cell Line

Nucleofector® Kit V reagent (cat. # VCA-1003, Lonza, Switzerland).

Ischemia-reperfusion-induced second-organ injury

Mice were anesthetized by i.p. injection of pentobarbital (80 mg/kg, Nembutal sodium solution NDC 0074-3778-04). Hind limb ischemia was induced using rubber band tourniquets tied on each hind limb (Chiang, Gronert et al. 1999). Hind limb ischemia was allowed for 1 h, after which the tourniquets were removed to begin reperfusion. Resolvin

D5 (RvD5) was administered at 0.5 µg/mouse in vehicle (5 µl of ethanol in 120 µl of sterile saline) and compared to vehicle alone or no ischemia reperfusion control. RvD5 was administered via I.V. retro-orbital injection at ~5 min before the start of the reperfusion period. At the end of this reperfusion period (2 h), the mice were euthanized with an overdose of anesthetic and the lungs were quickly harvested, frozen in liquid nitrogen and stored at -80°C or fixed and stored in 4% paraformaydehyde for tissue- sectioning and immunohistochemistry. The right lungs harvested from individual mice were homogenized and centrifuged, and the tissue levels of myeloperoxidase (MPO) in the supernatants were determined using a mouse MPO ELISA (Hycult Biotechnology;

Cell Sciences, Plymouth Meeting, PA, USA). The fixed tissues (left lungs) were sectioned and stained hematoxylin and eosin (H&E) (AML laboratories Inc., FL, USA) to study the lung pathology.

Macrophage and PMN phagocytosis and efferocytosis

Human polymorphonuclear neutrophils (PMN) were isolated by density-gradient Ficoll-

Histopaque from human peripheral blood. Peripheral blood was obtained from healthy human volunteers giving informed consent under protocol # 1999-P-001297 approved by

33 the Partners Human Research Committee. Apoptotic neutrophils were prepared by plating 1 x 107 cells/mL in 5 mL DPBSCa+/Mg+ for 24 h in 100 mm x 20 mm petri dishes.

Macrophages were prepared from peripheral blood leukopak. The pack rich in leukocytes was then used to isolate peripheral blood mononuclear cells (PBMCs) by histopaque density gradient. PBMCs were differentiated into macrophages using GM-CSF

(20 ng/mL) or M-CSF (20 ng/mL) in RPMI culture media (Lonza, Walkersville, MD,

USA) containing 10% FBS (Invitrogen, Carlsbad, CA, USA), 5 mM L-Glutamine

(Lonza), and 5% penicillin and streptomycin (Lonza) for 7 days with media change on days 3 and 5. Cells were then polarized to M1 or M2 macrophages by cytokine treatment:

10 ng/mL LPS + 20 ng/mL IFN-γ for 1 day to obtain M1 macrophages and 20 ng/mL IL-

4 for 2 days to obtain M2 macrophages. Cells were then plated in a 6-well plate at 1 x

106 cells/well or in 8-well chamber slides at 5 x 104 cells/well overnight before experiments.

Human macrophages were pre-incubated with vehicle (DPBSCa+/Mg+) or RvD (10 nM) for

15 minutes at 37 °C before addition with 1:50 green fluorescence protein (GFP) labeled E. coli (5 x 108 CFU/mL) (7 µg/mL, BacLight, Molecular Probes, Eugene, OR,

USA) or 1:3 fluorescently labeled apoptotic PMN (10 µM, CellTraceTM CFDA,

Molecular Probes, Eugene, OR, USA) for 1 hour (Oh, Pillai et al. 2011). Subsequently, E. coli (2.5 x 106 CFU/well) were added, and fluorescence was assessed after 60 min

(37 °C) using a BZ9000 microscope equipped with a 320x objective (Keyence, Itasca, IL,

USA). The apoptotic cells were added into the 6-well plates with macrophages for 1 hr, after which the cells were washed thoroughly and kept on ice, to stop phagocytosis.

These cells were then harvested and subjected to flow cytometry staining to measure

34 mean fluorescence intensity as measure of efferocytosis with appropriate controls.

Trypan blue (1:50 dilution) was added to quench extracellular fluorescence.

PLD activity assay

Lysed macrophages (50 µg) or lysed lung tissue samples (50 µg) (protein measured by

Bradford’s method) were processed for PLD2 activity in PC8 liposomes and [3H]n- butanol beginning with the addition of the following reagents (final concentrations): 1

3 µM PIP2, 3.5 mM PC8 phospholipid, 45 mM HEPES (pH 7.8), and 1.0 µCi [ H]n-butanol in liposome form, as described (Liscovitch, Czarny et al. 2000). Samples were incubated for 20 min at 30oC with continuous shaking. Addition of 0.3 ml ice-cold chloroform/methanol (1:2) stopped the reactions. Lipids were then isolated and resolved by thin layer chromatography. The amount of [3H]-PBut that co-migrated with PBut standards was measured by scintillation spectrometry.

Real-time (quantitative) Reverse Transcriptase-PCR

Reverse transcription coupled to qPCR was performed following published protocols

(Henkels, Miller et al. 2016). Total RNA was isolated from macrophages with the

RNeasy mini-kit following the manufacturer’s protocol (Qiagen, Valencia, CA, USA).

RNA concentrations were determined using a Nano-Drop spectrophotometer, and samples were normalized to 29 ng cDNA/µl. Reverse transcription was performed with 2

µg of RNA, 210 ng of random hexamers/primers, 500 µM dNTPs, 84 units of RNase

OUT and 210 units of Moloney murine leukemia virus reverse transcriptase and incubated at 42°C for 55 min. Quantitative PCR reactions were run with 100 ng of total input RNA (3.45 µl), 10 µl of the gene expression assay (RT2 SYBR Green ROX qPCR

Master Mix) (cat. # 330520, Qiagen) and 1 µl of the relevant mouse RT2 qPCR Primer

35 Assay. The following mouse primer sets were used from Qiagen (Valencia, CA, USA):

TBP (PPM03560F) (used as a housekeeping gene), PLD1 (PPH02835A), PLD2

(PPH02787A), S6K (PPH00791F), PLD3 (PPH02828A), PLD4 (PPH19319A) and PLD6

(PPH08933A). Q-PCR reactions were run with 100 ng total input RNA, 1 µl (which contained 250 nM of the probe and 900 nM of the primers) of either FAM-labeled PLD1

(Hs00160118_m1 Cat#: 4331182) or FAM-labeled PLD2 (Hs01093219_m1 Cat#:

4351372) gene expression assay multiplexed with the FAM-labeled housekeeping genes

Actin (Hs01060665_g1 Cat#: 4331182), GAPDH (Hs02758991_g1 Cat#: 4331182), and

TATA-Binding protein (Hs00427621_m1 Cat#: 4331182). Quantitative PCR conditions for the Stratagene Cycler were: 95°C for 10 min followed by 50 cycles of the next 2 steps: 30 s 95°C and then 1 min 55°C, followed by 1 cycle of 1 min 95°C, 30 s 55°C and

30 s 95°C to establish the dissociation curves. The “cycle threshold” Ct values were chosen from the linear part of the PCR amplification curve where an increase in fluorescence can be detected at >10 S.E. above the background signal. ΔCt was calculated as: ΔCt = Avg. PLD Ct - Avg. Housekeeping Ct and gene -fold expression

-( Ct) -(experimental condition Ct - control Ct) was calculated as: 2 ΔΔ = 2 Δ Δ .

SDS-PAGE and Western blot analysis

To confirm the presence of endogenous PLD1, PLD2 and S6K proteins in RAW264.7 macrophages or primary human macrophages, we performed SDS-PAGE and Western blot analyses specific for each of these three proteins, as well as TBP as the equal protein loading control. Approximately, 150 µg of protein lysate was loaded per each lane of the

SDS-gels that were then used for Western blot analyses. For western-blots, rabbit PLD1

(F-12) IgG (Santa Cruz Biotechnology, Santa Cruz, CA, USA) (cat. # sc-28314), rabbit

36 PLD2 (N-term) IgG (Abgent, San Diego, CA, USA) (cat. # AP14669a), rabbit S6K IgG

(49D7) (Cell Signaling Technology, Danvers, MA, USA) (cat. # 2708), rabbit TBP IgG

(Cell Signaling Technology) (cat. # 8515), rabbit anti-WASp IgG (D9C8) (Cell Signaling

Technology, Danvers, MA, USA) (cat. # 4271), rabbit anti-phospho-WASp (phospho

Y290) (Abcam, Cambridge, MA, USA) (cat. # ab59278), mouse anti-Grb2 IgG (3F2)

(EMD Millipore, Billerica, MA, USA) (cat. # 05-372) and rabbit anti-Actin (13E5) IgG

(Cell Signaling Technology) (cat. # 4970) were utilized as primary antibodies according to the manufacturers’ recommendations. Anti-rabbit IgG HRP and anti-rabbit IgG HRP antibodies were from Cell Signaling Technology (cat. # 7074 and 7076, respectively).

Immunoreactivities were detected using enhanced chemiluminescence (ECL) reagents from GE Heatlhcare (Pittsburgh, PA, USA) (cat. # RPN2106) and autoradiograph film.

Myeloperoxidase (MPO) Assay

MPO is an enzyme that is mostly present in neutrophils and correlates with the extent of neutrophil infiltration into tissues. The MPO assay was performed using the MPO Mouse

Myeloperoxidase DuoSet ELISA (R&D systems, Minneapolis, MN) (cat# DY3667).

Lungs were harvested from mice 2 hours after reperfusion. Samples were washed in cold phosphate-buffered saline (PBS, pH 7.4), immediately frozen on dry ice and stored at -

80ºC. For MPO assay, the lung samples were thawed, weighed, and homogenized in 1x

PBS (pH 7.4) and centrifuged (1000 x g, 5 min, 4°C). The resulting supernatant was used for the MPO assay. Assay plate were prepared by coating the plate with capture antibody overnight at room temperature. The plate were then washed and blocked. After blocking, samples were added with respective control and standards. After thorough washing, detection antibody was added and incubated at room temperature for 2 h. Then the wells

37 were washed thoroughly and streptavidin-HRP was added for 20 mins, followed by substrate for 20 min at room temperature. The reaction was stopped using a stop solution

(2N H2SO4). The samples were read in a micro-plate reader at 450 nm with wavelength correction set to 540 nm or 570 nm. Results were expressed as units per microgram (wet weight) of protein in tissue.

Isolation of bone marrow-derived monocytes

Bone marrow from WT, PLD1-/- and PLD2-/- euthanized mice was extracted from femurs and tibias according to (Swamydas and Lionakis 2013). The bones were washed once in

70% ethanol and twice in 1X PBS. The epiphyses (ends of the femur and tibia) were cut carefully with a new, single-edge razor, and then, using a 12 cc syringe and 25 G x 5/8 in. needle, RPMI media with 10% FBS and 2mM EDTA was used to flush out the bone marrow cells, which were passed through a 100 µm cell strainer placed on top of a 50 ml conical tube. This step was repeated from the other end of the bone to obtain the maximum number of cells possible. The bones were then cut into pieces, placed on top of the cell strainer, and crushed with the back of the syringe to recover any remaining cells.

The cells were then sedimented at 1400 rpm for 7 min at 4 oC, resuspended in 1X PBS, and counted. Cells were allowed to settle in the centrifuge tube and then the supernatant media carefully aspirated. The cells were resuspended in 20 ml of 0.2% NaCl to lyse red blood cells and incubated for 20 seconds. 20 ml of 1.6% NaCl was then added and the cells sedimented at 1400 rpm for 7 min at 4 oC. The pellet was resuspended in 1 ml RPMI media with 10% FBS and 2 mM EDTA, centrifuged at 1400 rpm for 7 min at 4 oC, and resuspended in 1 ml of RPMI media with 10% FBS and 100 ng/ml M-CSF.

38 Differentiation of bone marrow cells into bone marrow-derived macrophages

(BMDM)

Bone marrow cells were plated at a density of 2.0-2.5 x 107 cells/plate in 100 mm tissue culture-grade plates. Media was changed on Day 3 and again on Day 6 with RPMI-1640,

10% FBS, Pen-Strep, and 100 ng/ml M-CSF. On Day 7, the BMDM were washed twice with PBS and harvested by incubation in 10 ml PBS containing 10 mM EDTA, followed by vigorous pipetting using complete RPMI media with 100 ng/ml M-CSF. The BMDM were sedimented at 250xg for 5 min and counted using a hemocytometer. Cells were then plated into 12-well plate at 2.5 x 105 cells/well for foam cell assay or into 6-well plate at

2.5 x 105 cells/well for immunofluorescence (IF) staining.

Preparation of aggregated, oxidized LDL particles (Agg-Ox-LDL)

Extensively oxidized LDL can interact with scavenger receptors on phagocytic cells

(Steinbrecher, Parthasarathy et al. 1984, Steinbrecher, Witztum et al. 1987) leading to delivery of LDL to macrophages by phagocytosis and subsequent formation of foam cells

(Steinberg, Parthasarathy et al. 1989). LDL aggregation, fusion, and lipid droplet formation are important early steps in atherogenesis (Lu and Gursky 2013). As LDL oxidation increases aggregation (Oorni, Pentikainen et al. 2000) we used the method in

(Lougheed and Steinbrecher 1996), starting with high concentration of already oxidized

LDL (cat. # L34357) from Life technologies (Carlsbad, CA) at 1 mg/ml, and incubation with 20 µM copper for 24 h in a 15-ml conical tube with continuous rotation at medium speed setting. As per the manufacturer’s instructions each batch was tested for optimal levels of oxidation and functionally tested with bovine pulmonary artery epithelial cells for recognition by scavenger receptors (Cat# L34357) (Life technologies, Carlsbad, CA).

Phagocytosis of Agg-Ox-LDL by macrophage foam cells

BMDM were obtained from c57BL/6 mice or were differentiated into macrophages by continuous culture with 100 ng/ml M-CSF (Manzanero 2012). RAW264.7 macrophages were incubated as we have described in (Bruntz, Lindsley et al. 2014, Gomez-

Cambronero and Kantonen 2014, Frohman 2015). Cells were plated in a 12-well plate at a density of 2.5 x 105 cells/well and allowed to grow overnight in a humidified 37 oC incubator. Agg-Ox-LDL were added to the wells (0, 50 or 100 µg/ml) for 24h or 48h after which the media was aspirated and the cells washed in 1X PBS. Foam cells were fixed in 4% paraformaldehyde, washed with 1X PBS, and then incubated twice with 1 ml of 100% propylene glycol for 5 min each time. The supernatants were aspirated and the cells stained with 1 ml of 65 oC Oil Red O for 1 h. The stain was aspirated and the cells washed with 85% propylene glycol for 3 min followed by 2x in distilled water. The cells were counter-stained using hematoxylin for 2 min and washed 7x with water. After air drying, photomicroscopic imaging was conducted using an EVOS microscope (40x magnification). Oil Red O-stained foam cells were quantified in terms of fold-difference compared to control samples (Zhao, Li et al. 2006, Waldo, Li et al. 2008). Foam cell formation was measured using Image J. The bright-field images were opened and scale was set to 200 µm. The images were then set to grayscale using RGB stack, with threshold set to half the maximum threshold. The images were then analyzed by clicking measure to obtain the intensity of oil red staining.

40 Phagocytosis of Zymosan

Macrophage phagocytosis was performed as per our prior report (Kantonen, Hatton et al.

2011). Briefly, phagocytosis was measured using green fluorescent protein (GFP)-labeled zymosan A (Saccharomyces cerevisiae) (Life Technologies, Carlsbad, CA, USA). The particles were opsonized at 37 °C for 1 h using zymosan A BioParticles opsonizing reagent (derived from highly purified rabbit polyclonal IgG antibodies) (Invitrogen,

Carlsbad, CA) and added to 1 x 107 RAW264.7 macrophages such that there were approximately 20 zymosan particles per each cell. The 6-well plates containing the cells were sedimented at 800 x g for 5 min and incubated at 37 °C for 15 min. Fluorescent- labeled zymosan particles ingested by macrophages were counted manually under the microscope in five different fields and the averages + SEM calculated.

Immunofluorescence microscopy of phagocytosis and key proteins

Macrophage foam cells were incubated with Agg-Ox-LDL in a time-course experiment to measure phagocytic cup formation, similar to (Kantonen, Hatton et al. 2011). Samples were fixed, permeabilized, blocked and then incubated with fluorescent antibodies specific to WASp, Grb2 and PLD2 using AlexaFluor350-, FITC- or TRITC-labeling of two out of the three total proteins at any one time, allowing us to visualize fluorescent

Agg-Ox-LDL and two target proteins (Kantonen, Hatton et al. 2011, Reichard,

Cheemarla et al. 2015). Photomicrographs at 100x magnification were rendered using a

Nikon Eclipse 50i inverted fluorescence microscope, Infinity2 Lumenera digital camera and Infinity Analyze software (v. 6.2).

Human artery specimens

41 Our IRB protocol number is SC 5561 (Wright State University School of Medicine,

Dayton, Ohio). Small sections of normal human artery and popliteal vein controls, and also diseased popliteal artery and plaque from ileofemoral endarterectomy were obtained from deceased donor organs. Consent for research with the obtained tissue was by next of kin. Tissues were obtained and either flash-frozen or stored in formalin. The formalin fixed-tissue samples were used for sectioning for Immunofluorescent staining or tissue histology H&E staining (AML Laboratory Inc., FL). The flash-frozen sections were further processed with lysis buffer containing collagenase and phosphatase inhibitors by homogenization and ultrasonication for western blot analysis after SDS-PAGE or gene expression analysis (qRT-PCR).

Computational transcriptomic analysis

For the data presented in Figure 30, we used gene expression data from the human carotid artery atheroma study included in the NCBI Gene Expression Omnibus (GEO) microarray dataset (Ayari and Bricca 2013). We chose this study as it included carotid artery atheromatous plaque samples from hypertensive human patients compared to control companion healthy artery tissue. We used data from a single study to assure that it was generated using similar methodologies. The study we chose had 34 pairs of normal, adjacent carotid tissue and atheroma plaque. Per Ayari et al., gene expression values were scaled logarithmically (Ayari and Bricca 2013). All of the data from the genes of interest were downloaded to MS Excel spreadsheets from GEO during July 2016. Genes were defined as being over- or underexpressed when the fold-change in the atheroma plaque group was significantly different compared to the adjacent normal carotid tissue (p<0.05).

42 Results were analyzed using GraphPad Prism 6.0 (GraphPad Software, San Diego, CA,

USA). Unpaired t-tests were used to compare gene expression data.

PLD inhibitors

The PLD inhibitors VU0155069 (selective for PLD1), that has an IC50 of 9 nM (Ganesan,

Mahankali et al. 2015), VU0364739 (also called NFOT; selective for PLD2) that has an

IC50 of 10 nM (Ganesan, Mahankali et al. 2015), and FIPI (a dual PLD1/2 inhibitor) (Su,

Yeku et al. 2009) that has an IC50 of 10 nM for PLD1 and 8 nM for PLD2 (Ganesan,

Mahankali et al. 2015), were added to the cell cultures at 700 nM for 30 min prior to initiating experiments. We studied the effect of PLD inhibitors on foam cell formation after 24-48 h by Oil-red O staining.

Co-immunoprecipitation

RAW264.7 cells were cultured in reduced bicarbonate DMEM plus 10% fetal bovine serum. The plasmids used for transfections were pcDNA3.1-mycPLD2-WT, pcDNA3.1-

XGrb2 and pEGFP-C1-WASp. When cultured cells reached a confluence of 60%, they were transfected with the plasmid of interest. Transfections were performed using 5 µl

Superfect (Qiagen, Valencia, CA, USA) in Opti-MEM media previously mixed in sterile glass test tubes. The DNA and Superfect were mixed in post-transfection media (without antibiotics), applied to cells, and then transfected for 36 h. After transfection, the cells were harvested and lysed with Special lysis buffer (5 mM HEPES, pH 7.8, 100 µM sodium orthovanadate, and 0.1% Triton X-100 and protease inhibitors (aprotinin 2 mg/ml and leupeptin 5 mg/ml). The lysates were sonicated and treated with 1 µl monoclonal antibody for the respective protein and 10 µl agarose beads (Millipore, Billerica, MA)

43 and incubated at 4 °C overnight. After incubation, the immunoprecipitates were washed with LiCl wash buffer (2.1% LiCl, 1.6% Tris-HCl, pH 7.4) and NaCl wash buffer (0.6%

NaCl, 0.16% Tris-HCl, 0.03% EDTA, pH 7.4), respectively, and sedimented at 12,000 x g for 1 min. The resulting pellets were then analyzed using SDS-PAGE and Western blotting (WB).

44 Statistical Analysis

Data presented in the Figures as bars are means + Standard Error of the Mean (SEM)

(standard deviation/n1/2, were n is the sample size). Experiments were performed in technical duplicates (for PLD activity assays) for n=5 independent experiments. The difference between means was assessed by the Single Factor Analysis of Variance

(ANOVA) test or paired t-test, calculated using Prism 7 (Graphpad software Inc., La

Jolla, CA). Probability of p<0.05 indicates a significant difference. In the figures, the (*) symbols above bars denote statistically significant (P<0.05) ANOVA or t-test increase or decrease between samples and controls.

45 RESULTS

Chapter I: To study the regulation of Phospholipase D by resolvins in macrophage mediated inflammation-resolution

1.1. PLD activity and expression are altered in anti- and pro-inflammatory macrophages

Experimental design and results

Taking into consideration the fact that PLD protein expression was altered following macrophage activation, we determined the effect of each specific macrophage population on PLD in terms of protein and mRNA expression and PLD activity. As shown in Figure

11A, once Mφ(zero) were polarized into either pro-inflammatory M1 or anti- inflammatory/pro-resolving M2 macrophages by cytokine treatment, morphology changed significantly. Mzero macrophages are mostly rounded while M1 macrophages are larger in cell size and volume and are vastly vacuolated. On the other hand, M2 macrophages are large and stellate, possessing many pseudopodia-like extensions implicating their role in efferocytosis. This change in macrophage population also resulted in obvious changes in protein expression of PLD1, PLD2 and S6K (Figure 11B).

Both PLD1 and PLD2 protein and gene expression (Figure 11D-E) was increased in anti- inflammatory/pro-resolving M2 macrophages, as was S6K. However, while PLD expression was elevated in the M2 macrophages, PLD activity was highest in pro- inflammatory M1 macrophages (Figure 11C). Our preliminary data from M0, M1 and M2 macrophages indicates a varying pattern of PLD and S6K gene and protein expression in

RAW264.7 macrophages that also varies along with changes in morphology. Further, this

46 data suggests that elevated PLD activity is necessary for pro-inflammatory M1 macrophages, while upregulated PLD expression is necessary for anti-inflammatory M2 macrophages.

Figure 11. PLD activity and expression are altered in anti- and pro-inflammatory macrophages M1 and M2 macrophages were polarized from M0 RAW264.7 macrophages as in the Materials and Methods section. Polarized cells were either used for brightfield microscopy or pelleted and used for SDS-PAGE and subsequent western blotting, PLD lipase assays or qRT-PCR assay. The symbols * or ** denote statistically significant (p<0.05) between samples and controls. (A) Morphology of macrophage populations using bright-field microscopy of monolayer Mzero, M1 and M2 macrophages. Scale bar = 150 µm. (B) Relative levels of PLD1, PLD2 and S6K protein expression of macrophages using 50 µg protein per lane. ECL reagents were used to visualize immunoreactive products. GAPDH was used as the equal protein loading control. (C) PLD assay to measure the amount of TLC-isolated [3H]-PBut that co-migrated with PBut standards was measured by scintillation spectrometry. Results are mean + SEM and are expressed in terms of the total percentage of control (% of control). (D-E) Relative PLD1 (D) and PLD2 (E) gene expression. Results are quantified in terms of total relative gene expression (-fold) + SEM from n=3 performed in duplicate. GAPDH was used as housekeeping gene for qPCR experiments.

48 1.2. Effect of resolvins on PLD and S6 Kinase gene expression

Experimental design and results

Having observed the basal expression of PLD1 or PLD2 in M1 and M2 macrophages, we next determined if resolvins, which are a specific class of polyunsaturated fatty acid

(PUFA) metabolites and are considered to be specialized pro-resolving mediators

(SPMs), had any effect on PLD mRNA levels from pro-inflammatory M1 or anti- inflammatory/pro-resolving M2 macrophages. We focused our interest to resolvin Ds

(RvDs), which are polyhydroxyl metabolites of the 22-carbon PUFA, docosahexaenoic acid (DHA). Specifically, we looked at RvD1 (7S,8R,17S-trihydroxy-DHA), RvD2

(7S,16R,17S-trihydroxy-DHA), RvD3 (4S,7R,17S-trihydroxy-DHA), RvD4 (4S,5,17S- trihydroxy-DHA and RvD5 (7S,17S-dihydroxy-DHA) and their effect on PLD activity of differentiated macrophages. We treated Mzero, M1 and M2 macrophages with vehicle or

10 nM D-series resolvins (RvD1-5) for 6, 12, 18 and 24 h. These time points were chosen based on the speculation that resolvins are involved in acute inflammation resolution and will kick in early during inflammation soon after macrophages arrive to the site of inflammation (Lawrence, Willoughby et al. 2002).

We observed that RvD3, RvD4 and RvD5 significantly increased PLD1 gene expression in M1 macrophages (Figure 12A), while RvD1 and RvD5 significantly increased PLD1 mRNA levels in M2 macrophages (Figure 12B). We observed no effect of resolvin treatment on Mzero macrophages at 24 h treatment (data not shown). This data suggests that RvD3, RvD4 and RvD5 can contribute to PLD1 gene levels in pro-inflammatory M1 macrophages, while RvD1 and RvD5 contributed to PLD1 mRNA expression levels in anti-inflammatory M2 macrophages.

49 In regards to the effects of RvD treatment on PLD2 mRNA expression, we observed that while RvD5 treatment significantly downregulated PLD2 in M1 macrophages (Figure

12C), RvD5 significantly upregulated PLD2 expression in M2 macrophages (Figure

12D). RvD1, RvD3 and RvD4 also upregulated PLD2 expression in M2 macrophages albeit to a lesser extent than that of RvD5 (Figure 12D). We report no effect of resolvin treatment on Mzero macrophages at 24 h treatment (data not shown). These data suggest that an inverse relationship exists between M1 and M2 macrophages for PLD2 gene expression as a result of RvD5 treatment, whereby RvD5 may contribute to the anti- inflammatory/pro-resolving phenotype of M2 macrophages.

Figure 12. Effect of Resolvins on PLD gene expression Human M1 and M2 macrophages were treated with vehicle or 10 nM D-series resolvins

(RvD1-5) for 6 and 24 h. After resolvin treatment, M1 (A,C) and M2 (B,D) macrophages were pelleted and used for RT-qPCR assays to measure relative PLD1 (A-B) and PLD2

(C-D) gene expression levels normalized to control treated M0 macrophages. Results are quantified in terms of total relative gene expression (-fold) + SEM from n=3 performed in duplicate. GAPDH was used as housekeeping gene for qPCR experiments. The symbols (*) or (#) symbols denote statistically significant (p<0.05) increases or decreases, respectively, between samples and controls.

51 A somewhat similar effect was also observed for another gene/protein involved in PLD signaling, S6 kinase (S6K). S6K has been established as a morphogenic protein involved in cell shape change and is part of the underlying mechanism that regulates cytoskeletal structures needed for adhesion and cell locomotion during inflammation (Henkels,

Mallets et al. 2015). Similar to PLD2 upregulation by RvD5 in M2 macrophages, we observed a significant positive effect on S6K mRNA levels following RvD4 and RvD5 treatment in M2 macrophages (Figure 13C). In M1 macrophages, RvD4 and RvD5 had no effects on S6K gene expression, while RvD2 significantly upregulated S6K gene in

M1 macrophages (Figure 13B). Additionally, RvD4 and RvD5 downregulated S6K mRNA expression in undifferentiated M0 macrophages (Figure 13A). Data shown in

Figure 13 suggests S6K gene levels were differentially regulated following RvD4 and

RvD5 treatment depending on macrophage phenotype.

Figure 13. Effect of Resolvins on S6K gene expression Human Mzero, M1 and M2 macrophages that were differentiated and polarized as mentioned in materials and methods were treated with vehicle or 10 nM D-series resolvins (RvD1-5) for 6 and 24 h. After resolvin treatment, Mzero (A), M1 (B) and M2

(C) macrophages were pelleted and used for RT-qPCR assays to measure S6K gene expression levels. Results are quantified in terms of total relative gene expression (-fold)

+ SEM from n=3 performed in duplicate. GAPDH was used as housekeeping gene for qPCR experiments. The symbols (*) or (#) symbols denote statistically significant

(p<0.05) increases or decreases, respectively, between samples and controls.

53 We also considered other non-classical PLDs, such as PLD3, PLD4 and PLD6, in the different macrophage populations and what effect, if any, resolvin treatment might have on their regulation. While PLD3 was significantly downregulated in M1 macrophages by

RvD1, RvD4 and RvD5 (Figure 14A), PLD3 was significantly upregulated in M2 macrophages by RvD5 (Figure 14B).

Again, this is another example of inverse effects of RvD5 on gene expression of M1 and

M2 macrophages. Similar effects of RvD treatment on relative PLD4 and PLD6 gene expression levels were also observed in both M1 and M2 macrophages (Figure 14 C-F).

Figure 14. Effect of Resolvins on non-classical PLD gene expression Human M1 and M2 macrophages were treated with vehicle or 10 nM D-series resolvins

(RvD1-5) for 6 and 24 h. After resolvin treatment, M1 (A,C,E) and M2 (B,D,F) macrophages were pelleted and used for RT-qPCR assays to measure relative PLD3 (A-

B), PLD4 (C-D) and PLD6 (E-F) gene expression levels. Results are quantified in terms of total relative gene expression to M0 control macrophages (-fold) + SEM from n=3 performed in duplicate. GAPDH was used as a housekeeping gene for qPCR experiments. The symbols (*) or (#) symbols denote statistically significant (p<0.05) increases or decreases, respectively, between samples and controls.

55 1.3. Effect of resolvins on anti- and pro-inflammatory macrophage PLD activity

Experimental design and results

Using M1 and M2 macrophages that were derived from differentiated human monocytes, we determined the effect of resolvins on PLD activity of these M1 and M2 macrophages.

As shown in Figure 15, we found that the positive effects of RvD4 on PLD activity from

M1 macrophages and RvD5 on PLD activity from M2 macrophages was time-dependent

(significantly increased after 24 h incubation with RvD4 or RvD5, respectively) but observed no effect of RvD5 on M1 macrophages and a slight positive effect of RvD4 on

M2 macrophages. These data indicate that the pro-resolution lipid mediators, RvD4 and

RvD5, increased PLD activity in the anti-inflammatory M2 macrophages.

Figure 15. Effect of Resolvins on anti- and pro-inflammatory macrophage PLD activity Human M1 and M2 macrophages were treated with vehicle or 10 nM D-series resolvins

(RvD1-5) for 6 and 24 h. After resolvin treatment, M1 (A-B) and M2 (C-D) macrophages were pelleted and used for PLD lipase assays to measure the amount of TLC-isolated

[3H]-PBut that co-migrated with PBut standards was measured by scintillation spectrometry. Results are mean + SEM and are expressed in terms of the total percentage of control (% of control). The symbols * or ** denote statistically significant (p<0.05) between samples and controls.

57 1.4. Effect of resolvins on PLD-mediated inflammation resolution

Experimental design and results

Having observed the effects of resolvins on PLD activity and expression from anti- and pro-inflammatory macrophages, we wanted to see if resolvins mediated inflammation resolution via PLD. It is well documented that the human airway is highly susceptible to injury, which elicits an inflammatory response due to infectious agents, allergens or other damaging agents. Such sources of airway inflammation that proceed unchecked can yield tissue damage in the lungs (considered a first organ), while lung injury can also proceed as a result of inflammation/damage to another upstream organ that then negatively impacts the lungs (called second organ injury). Leukocytes play a critical role in both forms of lung damage. Our lab has demonstrated PLD’s role in cell migration in many cell types, including macrophages (Lehman, Di Fulvio et al. 2006, Gomez-Cambronero,

Di Fulvio et al. 2007, Peng, Henkels et al. 2011, Peng, Henkels et al. 2011). Uncontrolled inflammation is linked to defective generation of pro-resolving mediators, such as in asthma, which suggests that resolution of acute inflammation is critical. As the damaged, ischemic tissue initiates an intense innate immune response (inflammation), we proposed administration of resolvins to a second organ model of inflammation/injury that is mediated by PLD could keep this process at bay and yield less second organ tissue injury and a more favorable outcome.

To test this hypothesis, we administered vehicle or resolvins to PLD1-/-, PLD2-/- and WT mice in a mouse model of second organ injury to lungs using hind limb ischemia reperfusion (HLIR). Hind limb ischemia was performed for 1 h at which time vehicle or

500 nM concentration of resolvins were administered using retro-orbital injection and

58 then mice were reperfused for 2 h. Following completion of reperfusion, lungs were harvested and used for measuring myeloperoxidase (MPO) activity generated from ROS in the damaged lung tissues (Figure 16A). RvD5 reduced MPO activity in WT and PLD1-

/- mice. But RvD5 did not affect MPO activity in the PLD2-/- mice. Based on these results, we conclude that RvD5 functions via the PLD2 isoform and not the PLD1 isoform. We perfomed H&E staining on the lung tissue sections to observe lung pathology (second-organ injury) after hind-limb ischemia reperfusion injury (Figure

16B). We observed loss of honeycomb lattice structure in the WT, PLD1-/- and PLD2-/- mice after HLIR compared to non-ischemia controls. This loss of lattice structure was reverted in RvD5 treated samples in WT and PLD1-/-, but not in PLD2-/- suggesting RvD5 functions via PLD2 during HLIR.

Figure 16. Resolvin-PLD in Hind-limb ischemia reperfusion injury Lungs were harvested from WT, PLD1KO or PLD2KO HLIR subjected or no HLIR mice treated with vehicle or RvD5 at the time of reperfusion. (A) Bar graph showing absorbance values plotted in a bar graph from myeloperoxidase activity (units/ µg (wet weight) of protein in tissue) assay for lung lysates from hind-limb ischemia reperfused

WT, PLD1 -/- or PLD2 -/- treated with vehicle, 500 nM RvD2 or RvD5 at the start of reperfusion. (B) Photomicrographs showing pathology of lung sections from control or

HLIR mice treated with vehicle or RvD5 by H&E staining. Scale bar = 20 µm. All experiments were in 3 biological replicates.

60 1.5. Effect of PLD and resolvins on macrophage functions

Experimental design

We also did a macrophage functional assay using human macrophages that were stably silenced for PLD1 or PLD2 and subjected to resolvin treatment during the duration of the phagocytosis or efferocytosis assay. We found that silencing PLD2 significantly and negatively affected the effect of RvD5 on apoptotic neutrophil efferocytosis (Figure 17A) and bacterial phagocytosis by human macrophages compared to vehicle-treated, PLD2- silenced control (Figure 17B). Thus, PLD2 is important for RvD5-mediated effects on macrophage function and neutrophil activity.

Figure 17. Effect of PLD and Resolvins on macrophage functions M1 and M2 macrophages were differentiated from human Mzero macrophages as in the

Materials and Methods section. (A) Efferocytosis of apoptotic PMNs was decreased when human M2 macrophages were silenced for PLD1 or PLD2 and further decreased with RvD5 treatment. (B) Phagocytosis of E. Coli was significantly decreased in cultured human Mzero macrophages that were silenced for PLD expression using shRNAs specific for PLD2 and further decreased with 10 nM RvD5 treatment. The symbols * or

** denote statistically significant (p<0.05) between samples and controls.

62 Having discovered that RvD5 acts via PLD1 in affecting neutrophil activity and macrophage functions, we wanted to explore the signaling mechanism underlying this. In order to understand the signaling mechanism, RAW264.7 murine macrophage cells that were polarized into M1 or M2 macrophages were treated with 10 nM RvD4 or RvD5 for

6 h and subject to western blot analysis (Figure 18) for determining signaling mechanism involving PLD important for macrophage functions. ArgI is a marker for M2 macrophages and NOS2 is a marker for M1 macrophages. It was observed that RvD5 treatment increased PLD2 and phospho-S6 protein expression and decreased PLD1 protein expression in M2 macrophages.

Figure 18. Resolvins affect PLD protein expression in M1 and M2 macrophages Cultured RAW 264.7 macrophages that were differentiated into M1 or M2 were treated with vehicle, 10 nM RvD4 or RvD5 for 6 h. Cell lysates were subject to SDS-PAGE and western-blotting. Relative levels of PLD1, PLD2, NOS2, ArgI and phospho-S6 protein expression of macrophages using 50 µg protein per lane. ECL reagents were used to visualize immunoreactive products. GAPDH was used as the equal protein loading control. All experiments were in 3 biological replicates.

64 DISCUSSION

Resolution of inflammation is a multi-step process and resolvin-mediated inflammation resolution signaling mechanism is different in pro-inflammatory and anti- inflammatory macrophages exhibiting opposite effects on PLD signaling.

We know that PLD is upregulated in activated neutrophils and macrophages during inflammation (Kusner, Hall et al. 1996). Also a role of lipoxins in regulating PLD mediated inflammation is known, wherein lipoxins inhibit PLD activation by accumulation of pre-squalene diphosphate (PSDP) mainly in macrophages (Levy, Fokin et al. 1999). It is known that FPR2 is regulated by PLD. Whether or not the reverse is possible has not been tested. Since Resolvins act via GPCR (ALX/FPR2, GPR18, GPR32 or ChemR23) (Devchand, Arita et al. 2003, Serhan 2017), we studied the role of resolvins in regulating PLD protein, gene and functions. Also other studies have shown that resolvins increase inflammation initially and later promote resolution (Fredman and

Serhan 2011). As shown in Figure 19, Resolvin D5 in M2 macrophages inhibits PLD1 that is also evident from qPCR, PLD activity and Western blot data, with an alternative signaling via PLD2 suggesting a potential role in phosphorylation of S6 protein, protein synthesis with an increase in efferocytosis. A decrease in NOS2 levels and less bacterial phagocytosis in M1 macrophages suggests RvD5 suppresses M1 macrophages and their functions making them less pro-inflammatory. Thus, these results indicate that RvD5 regulates inflammation resolution by suppressing M1 macrophage activity and promoting

PLD2 activity in M2 macrophages to promote enhanced efferocytosis of apoptotic cells and thus resolution. This study thus shows a novel role of PLD in resolution of

65 inflammation, which can be used as a basis for exploring the mechanism by which RvD5 upregulates PLD2 in M2 macrophages to promote efferocytosis.

Second-organ injury is a major concern that results from ischemia reperfusion.

Lung is the second-organ most commonly affected from I/RI. Leukocytes mainly accumulate in the lungs during a second-organ injury exacerbating inflammation by releasing reactive oxygen species (ROS) and lysosomal enzymes. Myeloperoxidase

(MPO) is the enzyme important for ROS generation and is a measure of neutrophil infiltration and activity. We found that RvD5 could not reduce MPO activity in PLD2-/- lungs with hind-limb ischemia reperfusion, suggesting PLD2 is important for RvD5’s proresolving activity.

Figure 19. Model for Resolvin-PLD signaling in macrophages This model shows that resolvins activate PLD in M2 macrophages that results in increased phosphor-S6 and hence phagocytosis or efferocytosis.

67 Chapter II: To study the role of Phospholipase D in macrophage polarization

Macrophages are the first line of defense in the human body immediately after an inflammatory insult. A recent study from our lab has shown that PLD inhibitors decrease tumor-associated macrophages (more M2-like) in breast tumors and lung and liver metastasis. As PLD and PA are involved in tumor and inflammation, it would be of interest to establish whether or not PLD participates in macrophage polarization having observed their differential expression in the different macrophage populations.

Cytokines are a vast group of small proteins that are vital to cell signaling and their release affects the function/behavior of surrounding cells. Cytokines are produced by many different immune cells, such as mast cells, lymphocytes and macrophages, and by non-immune cells, such as endothelial cells, fibroblasts and stromal cells. Pro- and anti- inflammatory cytokines regulate the pathology of inflammatory diseases and are current topics of research mediating resolution of inflammation, namely as a result of ischemia/reperfusion injury (I/RI).

2.1. Effect of PLD on macrophage function

Experimental design and results

Taking this information into consideration, we induced macrophage cytocidal activity using LPS/IFNγ or MCP-1, both specific for pro-inflammatory macrophages, and measured their effects on macrophage PLD activity, which has been linked to lipid digestion and storage in phagocytic macrophages (Hume, Wells et al. 2007, Kantonen,

Hatton et al. 2011). Human Mzero macrophages were incubated with either 100 ng/ml

68 LPS plus 20ng/ml IFNγ or with 20ng/ml concentration MCP-1 for 7 -14 min. As shown in Figure 20A, increased stimulation with LPS/IFNγ slightly decreased Mzero macrophage PLD activity, while MCP-1 stimulation increased PLD activity of these

Mzero macrophages. As a result of Mzero macrophage activation, PLD1, PLD2 and S6K gene expression increased in response to MCP-1 treatment (Figure 20B, black bars).

IFNγ treatment upregulated both PLD2 and S6K gene expression but not PLD1, which remained unchanged following IFNγ stimulation. These data suggest differential effects of MCP-1 and IFNγ stimulation on gene expression of PLD1 versus PLD2 exists in human macrophages.

Figure 20. Cytokine stimulation affects PLD activity and expression Human M0 macrophages were stimulated with LPS/IFNg or MCP-1 for 0, 7 and 14 min and lipase activity was determined with [3H]-butanol or relative gene expressions were measured by qRT-PCR. (A) Total phospholipase activity of M0 macrophages. (B)

Relative gene expression of PLD1, PLD2 or S6K. GAPDH was used as the housekeeping gene. The symbols * or ** denote statistically significant (p<0.05) between samples and controls.

70 2.2. Macrophage polarization in inflammation

Experimental design and results

We know that PLD plays an important role in macrophage functions such as chemotaxis and phagocytosis (Kantonen, Hatton et al. 2011). Although PLD has always been implicated in inflammation, its role in resolution of inflammation is not known. In order to study the role of PLD in inflammation and resolution, it is important to know the expression levels of PLD in the different macrophage subpopulations that occur in the process. We also know that PLD expression is altered after cytokine stimulation. First to differentiate Mzero macrophages, RAW264.7 macrophages were polarized to M1 Mφ upon treatment with 100 ng/ml LPS and 20ng/ml IFNγ for 24 h or to M2 Mφ following treatment with 20 ng/ml IL-4 for 48 h. As shown in Figure 21A, morphology changed significantly once Mzero macrophages were differentiated into either pro-inflammatory

M1 or anti-inflammatory/pro-resolving M2 macrophages. Stimulation of Mzero macrophages with LPS/IFNγ yielded flattened, spherically-shaped M1 polarized macrophages, while IL-4 treatment yielded elongated macrophages with a distinct M2 phenotype. We then measured the gene expression of M1 specific marker gene NOS2

(Figure 21B) and M2 specific maker gene ArgI (Figure 21C) in M0, M1 and M2 macrophages. From findings in chapter 1, we can say that PLD1 expression levels correspond to M1 marker NOS2 and PLD2 expression levels correspond to M2 marker

Arg I.

Figure 21. Macrophage polarization in inflammation (A) Photomicrographs of representative morphology of M0, M1 and M2 RAW264.7 macrophages. Scale bar = 150 µm. (B-C) Gene expression levels of NOS2 (B) and ArgI

2 technical replicates. The symbols * denote statistically significant (p<0.05) changes between samples and controls.

72 2.3. PLD affects human M0 macrophage polarization

After finding out differential expression of PLD1 and PLD2 (Figure 11) in the different polarized macrophage subpopulations or by cytokine stimulation (Figure 20) and the correlation between PLDs and M1 or M2 markers (Figure 21), we wanted to investigate the role of PLD in macrophage polarization. For which, we transfected M0 macrophages with PLD1 or PLD2 plasmid, overexpression (Figure 22A) or silencing (Figure 22B) and measured the expression of cell surface markers specific for M1 macrophages (CD80 and

CD86) or M2 macrophages (CD163 and CD206) by flow cytometry. We observed that overexpressing PLD2 increased M1 markers and decreased M2 marker expression

(Figure 22A), while silencing increased M2 marker expression (Figure 22B). Thus, PLD2 induces macrophage polarization.

To establish the importance of PLD expression to macrophage function, we silenced PLD expression of human macrophages following differentiation from monocytes using overexpression of shRNA specific for PLD1 or PLD2 and measured PLD activity of silenced macrophages. As shown in Figure 22C, PLD activity was decreased to ~30-40% when each of the two predominant PLD isoforms was silenced individually.

Furthermore, to establish the importance of PLD expression to macrophage function, we also measured phagocytosis in real-time by microscopy after stimulating the macrophages with fluorescently labeled E.Coli. We observed that silencing PLD1 or

PLD2 affected not only the polarization, but also significantly decreased macrophage phagocytic function (Figure 22D-E).

Figure 22. PLD affects human M0 macrophage polarization (A) Representative bar graph showing the mean fluorescence intensity from flow cytometry staining for CD80 and CD86 (M1 markers) or CD163 and CD206 (M2 markers) by human M0 macrophages overexpessing PLD1 or PLD2. (B) Representative bar graph showing the mean fluorescence intensity from flow cytometry staining for CD80 and CD86 (M1 markers) or CD163 and CD206 (M2 markers) by human M0 macrophages silenced for PLD1 or PLD2. (C) Lipase activity of M0 macrophages stably silenced with shPLD1 or shPLD2. (D) Phagocytosis of green fluorescence labeled E.coli by macrophages that were silenced for PLD1 or PLD2. Phagocytosis of E.coli was performed in triplicate with sample size n~60 cells read for each condition. (E) Representative micrograph of real-time phagocytosis of E.Coli by macrophages. All experiments were in 3 biological replicates and 2 technical replicates. The symbols * denote statistically significant (p<0.05) changes between samples and controls.

74 2.4. M1 to M2 macrophage class-switch induced by PLD

In order to understand the underlying mechanistic role of PLD in macrophage polarization, we determined the effect of PLD expression on the inflammation-mediated phagocytic activity of fully differentiated M1 and M2 macrophages. Also, studying the role of PLD altering M1 and M2 specific marker expression in the dormant M0 macrophages, lead us to further investigating the role of PLD later in inflammation, i.e., after an inflammatory insult, we know that macrophages polarize to M1 for proinflammation via phagocytosis of bacteria (Chung and Kocks 2012, Labonte, Tosello-

Trampont et al. 2014) and later undergo transient polarization to M2 macrophages to promote anti-inflammation via efferocytosis (Ogden, deCathelineau et al. 2001,

Nakanishi, Henson et al. 2009, Korns, Frasch et al. 2011). We also know that PLD is upregulated during inflammation. Till date, a phase and time specific role of PLD in inflammation and resolution has not been studied. In order to investigate an underlying link for the inflammation-resolution transition and the role of PLD in these processes, we polarized RAW264.7 macrophages into either M1 or M2 macrophages. M1 macrophages were transfected with plasmids to overexpress PLD1 or PLD2. As shown in Figure 23A-

23B, interestingly we observed that overexpression of PLD2, but not PLD1 significantly increased M2 marker expression (CD23 and CD206), and decreased M1 marker expression (CD38 and CD80). We quantified the number of M1 positive or M2 positive cells (Figure 23C and 23D). This data suggested a novel anti-inflammatory role of PLD2.

Figure 23. M1 to 2 macrophage class-switch induced by PLD Scatter plot showing the mean fluorescence intensity from flow cytometry staining for

CD80 and CD38 (M1 markers) (A) or CD206 and CD23 (M2 markers) (B) by

RAW264.7 macrophages M1 macrophages overexpessing PLD1 or PLD2.

Representative bar graph showing number of CD80/CD38 positive cells (C) and

CD23/CD206 positive cells (D). All experiments were in 3 biological replicates and 2 technical replicates. The symbols * denote statistically significant (p<0.05) changes between samples and controls.

76 Depending on the cytokine or chemokines signals present in an inflammatory microenvironment, macrophages may be either pro-inflammatory (M1-like) that phagocytose bacteria or anti-inflammatory (M2-like) that phagocytose apoptotic polymorphonuclear neutrophils (PMNs). In order to verify PLD has a role in macrophage function besides its role in macrophage morphology and class-switch, we analyzed the effect of either overexpressing or silencing PLD1 or PLD2 on macrophage function, such as phagocytosis of E. Coli (M1 function) or efferocytosis of apoptotic neutrophils (M2 function) (Figure 24A-B). We found that when M1 macrophages that overexpressed

PLD2 showed an increase in apoptotic PMN efferocytosis (Figure 24B) at par with M2 macrophages and when PLD2 was silenced in M1 macrophages there was a decrease in

E. coli phagocytosis (Figure 24A). Overexpressing or silencing PLD1 did not have an effect on both phagocytosis and efferocytosis. Both these results confirmed our previous results indicating a role of PLD in macrophage polarization and thus function by promoting resolution of inflammation by efferocytosis.

Figure 24. Effect of PLD on M1 and M2 macrophage functions (A) Phagocytosis of green fluorescence labeled E.coli by macrophages that were silenced for PLD1 or PLD2. Phagocytosis of E.coli was performed in triplicate with sample size n~60 cells read for each condition. (B) Representative bar graph showing mean fluorescence intensity from efferocytosis of green fluorescence labeled apoptotic neutrophils (PMNs) by macrophages that were overexpressing PLD1 or PLD2 measured by flow cytometry. All experiments were in 3 biological replicates and 2 technical replicates. The symbols * denote statistically significant (p<0.05) changes between samples and controls.

78 DISCUSSION

Macrophages are one of the key phagocytes in the body along with dendritic cells, neutrophils and NK cells. Macrophages are polarized to either M1 or M2 depending on the environment and stimulus. M1 macrophages are pro-inflammatory and contain many granules rich in ROS and proteolytic enzymes. These M1 macrophages secrete many pro- inflammatory mediators and their main function is to phagocytose bacteria. On the other hand, M2 macrophages are anti-inflammatory and carry out efferocytosis of apoptotic cells (Wynn and Barron 2010, Braga, Agudelo et al. 2015, Tan, Wang et al. 2016). In this study, for the first time we found that PLD induces macrophage class-switch from M0 to

M1-like and then M1 to M2-like by modulating cell surface marker expression and macrophage function. This suggests a time dependent role of PLD in the process of inflammation and later in resolution of inflammation (Figure 25). Earlier studies from our lab have shown increased infiltration of tumor-associated macrophages in secondary tumors, with an increase in PLD levels in these TAMs and also tumor-associated neutrophils (TANs) (Henkels, Muppani et al. 2016). Based on this study and the previous findings obtained it is evident that phospholipase D plays a role in macrophage polarization during inflammation in both acute and cancerous scenario. Several transcription factors have been shown to be up- or down-regulated during macrophage polarization including PU.1, NF-κB, AP-1, STAT1, STAT3 and PPARγ (Valledor,

Borras et al. 1998, Chawla, Barak et al. 2001, Lavin, Mortha et al. 2015). Previous findings from our lab has shown that PLD regulates PPAR family of transcription factors including, PPARα, LXRα and RXR both as monomers and homo- or hetero-dimers

(Mahankali, Farkaly et al. 2015). Thus, it would be interesting to explore the regulation

79 of PPARγ by PLD during macrophage polarization. Other studies indicate the upregulation of PPARγ not only in M1 to M2 class-switch, but also in tumor associated macrophages (Odegaard, Ricardo-Gonzalez et al. 2007, Wang, Liang et al. 2014).

Pharmacological inhibition of PLD can be used as a therapeutic intervention to inhibit

PLD from promoting tumor associated macrophages in the tumor microenvironment.

Figure 25. Model for PLD induced macrophage polarization This model shows that PLD induces macrophage polarization by affecting M2 markers and decreasing M1 markers.

81 Chapter III: To study the role of PLD in macrophage foam cell formation

PLD is involved in phagocytic cup formation, mediated by phosphatidic acid (PA), the product of PLD enzyme action and actin-regulatory protein WASP (Kantonen, Hatton et al. 2011). Studies have shown that Phospholipase D (PLD) regulates endothelial permeability by downregulating occludin, but it is still unclear as to what regulates this action of PLD2. Vascular leaks are a major concern in the development of atherosclerosis and other cardiovascular diseases. Hence it is important to understand the potential role of PLD in foam cell formation.

3.1. Macrophage phagocytosis of Agg-Ox-LDL is reduced in the absence of PLD2 activity

Based on earlier work from our laboratory that outlined the importance of PLD in macrophage phagocytosis and phagocytic cup formation (Corrotte, Chasserot-Golaz et al.

2006, Kantonen, Hatton et al. 2011, Ali, Chen et al. 2013), we examined the potential requirement of PLD for the ability of macrophages to differentiate into Agg-Ox-LDL- laden foam cells. Cholesterol can be taken up by macrophages by receptor-mediated processes, pinocytosis or phagocytosis. To ensure that we studied here phagocytosis, we prepared aggregated, oxidized LDL particles (Agg-Ox-LDL) of ~200 nm as detailed in

Methods, whose size is optimized for phagocytosis vs. receptor mediated endocytosis or pinocytosis. Bone marrow-derived monocytes from wild-type (WT) and PLD1- and

PLD2-deficient mice were differentiated into macrophages (Manzanero 2012) and then stimulated to undergo foam cell formation by incubation with 50 mg/ml aggregated, oxidized LDL (Agg-Ox-LDL) cholesterol (Lougheed and Steinbrecher 1996, Relevy,

82 Bechor et al. 2015). We used BMDMs because they are uniformly quiescent until stimulated, whereas intraperitoneal macrophages are activated to varying levels. Foam cell formation was monitored by staining for internalized Agg-Ox-LDL using Oil Red O, a fat-soluble dye that binds to neutral triglycerides and lipids (Xu, Huang et al. 2010, Wu,

Chen et al. 2015).

Foam cells have an enlarged, red appearance in comparison to the relatively smaller, pale-colored macrophages (Figure 26A). WT and PLD1-/- macrophages both readily took up Agg-Ox-LDL and formed foam cells to similar extents as determined by visual inspection (Figure 26B) and were not significantly different in uptake based on quantitation (Figure 26C). In contrast, PLD2-/- macrophages phagocytosed significantly less Agg-Ox-LDL compared to WT mice and formed foam cells at a reduced frequency over the time-course of the assay. Similar results were observed when the BMDM were presented with opsonized GFP-tagged Zymosan for phagocytosis (Figure 26D), indicating that the reduced rate phagocytosis is not restricted to Agg-Ox-LDL uptake.

This substantiates the importance of PLD and, specifically, the PLD2 isoform, in phagocytosis of Agg-Ox-LDL by foam cells.

Figure 26. Macrophage phagocytosis of oxidized LDL is PLD2-dependent (A) Photomicrographs of representative Oil Red O-stained macrophages versus Ox-LDL- laden foam cells. Scale bar = 20 µm. (B) Representative fields of view from photomicrographs of untreated and Ox-LDL -treated (50 µg/ml) BMDM from WT and

PLD knockout mice. Scale bar = 200 µm. (C) Quantification of the relative densities of

Oil Red O staining from foam cells form panels similar to those shown in (B) in terms of the mean density (-fold) + SEM. Phagocytosis of Ox-LDL by BMDM was performed in triplicate with sample size n~120 cells read for each condition. (D) Phagocytosis of opsonized GFP-tagged Zymosan beads by macrophages that were silenced for PLD1 or

PLD2. Phagocytosis of Zymosan was performed in triplicate with sample size n~50 cells read for each condition.

84 3.2. PLD inhibitors reduce Agg-Ox-LDL phagocytosis

PLD2 interacts with many different proteins and has been proposed in some contexts to have non-catalytic roles (Lee, Kim et al. 2009). To address whether PLD activity is key for phagocytosis, we investigated the ability of small molecule PLD inhibitors to reduce

Agg-Ox-LDL uptake from RAW264.7 macrophages. WT macrophages were cultured in media containing inhibitors that block activity of both PLD1 and PLD2 (FIPI) (Su, Yeku et al. 2009), are relatively selective for PLD1 NBOD (VU0155069) (Scott, Selvy et al.

2009, Bruntz, Lindsley et al. 2014), or are selective for PLD2 NFOT (VU0364739). Agg-

Ox-LDL uptake by macrophages was significantly inhibited by all three inhibitors, with the PLD2-selective inhibitor (VU0364739) being the most potent and the PLD1-selective inhibitor (VU0155069) being the least potent (Figure 27A). At 700nM VU0155069, the lowest concentration that significantly inhibited uptake, PLD2 is also partially inhibited

(Scott, Selvy et al. 2009), suggesting that PLD1 may have a weak if any role in the phagocytosis observed. IC50 of VU0155069 is 11 nM for PLD1 and 933 nM for PLD2 and the IC50 of VU0364739 is 20 nM for PLD2 and 1500nM for PLD1.

3.3. Agg-Ox-LDL phagocytosis via PLD2-facilitated pathways is CD36-mediated

More than half of Agg-Ox-LDL phagocytosis is thought to occur via action of the type B scavenger receptor CD36 (Collot-Teixeira, Martin et al. 2007, Montano, Boullier et al.

2013, Yu, Fu et al. 2013), which signals through Src and Jnk2 in the pathway that leads to foam cell formation (Rahaman, Lennon et al. 2006, Silverstein, Li et al. 2010). PLD has been reported to act downstream of Src (Jiang, Lu et al. 1995), presenting a possible mechanism for the effects of PLD2 inhibition on foam cell formation. To examine this

85 hypothesis, we generated BMDM from WT, PLD1 and PLD2 mice and blocked CD36 using a monoclonal antibody during culture of the cells with Agg-Ox-LDL. The CD36 antibody was effective in reducing Agg-Ox-LDL uptake for WT and PLD1-/- BMDMs, but did not reduce uptake by PLD2-/- BMDMs beyond the suppression already created by the absence of PLD2 (Figure 27B). Both these results, along with the fact that we used in the experiments Agg-Ox-LDL (particles 10x larger than individual LDL molecules) indicate that the uptake occurred through phagocytosis (as opposed to pinocytosis or receptor-mediated endocytosis). These findings also indicate that all of the CD36- mediated phagocytosis proceeds through a PLD2-regulated pathway, and that PLD2 is critical for this component of Agg-Ox-LDL uptake.

Figure 27. Inhibition of PLD reduces LDL phagocytosis (A) BMDM from WT mice were used for foam cell phagocytosis assays of Agg-Ox-LDL in the presence of 50 nM or 200 nM PLD-specific inhibitors: a PLD1-selective NBOD

(VU10155069), a PLD2-selective NFOT (VU0364739) and a dual PLD1/2 inhibitor

(FIPI). Results are expressed in terms of relative Oil red-O intensity compared to vehicle treated samples in a bar graph. Phagocytosis of Agg-Ox-LDL with PLD inhibitors was performed in triplicate with sample size n~120 cells read for each condition. (B) BMDM from WT, PLD1 KO and PLD2 KO mice were differentiated into macrophages, blocked with anti-CD36 antibody, and subject to foam cell formation assay with Agg-Ox-LDL.

All experiments were in 3 biological replicates and 2 technical replicates. The symbols * denote statistically significant (p<0.05) changes between samples and controls.

87 3.4. The heterotrimeric protein complex of PLD2-Grb2-WASp

PLD2, through its product phosphatidic acid (PA), mediates cell migration of macrophages and neutrophils (Lehman, Di Fulvio et al. 2006, Frondorf, Henkels et al.

2010). This signaling pathway is also dependent on Wiskott-Aldrich syndrome protein

(WASp) (a crucial protein needed for actin polymerization and, as such for phagocytosis) and on Growth factor receptor-bound protein 2 (Grb2), with Grb2 functioning as the docking protein that mediates the physical interaction between PLD2 and WASp (Figure

28A). The SH2 domain of Grb2 interacts with the Y169 and Y179 residues of PLD2, while the two SH3 domains of Grb2 interact with the polyproline region of WASp. This heterotrimeric protein binding interaction is dependent on phosphatidic acid (PA), as

RAW264.7 macrophages incubated in the presence of 300 nM PA exhibited increased interaction of PLD2-Grb2-WASp as a function of time, as shown using co- immunoprecipitation (Figure 28B). A recent study has shown that exogenously-supplied

PA can be detected by a PA sensor inside cells during phagocytosis (Kassas, Tanguy et al. 2017). Moreover, from previous studies in our laboratory, we know that exogenously- supplied PA activates PLD, resulting in increased activity and downstream cellular signaling cascades (Henkels, Miller et al. 2016, Miller and Gomez-Cambronero 2017).

To explore whether the same proteins are involved in Agg-Ox-LDL phagocytosis, we used 50 µg/ml Agg-Ox-LDL to induce BMDM cells to form foam cells and performed immunofluorescence staining to visualize PLD, Grb2, WASp and actin localization. In the absence of culture with Agg-Ox-LDL, Grb2 and PLD2 did not visually co-localize

(Figure 28C, the red (PLD2) and green (Grb2) signals are distinct).

Figure 28. The adapter protein Grb2 links PLD2 to Wiskott-Aldrich Syndrome protein (WASp) (A) Schematic of the interaction sites between PLD2, Growth factor receptor-bound protein 2 (Grb2) and actin-polymerizing Wiskott-Aldrich syndrome protein (WASp). (B)

Cultured macrophages were treated with 300 nM phosphatidic acid (dioleoyl-PA) for 0,

1, 7 or 14 min. Cell lysates were immunoprecipitated using Protein G agarose linked to antibodies specific for PLD2, WASp or Grb2 and used for SDS-PAGE. Detection of the protein-protein interactions on subsequent Western blots using a two protein approach indicates protein-protein interaction occurred between PLD2 and WASp, between PLD2 and Grb2 and between WASp and Grb2, which suggests formation of a protein heterotrimeric complex (PLD2-Grb2-WASp) in the presence of PA (i.e., in conditions where PLD2 would be activated). (C) BMDMs with or without 50 µg/ml oxLDL treatment were subject to immuno-staining with PLD2-TRITC and Grb2-FITC. Scale bar

= 50 µm.

89 In contrast, with Agg-Ox-LDL treatment, PLD2 broadly co-localized with Grb2 in WT and PLD1KO BMDMs (as evidenced by the yellow fluorescence ad indicated by the arrows). PLD2 interaction with WASp and actin was also observed to increase with Agg-

Ox-LDL treatment (Figure 29), and blockade of CD36 using the monoclonal antibody prevented the PLD2-WASp and –actin interaction. Finally, in PLD2KO BMDMs, no change in the localization pattern for WASp and actin in the presence or absence of Agg-

Ox-LDL with or without CD36 treatment was observed. Taken together, these findings indicate that the PLD2 and partner protein complex formation is CD36-dependent and that mobilization of the partner proteins requires PLD2 activity.

Figure 29. PLD2 interacts with Wiskott-Aldrich Syndrome protein (WASp) and actin (A) BMDMs were treated with 50 µg/ml Agg-Ox-LDL or Agg-Ox-LDL and CD36 followed by immuno-staining with PLD2-TRITC, actin-FITC or WASp-FITC. Scale bar

= 50 µm.

91 3.5. Arterial gene expression during atherosclerosis

To pursue clinical correlates, we mined gene expression profiles of PLD and some of its key signaling protein partners in a human disease that relies heavily on macrophage function and signaling. Through comparing relative mRNA expression levels in human atheroma plaque tissue and adjacent, non-diseased carotid artery samples, using the

Carotid Artery Atheroma Dataset at the Gene Expression Omnibus (GEO) microarray

(accession number GDS5083) (Ayari and Bricca 2013), we determined that that NFκB1,

PLD2, WASp and Grb2 (p=0.0016, p=0.005 and p<0.0001, respectively) undergo upregulation in diseased vascular tissue (Figure 30A-D). NFkb1 represented a positive control, as it is a well-known marker of inflammation. In contrast, there was no change in expression of PLD1 (Figure 30E, p=0.24), further supporting its lack of relevance in this disease context.

Figure 30. Bioinformatic analysis of PLD and its signaling proteins in human atheroma plaque tissues Microarray data were from the NCBI Gene Expression Omnibus (GEO) and specifically from the carotid artery atheroma dataset (Ayari and Bricca 2013). (A-E) NFκβ

(p=0.0016), PLD2 (p=0.005), Grb2 (p<0.0001) and WASp (p<0.0001) are significantly upregulated in diseased atheroma plaques, while PLD1 is not (p=0.2432). The symbols * denote statistically significant (p<0.05) increases between atheroma plaque and control adjacent carotid artery samples.

93 3.6. PLD2 but not PLD1 is upregulated in diseased artery tissues

To confirm that hypoxic conditions actually do trigger PLD expression in diseased human artery and atheromatous plaques, we used human undamaged control and diseased atherosclerotic artery specimens to assess both relative mRNA and protein expression levels. We performed quantitative RT-PCR and/or SDS-PAGE/western blot analyses on human patient samples and measured the expression levels of PLD, Grb2 and WASp. We analyzed samples for both PLD isoforms and found that while PLD2 mRNA was significantly upregulated in the diseased artery and plaque samples (Figure 31A), PLD1 mRNA was virtually undetectable (Figure 31B). Similar findings were observed for

PLD2 and PLD1 protein expression levels by western blot (Figure 31C); PLD2 was upregulated in the diseased artery samples that contained atherosclerotic plaque (both from diseased femoral artery and ileofemoral endarterectomy), while PLD1 was downregulated.

3.7. Upregulation of Wiskott-Aldrich Syndrome protein (WASp) and Grb2 in diseased artery tissues

We similarly investigated the expression of WASp and Grb2 in human undamaged control and diseased atherosclerotic artery specimens. WASp and Grb2 protein expression levels were increased in the diseased artery samples compared to the non- diseased companion tissues (Figure 31C-D). Furthermore, WASp activation was increased, as assessed by detecting phosphorylation at the WASp tyrosine 290 residue, using a phosphotyrosine-specific antibody (Figure 31C-D). These data suggest a role for activation of WASp protein in the diseased artery and plaque samples, which could play a

94 role in membrane-associated actin polymerization at the leading edge of phagosomes involved in phagocytosis of Agg-Ox-LDL and foam cell formation.

Figure 31. PLD2, Grb2 and phospho-WASp are upregulated in diseased artery tissues Human patient samples comprised of normal, non-diseased popliteal artery and diseased popliteal artery and the atheromatous plaque contained within it were used for qRT-PCR and SDS-PAGE/western blot analyses. (A-B) Relative mRNA expression levels of PLD2 (A), PLD1 (B), expressed in terms of mean (-fold) expression + SEM relative to housekeeping gene (TATA Binding Protein, TBP). (C) Protein expression levels of PLD2 and Actin, PLD1, WASp and phospho-WASp and Grb2. Actin is the equal protein loading control. The Western blots and gene expression assays were done in biological triplicates (n=3). (D) Densitometry ratio of p-WASp/WASp shown in bar graph. The symbols * denote statistically significant (p<0.05) changes, respectively, between samples and controls using one-way ANOVA or t-test.

96 3.8. Validation of PLD2, WASp and Grb2 in diseased human artery samples

Formalin-fixed companion samples to those diseased tissues shown in the Fig. 6 mRNA and protein expression assays were used for H&E staining and immunofluorescence microscopy (IF). As shown in Figure 32A, H&E staining revealed positive staining in the diseased artery towards the lumen, adjacent to but not in the acellular area. The positive staining in the plaque, which is from the same vessel, looks contiguous with the staining in the diseased artery, and that area is not highly cellular (based on DAPI). PLD1 IF staining of tissues with rabbit α-PLD1-FITC-conjugated IgG antibodies showed little-to- no PLD1 expression in the plaque and endarterectomy arteries, similar to the diseased artery alone (Figure 32B).

In contrast, immunofluorescence staining of these tissues using rabbit α-PLD2-FITC- conjugated IgG antibodies indicated that PLD2 expression is concentrated in the plaque in diseased arteries (Figure 32C) and in the plaque that was removed during endarterectomy when compared to the diseased artery alone. The positive PLD2 staining in the diseased artery (Figure 32C, left panel) appears to be towards the lumen. The positive staining in the plaque (Figure 32C, middle panel), appears to be adjacent to the staining in the diseased artery. Thus, PLD2 appears in the area of the plaque underneath the fibrous cap.

Figure 32. PLD2, WASp and Grb2 are overexpressed in diseased human artery samples Diseased human arteries, plaque from that diseased arteries and plaque removed during endarterectomy samples, which are companions to those shown in the previous figure; they were fixed in paraformaldehyde, paraffin-embedded, sectioned, mounted and used for H&E and immunofluorescence (IF) stainings. (A) H&E stained samples were magnified at 10x. Scale bar = 200 µm. (B-C) IF stained samples for PLD1 (B) or PLD2

(C). Proteins of interest were FITC stained, while nuclei were stained with DAPI.

Merged images of FITC and DAPI staining are shown for each sample. Arrows point at zones were accumulation of PLD2 staining was obvious. IF samples were magnified at

100x. Scale bar = 200 µm.

98 DISCUSSION

Phospholipase D and its catalytic product PA have been linked to cell signaling, cell proliferation, inflammation and cancer. Phospholipase D (PLD) is a major signaling component in myeloid leukocytes, where it plays a key role in cytoskeletal actin polymerazation and phagocytosis where both isoforms, PLD1 and PLD2 coordinately regulate macrophage phagocytosis (Iyer, Barton et al. 2004, Kantonen, Hatton et al.

2011). The findings reported in this study center on macrophage phagocytosis of oxidized-LDL and their conversion into foam cells. The data suggest PLD2-specific mechanism in foam cell formation, which is important in the progression of atherosclerosis.

Chronic inflammation is responsible for the development and continued progression of atherosclerosis (Nabel and Braunwald 2012), which encompasses a great portion of current healthcare costs (Mozaffarian, Benjamin et al. 2015). Thus, it is very important to limit or terminate inflammation and start vascular healing. Plaque growth and swelling in a diseased artery involves migration of immune cells and the conversion of macrophages into foam cells (Ross 1999). In atheroma formation, cholesterol in the form of LDL accumulates in the intima layer of arteries. The plaque is a complex mixture of cholesterol crystals, extracellular matrix, smooth muscle cells, endothelial cells, monocytes, macrophages and foam cells (Schwartz, Valente et al. 1992, Finn, Nakano et al. 2010). Macrophages are recruited to the LDL accumulation and attempt to clear it.

Foam cell formation occurs as a result of receptor-dependent and receptor-independent

(phagocytosis) uptake of Agg-Ox-LDL, causing cholesteryl ester accumulation in these cells (Mietus-Snyder, Friera et al. 1997, Barbieri, Cavalca et al. 2004, Hansson,

99 Robertson et al. 2006). Although much is known about receptor-mediated endocytosis, what role PLD might have in phagocytosis of LDL is less clear.

The findings reported here are a substantial departure from the status quo of eliminating the plaque in atheroma and instead defines an entirely new angle of approaching the problem, which is to avoid formation of functional foam cells. The proposed research strategy shifts existing paradigms by characterizing a heretofore-unsuspected link between PLD2 and the progression of inflammation in atherosclerosis. We identified that

PLD2 makes macrophage foam cells phagocytose plaque Agg-Ox-LDL more efficiently in a process that is dependent on mechanism that includes PLD2-Grb2-WASp.

Other studies have shown that PLD Agg-Ox-LDL uptake is increased in J774.1 murine macrophages (Mori, Itabe et al. 2001), but a direct role of PLD in foam cell formation has not been shown to date. Our study reveals that PLD2 not only regulates macrophage phagocytosis by phagocytic cup formation, but also promotes receptor-mediated phagocytosis of Agg-Ox-LDL via CD36 (a type II scavenger receptor). Having learned the molecular mechanisms of PLD2-WASp mediated phagocytosis of Agg-Ox-LDL enabled us to conclude how foam cell formation is affected by PLD2. Absence of PLD2 prevents action of CD36 to induce foam cell formation.

The data presented in this chapter support a key role for PLD2, as PLD2-/- macrophages phagocytose aggregated Agg-Ox-LDL (Figure 26B-C) and zymosan (Figure 26D) less efficiently and PLD2-specific inhibitors diminish phagocytosis (Figure 27B). Gene and protein expression from diseased human artery and plaque samples indicate PLD2 but not

PLD1 correlates with diseased vessels (Figure 31). Thus, we have demonstrated that

PLD2 plays a role in foam cell phagocytosis whereas PLD1 does not.

100 WASp is linked with actin polymerization, and we show that phospho-WASp is present in the atheromatous plaque removed during ileofemoral endarterectomy from diseased arterial samples (Figure 31). Likewise, we also observed that PLD2 and Grb2 were upregulated in the diseased artery/plaque samples with very low basal expression in non- diseases artery (Figure 31). Based on this, we propose that once activated by hypoxia,

PLD hastens the phagocytic capability of foam cells through a mechanism that involves

CD36, PLD, Grb2 and WASp interaction and subsequent actin remodeling to form phagocytic cups (Figure 33).

Overall, we present a new paradigm: that PLD-Grb2-WASp heterotrimeric signaling axis in macrophages is a key mediator of macrophage phagocytosis of Agg-Ox-LDL and subsequent foam cell formation, as indicated in the Model presented in Figure 33. This knowledge provides specific new protein targets in vascular biology that have potential implications for improving human health. The proposal that elevated PLD, Grb2 and

WASp expression/activation may support progression of the atherosclerosis is significant, as potent, new small-molecule inhibitors are currently available that could be used to pharmacologically inhibit foam cell formation.

According to the present study, using the PLD1-KO and PLD2-KO mice from references

(Chen, Hongu et al. 2012) and (Oliveira, Chan et al. 2010), respectively, the PLD2-/- macrophages do not take up Agg-Ox-LDL as well as WT macrophages do. However, it is entirely possible that they would eventually (meaning days or weeks, as atherosclerosis progresses slowly) take Agg-Ox-LDL up, permitting the formation of foam cells. Such delay in clearing would raise the local concentration of Agg-Ox-LDL in the extracellular

101 space under the endothelial cells, which could help activate them to create a pro- inflammatory environment.

There is always the intriguing possibility that time progression could cause a switch of the role of PLD from harmful to protective (or vice versa). In a related model of ischemia reperfusion indicate PLD1 found to be protective –when experiments are done after 28 days after initial reperfusion (chronic phase of reperfusion) (Schonberger, Jurgens et al.

2014). However, the absence of PLD1 or PLD2 (from KO mice) are found to be protective, when experiments are performed for shorter (6-24 hr) periods of time (acute phase of reperfusion, our data). For the case at hand on PLD and athrosclerosis, it will be interesting to see what role PLD’s isoforms would play in long-term studies by using

ApoE/PLD double KOs.

102

Figure 33. Model of PLD2-mediated foam cell atherogenesis Foam cell accumulation leads to atherogenesis. Foam cell formation by Agg-Ox-LDL uptake by scavenger receptor CD36 on macrophages mediates PLD2 interaction with

WASp and Grb2 resulting in actin polymerization, that allows phagocytosis as indicated for the first time in this study.

103 CONCLUSIONS

From the studies above, we have an understanding of i) potential regulation of phospholipase D by resolvins in macrophages; ii) role of PLD in macrophage polarization; and iii) role of PLD in foam cell formation and the signaling mechanism involved in foam cell formation during atherosclerosis (Figure 34).

104

Figure 34. Model of PLD2-mediated inflammation and resolution in macrophages This model shows how PLD regulates macrophage polarization and functions during inflammation (atherogenesis) and how resolvins facilitate resolution via PLD in macrophages.

105 LIST OF ABBREVIATIONS

Agg-ox-LDL- Aggregated oxidized low density lipoprotein

ALX/FPR2- Lipoxin A4/N-Formyl peptide receptor 2

ArgI- Arginase I

BMDM- Bone marrow derived macrophages

CD- Cluster of differentiation

CFSE- Carboxyfluorescein succinimidyl ester

ChemR23- Chemerin receptor 23

CVD- Cardiovascular disease

DHA- Docosahexanoic acid

DPBS- Dulbecco’s Phosphate-Buffered Saline

EPA- Eicosapentanoic acid

FAK- Focal adhesion kinase

FBS- Fetal Bovine Serum

FPR- Formyl peptide receptor

GAPDH- Glyceraldehyde-3-phosphate dehydrogenase

GEF- Guanine nucleotide exchange factor

106 GEO- Gene expression Omnibus

GM-CSF- Granulocyte macrophage colony stimulating factor

GPCR- G-protein coupled receptor

Grb2- Growth factor bound receptor 2

HLIR- Hind-limb ischemia reperfusion

I/RI- Ischemia reperfusion injury

IFN- Interferon

IL- Interleukin

LPS- Lipopolysaccharide

LXR- Liver X receptor

MCP- Macrophage chemoattractant protein

M-CSF- Macrophage colony stimulating factor

MDSC- Myeloid-derived suppressor cells miR- Micro RNA

MMP- Matrix metalloproteinases

MPO- Myeloperoxidase

NADPH- Nicotinamide adenine dinucleotide phosphate

107 NOS2- Nitric oxide synthase 2

Ox-LDL- Oxidized low density lipoprotein

PA- Phosphatidic acid

PC- Phosphatidyl choline

PH- Pleckstrin homology

PLD- Phospholipase D

PMN- Polymorphonuclear neutrophils

PPAR- Peroxisome proliferator-activated receptor

PUFA- Polyunsaturated fatty acids

PX- Phox homology

ROS- Reactive oxygen species

RvD- Resolvins D

RXR- Retinoid X receptor

SPM- Specialized pro-resolving lipid mediators

TAM- Tumor associated macrophages

TAN- Tumor associated neutrophils

TBP- TATA-box binding protein

108 TGF- Tumor growth factor

TNF- Tumor necrosis factor

TSC- Tuberous Sclerosis Complex

VEGF- Vascular endothelial growth factor

VSMC- Vascular smooth muscle cells

WASp- Wiskott Aldrich Syndrome protein

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