Sectm1a Deficiency Aggravates Inflammation-Triggered Cardiac Dysfunction
Through Disruption of LXRα Signaling in Macrophages
A dissertation to be submitted to the
Graduate School of the University of Cincinnati
In partial fulfillment of the requirements for the degree of
DOCTOR OF PHILOSOPHY
in the Department of Pharmacology and Systems Physiology, College of Medicine, 2020
By:
Yutian Li
B.S. in Pharmaceutical Science from The Ohio State University, 2012
M.S. in Molecular, Cellular and Biochemical Pharmacology from University of Cincinnati, 2014
Committee Members: Guo-Chang Fan, Ph.D (Chair)
David Hui, Ph.D
Terence Kirley, Ph.D
Diego Perez-Tilve, Ph.D
Jack Rubinstein, MD
David Wieczorek, Ph.D Abstract
Acute and chronic inflammation are reflected by systemically greater abundance of proinflammatory cytokines and increased infiltration and activation of immune cells in various tissues. In particular, cardiac dysfunction is a common ailment associated with both acute and chronic inflammatory states. As a fundamental component of innate immunity, macrophages play critical roles in both initiating and resolving inflammation in the heart. In fact, macrophages are prominent cells that drive septic cardiomyopathy in animal models; and human monocytes/macrophages secrete more inflammatory cytokines in type 2 diabetic patients and positively correlate with atherosclerosis severity. Secreted and transmembrane protein 1 (Sectm1, also referred to as K12) is a type 1 transmembrane protein. The knowledge of Sectm1 function in human diseases is currently limited to its role as an alternative CD7 ligand to stimulate T cell proliferation. Whether Sectm1 plays a role in normal macrophage biology and inflammatory diseases has never been investigated.
In this dissertation, we observed that mRNA levels of Sectm1a (mouse homolog of human
Sectm1) was significantly increased in early time points (peak at 6 h), but reduced at later time points in LPS-treated bone marrow-derived macrophages (BMDMs) and spleen of wild-type
(WT) mice injected with LPS. To determine the role of Sectm1a in macrophage activation and inflammation-induced cardiac injury, we generated a Sectm1a-knockout (KO) mouse model in which LPS-induced cardiac injury and mortality were greatly augmented. Further analysis revealed that inflammatory macrophages in hearts of KO-LPS mice was greatly accumulated, compared to WT-LPS controls. In accordance to the activated macrophage phenotype, lack of
Sectm1a dramatically increased the production of inflammatory cytokines (TNFα, IL-6, and IL-
i 1β) and MCP-1 levels both in vitro (BMDMs) and in vivo (in serum and myocardium) after LPS challenge. Moreover, we detected significantly lower levels of proinflammatory cytokines when overexpressing Sectm1a in BMDMs, but not in cardiomyocytes. Most importantly, transplantation of Sectm1a-KO bone marrow cells into WT mice resulted in increased accumulation of inflammatory macrophages in the heart and aggravated cardiac dysfunction upon LPS challenge. These data suggest that ablation of Sectm1a induces cardiac dysfunction through activation of immune responses mediated by macrophages.
Furthermore, RNA-sequencing results, along with bioinformatics analyses showed that many of the LXRɑ target genes are significantly downregulated in Sectm1a KO BMDMs.
Furthermore, ablation of sectm1a hinders the nuclear translocation of LXRα in response to
GW3965 (LXR agonist), resulting in higher levels of inflammatory cytokines. In addition, administration of GW3965 fails to rescue cardiac function in KO mice upon LPS injection.
Notably, coimmuno- precipitation (Co-IP) results suggest potential physical interaction between
Sectm1a and LXRα. Lastly, using chronic inflammation model induced by high-fat diet (HFD,
18-24 week) feeding, we also observed that infiltration of inflammatory monocytes/ macrophages in KO-hearts was dramatically increased, leading to aggravated cardiac dysfunction, compared to WT-HFD controls.
In summary, this study defines a novel function of Sectm1a in macrophage biology, and identifies a new cellular mechanism for Sectm1a in the regulation of macrophage activation via
LXRα signaling cascade, and its relationship to inflammation-induced cardiac injury.
ii (Blank Page)
iii Acknowledgement
Throughout the entire journey towards earning my Ph.D. degree, I have been very fortunate to meet different people and make many friends, who have played significant role in my professional and personal development. Their support and encouragement are absolutely indispensable, making this endeavor much more enjoyable and fruitful than it could possibly be, and for this I will always be grateful.
First, I want to express my sincere gratitude to my thesis advisor, Dr. Guo-Chang Fan, for providing me a strong platform and countless guidance along my path to becoming an independent scientist. Being a great mentor as he is, Dr. Fan has always been extremely supportive and inspiring, his passion to science and rigorous work ethic had motivated me to maximize my potential on the daily basis. He is attentive to details and has regularly trained me on reviewing others’ publications and writing protocols/fellowship applications. All these had significant improved my critical thinking, writing, and presentation skills, and has been invaluable tools to my professional and personal development that will benefit me for the rest of my life. Apart from being an exemplary scientist, Dr. Fan is also a great host and organizer for many fun events that I enjoyed during the past few years: celebrating birthdays and holidays, or when manuscripts had been accepted for publication or new funding was awarded.
Furthermore, I would like to acknowledge my committee members, Drs. David Hui,
Terence Kirley, Jack Rubinstein, Diego Perez-Tilve, and David Wieczorek, for their constructive and idea-inspiring criticism, which kept me moving forward with my projects and ensured the timely completion of this dissertation. I am very thankful that they were always encouraging me and available to discuss my projects.
iv Next, I’d also like to extend my appreciation to many great collaborators. First, I am greatly appreciative of the mentoring and training on metabolic studies received from Dr. Hui’s lab when I did my lab rotation in his lab and throughout my graduate study; particularly, I’d like to thank Dr. Allyson Hamlin, David Kuhel, James Cash, and Joshua Basford, who taught me all the basic techniques commonly used in metabolic studies and answered endless question that I had. In addition, I would like to thank Jenna Holland and Emily Yates, in Dr. Diego Perez-
Tilve’s lab, who helped me with the project of Hsp20’s role in regulation of adipocyte function, which broadly expand my knowledge in lipid studies. Moreover, I would like to extend my gratitude to Dr. Rubinstein, and his lab members, Mr. Nathan Robins and Dr. Sheryl Koch, for their time and effort in performing and analyzing echocardiography as well as explaining the technique for me to understand the project better. Also I want to thank Dr. Yigang Wang for allowing me to use some of their equipment; particularly, I’d like to thank Wei Huang for assisting me with some of the echocardiography and Co-IP experiments.
I’d also like to acknowledge many great people in the Department of Pharmacology and
Systems Physiology. I am very appreciative to Drs. John Maggio, Abdul Matlib, and Robert
Rapoport for their guidance and mentorship since the beginning of the Master Program and throughout my graduate study. I also like to express my gratitude to Nancy Thyberg and Jeannie
Cummins, who are extremely supportive in all matters and keep me in track to finish things in time, they are the first go-to person whenever I have any question, and they will always be ready to help. I’m also very thankful to my classmates and fellow graduate students, Kobina Essandoh,
George Gardner, Fawzi Alogaili, Jiuzhou Huo, for their collaboration and friendship for the past several years.
v I am fortunate to have encountered many great people in the Fan lab. Particularly, I want to thank Dr. Xiaohong Wang for her mentorship and endless support throughout my graduate study, making it more enjoyable. I also owe my sincere gratitude to the past and present members, Drs. Liwang Yang, Dongze Qin, Haitao Gu, Jiangtong Peng, Shan Deng, Xingjiang
Mu, Peng Wang, Hongyan Zhao, Lu Wang, and Shunan Cui for their assistance and friendship.
I am honored to be awarded with the University Research Council Award, Albert J. Ryan
Fellowship, and the American Heart Association Pre-Doctoral Fellowship, which all provided me valuable resources and allowed me to network with fellow students and former awardees at annual symposiums. In addition, I’d like to thank Drs. Hong-Sheng Wang, Yigang Wang, and
David Wieczorek for their full support on my AHA Pre-Doc Fellowship application.
Lastly, I would like to thank my parents for their unconditional love and support, I’m thankful for their patience, encouragement and guidance throughout my life, and I’m forever grateful for their sacrifice for me to pursue my career. Next, I want to thank my wife, Yuqiu, for her overwhelming support and being the best cook in the world, thank you for believing in me and talking care of my life, I’m very blessed to have you in my life.
vi (Blank page)
vii Table of Contents
Abstract i
Acknowledgement iv
Table of Contents 1
List of Abbreviations 4
List of Figures and Tables 8
Chapter I: Introduction 11
Section 1: Introduction of Inflammation-Associated Cardiac Dysfunction 11
I.1.A. Overview of Acute and Chronic Inflammation 12
I.1.B. Effects of Inflammation on Cardiac Function 16
I.1.C. Functional Roles of Macrophages in the Heart 21
Section 2: Human Secreted and Transmembrane Protein 1 (Sectm1) and 23
Its Mouse Homologs Sectm1a/Sectm1b
I.2.A. Discovery, Structure and Expression of Sectm1 23
I.2.B. Functional Roles of Sectm1 in Immunity 25
I.2.C. Mouse Homologs Sectm1a and Sectm1b 26
Section 3: Role of Liver X Receptor in Macrophages 28
I.3.A. Overview of Anti-Inflammatory Functions of LXR 28
I.3.B. Effects of LXR in Cardiac Function 30
Chapter II: Materials and Methods 34
Section 1. Generation of Sectm1a-Knockout (KO) Mouse Models 34
Section 2. Mouse Model of Acute Inflammation Induced by Endotoxemia 36
1 Section 3. High Fat Diet (HFD)-Induced Chronic Inflammation Model 36
of Obesity
Section 4. Cell Isolation, Culture, and Treatments 37
II.4.A Culture of cell lines 37
II.4.B Isolation of bone marrow-derived macrophages (BMDMs) 37
II.4.C Isolation of adult rat cardiomyocytes (ARCMs) 38
II.4.D Construction and infection of Sectm1a adenovirus vector 38
II.4.E Treatments on macrophages 39
Section 5. In Vivo Assessment of Cardiac Function 39
Section 6. Gene Expression and Protein Measurements 40
II.6.A qRT-PCR experiments 40
II.6.B RNA sequencing 41
II.6.C Western-blotting experiments 42
Section 7. Cytokine Measurements using ELISA Assays 43
Section 8. Immunofluorescent Staining 43
Section 9. Flow Cytometry 44
Section 10. Co-Immunoprecipitation Assay 46
Section 11. Bone Marrow Cell Transplantation 47
Section 12. Statistical Analysis 47
Chapter III: Results
Section 1. Expression Profiles of Sectm1a in Different Organs and 49
BMDMs with or without LPS Challenge
Section 2. Sectm1a Deficiency Aggravates LPS-Induced Systemic 54
2 Inflammation and Mortality
Section 3. Ablation of Sectm1a Leads to Exacerbated Cardiac 57
Inflammation and Dysfunction
Section 4. Lack of Sectm1a Augments LPS-Induced Inflammation 64
via Skewing BMDMs toward Proinflammatory Phenotype
Section 5. Gene Enrichment Analysis of Sectm1a-KO BMDMs 73
Section 6. LXR Agonist Fails to Rescue LPS-Induced Inflammation 77
and Cardiac Dysfunction in Sectm1a-KO Model
Section 7. Sectm1a-KO Provokes HFD-Induced Inflammation and 84
Cardiac Dysfunction
Section 8. Sectm1a Deficiency-Mediated Cardiac Dysfunction is Mainly 89
Ascribed to Augmented Inflammatory in the Heart
Chapter IV: Discussion 97
Section 1. Dissertation Summary 97
Section 2. The role of Sectm1a in regulating inflammatory response 99
of macrophages
Section 3. Effects of Sectm1a on LXR signaling pathway 102
Section 4. Limitations and future directions 103
Section 5. Conclusion of the dissertation 104
References 106
3 List of Abbreviations
Abbreviation Full Name ABCA1 ATP-bindin cssette transporter A1 ACE Angiotensin-converting enzyme Ad. Adenovirus Ang II Angiotensin II ANP Atrial natriuretic peptide ApoE Apolipoprotein E ARCM Adult rat cardiomyocyte ASC2 Activating signal co-integrator 2 AT1R Angiotensin type 1 receptor BMDM Bone marrow-derived macrophage BNP B-type natriuretic peptide bp base pair BW Body weight CANTOS Canakinumab anti-inflammatory thrombosis outcomes study Cas9 CRISPR associated protein 9 CCR2 C-C chemokine receptor type 2 CD Chow diet CIRT Cardiovascular inflammation reduction trial CLP Cecal ligation and puncture Co-IP Co-immunoprecipitation ConA Concanavalin A CRISPR Clustered regularly interspaced short palindromic repeats CTL Cytotoxic T lymphocyte CXCL1/2 C-X-C motif chemokine ligand 1/2 CXCR2 C-X-C motif chemokine receptor 2
4 CX3CR1 C-X3-C motif chemokine receptor 1 DAMP Damage-associated molecular pattern DIO Diet-induced obesity DMSO Dimethyl sulfoxide ECM Extracellular matrix Eid1 EP300-interacting inhibitor of differentiation ELISA Enzyme-linked immunosorbent assay EP300 E1A-associated protein p300 ER Endoplasmic reticulum FASN Fatty acid synthase FBS Fetal bovine serum GFP Green fluorescent protein GLUT Glucose transporter GITR Glucocorticoid-induced TNFR-related protein gRNA guideRNAs HDL High density lipoprotein HFD High fat diet HFrEF Heart failure with reduced ejection fraction HSC Hematopoietic stem cell IAP Inhibitor of apoptosis ICAM1 Intracelllular adhesion molecule 1 IFN-γ Interferon γ IL-1β Interleukin 1β IL-6 Interleukin 6 i.p. Intraperitoneally KO Knockout LDL Low-density lipoprotein LOX Lipoxygenase LPS Lipopolysaccharide
5 LTB4 Leukotriene B4 LV left ventricle LVEF LV ejection fraction LXA Lipoxin A LXR Liver X receptor LXRE LXR response element MAMP Microorganism-associated molecular pattern MCP-1 Monocyte chemoattractant protein-1 M-CSF Macrophage colony-stimulating factor MDSC Myeloid-derived suppresive cell min Minute MOI multiplicity of infection MNC Mononuclear cell NCoR Nuclear receptor co-repressor NF-κB Nuclear factor κ-light-chain-enahncer of activated B cells NK Cell Natural killer cell NO Nitric oxide PAM protospacer adjacent motif PAMP Pathogen-associated molecular pattern PBS Phosphate buffered saline PCR Polymerase chain reaction PEPCK Phosphoenolpyruvate carboxykinase PFA Paraformaldehyde PMN Polymorphonuclear neutrophil PMSF Phenylmethylsulfonyl fluoride PRR Pattern recognition receptor PUFA Polyunsaturated fatty acid qRT-PCR Quantitative real time PCR
6 RAAS Renin-angiotensin-aldosterone system RBC Red blood cell RCT Reverse cholesterol transport RFP Red fluorescent protein RNAseq RNA sequencing ROS Reactive oxygen species RT Room termperature RXR Retinoid X receptor Sectm1(a/b) Secreted and transmembrane protein 1(a/b) SERCA Sarcoplasmic reticulum Ca2+ ATPase SIRS Systemic inflammatory response syndrome SIRT1 Silencing information regulator 1 SMRT Silencing mediator of retinoic acid and thyroid hormone receptor SPM Specialized proresolving lipid mediator SREBP1c Sterol regulatory element binding protein 1c TEs Thymic epithelial cells TGFβ Transforming growth factor β TNFɑ Tumor necrosis factor ɑ VCAM1 Vascular cell adhesion molecule 1 WGA Wheat germ agglutinin WT Wild type
7 List of Tables and Figures
Table 1. PCR protocol for Sectm1a-KO genotyping 35 Table 2. Primers for measurement of Sectm1a and Sectm1b gene expression 35 Table 3. Primers used for qRT-PCR analysis 40 Table 4. Antibodies used for Western Blotting experiments 42 Table 5. Antibodies used for immunofluorescent staining 44 Table 6. Antibodies used for Flow Cytometry 45 Table 7. Primary antibodies used for co-immunoprecipitation 46 Table 8. Echocardiographic measurements of WT and Sectm1a-KO mice 59 12 h after LPS injection Table 9. Echocardiographic measurements of WT and Sectm1a-KO mice with 82 GW3965 and LPS injections Table 10. Echocardiographic measurements of WT and Sectm1a-KO mice with 87 20 weeks of high fat diet feeding Table 11. Echocardiographic measurements of LPS-treated recipient mice 93
after transplantation of bone marrow cells from WT (WT-LPS)
and Sectm1a-KO (KO-LPS) mice
Figure 1. The temporal variation in acute and chronic inflammatory response 11 Figure 2. Distribution of major nonmyocyte types in the heart 20 Figure 3. Structure of classic Ig-domain and predicted Sectm1a Ig-like domain 24 Figure 4. CRISPR/CAS9 cassette for Sectm1a-KO mouse model 34 Figure 5. Tissue distribution of Sectm1a in WT mice 51 Figure 6. Kinetics of LPS-stimulated gene expression of Sectm1a 52 Figure 7. Expression of Sectm1a in whole blood and spleen after LPS treatment 53 Figure 8. Validation of Sectm1a-KO mouse model 55
8 Figure 9. Systemic inflammation is increased in Sectm1a-KO mice after 56 LPS injection Figure 10. Sectm1a deficiency increases LPS-induced mortality 56 Figure 11. Knockout of Sectm1a exacerbates LPS-triggered cardiac dysfunction 58 Figure 12. Gating Strategy for Flow Cytometry analysis of cardiac macrophages 60 Figure 13. Ablation of Sectm1a enhances accumulation of inflammatory macrophages 61 in hearts from LPS-treated mice Figure 14. Sectm1a deficiency causes increased macrophage infiltration in the heart 62
upon LPS challenge
Figure 15. Lack of Sectm1a increases cardiac cytokine levels in Sectm1a- 63
KO mice after LPS injection
Figure 16. Ablation of Sectm1a has no effect on macrophage maturation in vitro 66
Figure 17. Knockout of Sectm1a augments cytokine release from BMDMs 67
Figure 18. Absence of Sectm1a skews BMDMs toward proinflammatory phenotype 68
Figure 19. Sectm1a deficiency activates NF-κB pathway 69
Figure 20. Overexpression of Sectm1a in BMDMs suppresses NF-κB pathway 70
Figure 21. Overexpression of Sectm1a reduces cytokine production in BMDMs 71
Figure 22. Upregulation of Sectm1a does not affect cytokine gene expression in ARCMs 72
Figure 23. RNA-seq analysis using WT and Sectm1a-KO BMDMs 74
Figure 24. Sectm1a deficiency downregulates LXRα-targeted genes 75
Figure 25. Deletion of Sectm1a affects LXRα, but not LXRβ pathway 76
Figure 26. Lack of Sectm1a impairs LXRα translocation to nucleus after stimulation 79
with agonist
Figure 27. LXR agonist fails to rescue LPS-induced inflammation in 80
Sectm1a-KO BMDMs
9 Figure 28. Administration of LXR agonist shows no effect on cardiac function 81
in Sectm1a-KO mice upon LPS injection.
Figure 29. Sectm1a interacts with LXRα 83
Figure 30. Lack of Sectm1a promotes palmitate-induced macrophage activation 85
Figure 31. Sectm1a-KO mice show impaired cardiac function upon HFD feeding 86
Figure 32. Absence of Sectm1a leads to increased accumulation of proinflammatory 88
macrophages in hearts of obese mice
Figure 33. Transplantation of WT and Sectm1a-KO bone marrow cells 91
Figure 34. Transplantation of Sectm1a-deficient bone marrow cells aggravated 92
cardiac dysfunction after LPS injection
Figure 35. Transplantation of bone marrow cells from Sectm1a-KO mice has no 94
effect on cardiac neutrophil infiltration upon LPS stimulation
Figure 36. Transplantation of bone marrow cells from Sectm1a-KO mice increases 95
monocyte-derived macrophage population in the heart upon LPS stimulation
Figure 37. Sectm1a-deficient bone marrow cells give rise to inflammatory macrophages 96
with increased CCR2 expression after LPS treatment
Figure 38. Graphic schema depicting effects of Sectm1a deficiency on macrophage 105 activation and subsequently cardiac injury.
10 Chapter I: Introduction
Section 1: Introduction of Inflammation-Associated Cardiac Dysfunction
The term inflammation is derived from the Latin word “Inflammare” (to burn).
Inflammation is generally regarded as host responses to contain, dilute, and neutralize a foreign agent or injurious tissue, followed by a series of healing processes including regeneration of native parenchymal cells or formation of fibroblastic scar tissue to replace injured tissue (1, 2).
Immediately after detection of injury or infection, changes in vascular flow and permeability occur, allowing the migrating immune cells to enter the inflammatory site. The major cell type(s) responding to the inflammatory stimuli varies with the time of the onset of inflammation (Fig. 1).
Neutrophils are generally the predominant cell type recruited to the injured area during the first few days, because chemokine signals for neutrophil migration are increased and activated at Figure 1. The temporal variation in acute and chronic inflammatory early stage of inflammation response. Adapted from James et al., 2013 (Ref. 1)
(3). However, since neutrophils are short-lived and disintegrate after 48 h, they are replaced by monocytes/macrophages at later time points. Activation of chemokine factors for monocytes lasts longer, resulting in continuous migration of monocytes until inflammation is resolved (1).
Despite the development of many therapeutic approaches to treat inflammation-associated diseases, new strategies aiming to halt aberrant inflammatory signaling are gaining more attention due to growing number of patients that have become intolerant or show serious adverse
11 effects of existing treatment options. However, the overall benefits of such strategy remains inconclusive based on the results from different clinical trials. For example, the Cardiovascular
Inflammation Reduction Trial (CIRT) reported that effects of low-dose of methotrexate in patients with stable atherosclerosis did not differ from placebo treatments (4). However, according to the results from Canakinumab Anti-Inflammatory Thrombosis Outcomes Study
(CANTOS), targeting IL-1β pathway with Canakinumab was able to lower the rate of recurrent cardiovascular events without affecting lipids (5). Therefore, more efforts are required to delineate the molecular and cellular regulation of inflammatory responses, which will provide valuable insights into the effects of inflammation in a variety of diseases and may guide the development of new treatment strategies.
I.1.A. Overview of Acute and Chronic Inflammation
Acute inflammation is a rapid host-protective response of relatively short duration, lasting for minutes to days depending on the type and extent of infection or injury, and this will lead to signs and symptoms such as heat, swelling, pain, and loss of function (6). Some common stimuli for acute inflammation include: microbial invasion, ingestion of noxious/toxic compounds, or trauma (2). The hallmark for acute inflammation initiation is tissue edema following increased blood flow and vascular permeability to the injured site. Subsequently, polymorphonuclear neutrophils (PMNs) and relatively smaller number of monocytes migrate to the area to defend against foreign objects. Meanwhile, professional phagocytes, neutrophils and monocyte-derived macrophages are activated upon arrival to the inflammatory site and undergo the phagocytosis process: recognition, attachment, engulfment, and degradation of the injurious agent or pathogens (7, 8).
12 Systemic acute inflammation is often investigated experimentally in animal models.
Three of the most common procedures are: 1) exogenous administration of endotoxin (i.e. LPS);
2) treatment with viable pathogens (i.e. injection of Escherichia Coli); and 3) disruption of the protective barrier (i.e. cecal ligation and puncture model, CLP) (9). Among those, the LPS model offers several essential advantages when exploring the effects of new molecules in acute inflammation (10). Comparing to other models, administration of LPS is widely used as experimental animal or human endotoxemia model for studies of acute systemic inflammation
(11, 12). Injection of LPS is an easy technique with high reproducibility, particularly when assessing the inflammatory responses induced by LPS; and the extent and onset of systemic inflammatory response syndrome (SIRS) as well as lethality can be induced depending on the dose choices of LPS (9, 13). High levels of cytokines (both proinflammatory and anti- inflammatory) can be detected in circulating serum shortly after LPS injection. For instance, serum levels of TNFα, IL-6, and IL-10 peaks at 2 h after LPS (5 mg/kg of body weight) injection and gradually returns to baseline over 72 h, whereas IFNγ peaks at 6 h after same dose of LPS injection (10). In addition, the inflammatory responses elicited by LPS also include increased infiltration of neutrophils and monocytes/macrophage into many organs (i.e. liver, spleen and heart) and subsequent tissue damage and loss of function (10, 14, 15).
Chronic inflammation is oftentimes referred as slow, long-term inflammation that is perpetuated by persistent stimuli (i.e. obesity, cancer, or exposure to low level of irritant or foreign objects that could not be eliminated by the host). The duration may range from several months to years depending on the cause of injury and the capacity of the host to overcome and repair the damage (1, 2). Failure to resolve inflammation in a timely manner would lead to irreversible tissue damage and substantially increase the risk of chronic inflammatory diseases
13 such as cardiovascular diseases, arthritis, and asthma (16, 17). Due to its increasing prevalence worldwide, the World Health Organization has ranked chronic inflammation-mediated diseases as the greatest threat to human health, the prevalence of some specific chronic diseases includes:
1) cardiovascular diseases that account for 31% of all death globally; 2) diabetes that is affecting more than 30 million people in the US; and 3) allergies that rank the 6th leading cause of chronic human diseases in the US (2).
Many of the features of acute inflammation remain as inflammatory responses develops into chronic condition, and chronic inflammation becomes histologically less uniform due to the presence of a collection of immune cells (monocytes/macrophages and lymphocytes) and increased angiogenesis and proliferation of connective tissue (1). Some common signs and symptoms present during chronic inflammation includes: body pain, fatigue, GI track complications and frequent infections (2). Rodent models of chronic inflammation include the antigen-induced arthritis or dextran sodium sulfate-induced colitis. However, active inflammatory responses in these models resolves after several days or few weeks. In contrast animal models with diet-induced obesity (DIO) using high fat diet (HFD) are easily maintained and very commonly used to mimic different pathophysiological conditions in human. Feeding mice with HFD (60% energy from fat) for prolonged period of time could increase adipocyte proliferation and growth, promote infiltration of monocytes/macrophages and subsequently stimulate chronic inflammatory responses as evidenced by increased plasma levels of cytokines
(18).
Timely resolution of inflammation is critical to prevent damage to the host by immune system and to restore tissue homeostasis. Acute inflammation is generally resolved quickly, usually less than 1 week, due to its inherent negative feedback regulation (1, 19). The initiation
14 of resolution phase is marked by cessation of neutrophil recruitment, which are the first responders and the most abundant leukocyte population at inflammatory area during early stage of injury (28). This process is largely regulated by “proresolving” mediators including lipid metabolites from omega-3 and omega-6 polyunsaturated fatty acids (PUFA) such as resolvins and lipoxins, protein and bioactive peptides (i.e. Annexin A1 and Galectin 1) and gases (i.e. nitric oxide and carbon monoxide (21, 22). Indeed, neutrophil infiltration is blocked partially due to the downregulation of chemokine, CXCR2, facilitated by the proresolving mediators (23).
Furthermore, these mediators actively regulate PMN apoptosis, stimulate phagocytosis, promote chemokine scavenging, and promote tissue repair and regeneration (24-26). Importantly, as terminally differentiated cells, infiltrated neutrophils start undergoing apoptosis shortly after reaching the target tissue, and along with other injured/apoptotic cells, they will release signals such as nucleotides and externalized phosphatidylserine that can be readily recognized by monocyte-derived macrophages (27, 28). Furthermore, the uptake and digestion of apoptotic cells (or efferocytosis) is an anti-inflammatory process that is coupled with reduction of pro- inflammatory signaling. Key mediators for efferocytosis includes the TAM (Tyro3, Axl, and Mer) tyrosine kinases (19, 29).
Importantly, specialized proresolving lipid mediators (SPMs) play a central role in inflammation resolution because of their rapid production by different immune cells. As the production of pro-inflammatory mediators initiates in leukocytes, various enzymes responsible for SPM generation are also produced and activated. For example, 5-lipoxygenase (5-LOX) is required for production of leukotriene B4 (LTB4, a leukocyte chemoattractant) from omega-6
PUFA; meanwhile, 5-LOX can also convert the intermediates in this process into lipoxins in the presence of 12-LOX. Moreover, 5-LOX can cooperate with 15-LOX (derived from
15 monocyte/macrophage, PMNs, epithelial cells, or eosinophils) to produce lipoxins (19, 26, 30-
32). Nonetheless, the action of these specialized proresolving mediators varies depending on the type of target cells. For example, Lipoxin A4 (LXA4) could inhibit chemotaxis of neutrophils while stimulating chemotaxis and adhesion of monocytes; it can also inhibit IL-1β-induced IL-6 production from fibroblasts and inhibits IL-12 release from dendritic cells (26). Given the wide range of cellular targets and the ability to promote host defense and healing process while suppressing inflammatory responses, members of SPMs or their bioactive analogues represent potential therapeutic targets to treat inflammation-associated diseases.
I.1.B. Effects of Inflammation on Cardiac Function
Cardiac dysfunction is a common feature associated with both acute and chronic inflammatory states. The first evidence that linked heart failure with inflammatory response was reported in 1990, when patients with heart failure with reduced ejection fraction (HFrEF) showed increased serum levels of TNFα compared with healthy individuals (33). Since that seminal study, the role of inflammation on cardiac function has been a topic of intensive research. A recent study has demonstrated that functional recovery of the heart from ischemia/reperfusion injury following intracardiac injection of stem cell or Zymosan requires infiltration and induction of CCR2+ and CX3CR1+ macrophages (34), rather than stem cell differentiation into cardiomyocytes as previously proposed (35-37). In addition, results from the CANTOS clinical trial showed that, comparing to placebo, anti-IL-1β treatment could lower the rate of recurrent cardiovascular events and heart failure-related hospitalization and mortality in patients with previous myocardial infarction (5, 38). This evidence suggests that targeting inflammatory processes may provide efficacious therapies for patients with heart failure.
16 There are numerous different causes for myocardial injury, including myocarditis caused by infectious pathogens, cardiomyopathy associated with metabolic syndrome and diabetes or, most commonly, ischemic cardiomyopathy following coronary artery blockage. Upon injury, physiological inflammation mediated by the innate and adaptive immune systems are activated with upregulation of cardioprotective responses, providing the heart with temporary adaptation to increased stress. However, the condition will become pathological if the inflammatory responses are not resolved in a timely manner, resulting in secondary damage to the heart and ultimately heart failure (17, 39).
Major mediators of inflammatory responses can be categorized into non-cellular and cellular components. Pattern recognition receptors (PRRs) are expressed by different cell types in the heart such as cardiomyocytes, endothelial cells, and tissue resident immune cells.
Immediately after injury, these receptors are activated upon recognition of signals from pathogen-associated molecular patterns (PAMPs), damage-associated molecular patterns
(DAMPs, released from dying cells), and microorganism-associated molecular patterns
(MAMPs), resulting in augmented production of proinflammatory cytokines (TNFα, IL-1β, and
IL-6) and chemokines (MCP-1), which are all critical elements of the non-cellular component
(40, 41).
Pro-inflammatory cytokines are highly potent endogenous polypeptides produced mainly by activated macrophages; other cellular origins include lymphoid cells, mast cells, endothelial cells, and fibroblasts (42, 43). Patients with heart failure exhibit increased levels of proinflammatory cytokines, and those levels are positively correlated to heart failure severity and prognosis. More importantly, proinflammatory cytokines have been shown to produce negative inotropic effects in isolated cardiomyocytes, ex vivo hearts, and in vivo animal models (44-46,
17 47). Specifically, TNFα suppresses Ca2+ handling in cardiomyocytes through two main mechanism: 1) inhibiting the expression and activity of sarcoplasmic reticulum Ca2+ ATPase
(SERCA) mediated by activation of NF-κB cascade (48); 2) by provoking Ca2+ leakage through activation of caspase-8, which leads to nitric oxide (NO) and reactive oxygen species (ROS) production and subsequently causes S-nitrosylation and destabilization of Ryanodine receptor
(49). In addition, stimulation of cardiomyocytes with TNFα strongly induces apoptosis through activation of sphingomyelinase pathway (50). Similarly, Schulz et al. demonstrates that the negative inotropic effects of IL-1β are also mediated, at least partially, through NO production
(51). Additionally, treatment of neonatal rat cardiomyocytes with IL-1β has been shown to induce apoptosis through caspase-dependent (increasing secretion of cytochrome c and caspase 3 activation) or caspase-independent [upregulating endonuclease G while downregulating surviving and inhibitors of apoptosis (IAP)] pathways (52, 53). In contrast, the effects of IL-6 on cardiomyocytes are less well understood and remain controversial, in part due to the fact that IL-
6 has both pro- and anti-inflammatory properties (54). Remarkably, the effects of IL-6 on cardiac tissue (either protective or pathogenic) depend on the duration of signaling (from acute to chronic). Acute activation of IL-6 signaling protects cardiomyocytes from oxidative stress and induces anti-apoptotic program as adaptive mechanisms for increased stress (55, 56). Yet, while
IL-6 signaling protects cardiomyocytes from damage, it decreases the basal contractility and responsiveness to β-adrenergic stimulation, thus limiting the damage and preserving cardiac tissue (57, 58). As the signal persists in the long term, myocardium undergoes genetic reprogramming to increase contractility, mainly through hypertrophy mediated by activation of
IL-6R-gp130 signaling cascade, which ultimately leads to heart failure (56, 59, 60). This postulate is consistent with findings in patients with chronic heart failure, who exhibit elevated
18 plasma levels of IL-6 associated with reduced LV ejection fraction (LVEF), atrial fibrillation, iron deficiency, and worse clinical outcome (55).
As a distinct family of cytokines, chemokines interact with their corresponding receptors to regulate diverse biological processes such as chemotaxis, collagen turnover, and angiogenesis
(17). Like cytokines, although properly activated chemokine signaling is critical for controlling infection, wound healing and restoring homeostasis, unresolved chemokine activation could lead to excessive inflammatory responses, cell death and tissue damage. One major function of chemokines is to recruit and activate specific leukocytes to the sites of injury. For example,
CXCR2 is required in TNF-induced neutrophil migration, which is dependent on CXC- chemokine ligand 1 (CXCL1) and CXCL2 as the directional cue (61); and neutrophils produces large amount of CXCL1 to recruit monocytes to myocardium during hypertrophy (62).
Additionally, when comparing to healthy individuals, patients with congestive heart failure showed significantly elevated levels of CC-motif chemokines, such as CC-chemokine ligand 2
(CCL2; also known as MCP1), which is the principal mediator for monocyte migration (63, 64).
Therefore, dysregulated chemokine signaling can contribute to heart failure progression by exaggerating inflammatory responses, and therapies targeting chemokine-receptor signaling provide great potential in attenuating inflammation-associated cardiac dysfunction.
The cellular component of the inflammatory response includes various immune cells (i.e.
T cells, B cells, or NK cells) that are recruited to the heart. As mentioned in previous section, the initial phase of immune response is marked by rapid influx of neutrophils and monocytes/ macrophages from circulation to injured sites (3, 8, 41), as the first responders to contain and neutralize foreign pathogens or injured tissues (7). This is followed by the resolution phase characterized by removal of necrotic cells by monocyte-derived macrophages, activation of
19 myofibroblasts and generation of anti-inflammatory mediators (i.e. lipoxins, IL-10) and pro- reparative molecules such as TGFβ (19, 65). The last stage of the immune response is left ventricular remodeling, distinguished by scar maturation and reduction of reparative immune cells (66). In human and mouse hearts, leukocytes account for about 9% of total cell population with exclusion of cardiomyocytes, and the majority of these cells are identified as myeloid cells (~7% in mouse heart), in particular, macrophages; whereas B cells and T cells comprise ~0.8% and 0.3% of all nonmyocytes, respectively (Fig. 2) (67).
Interestingly, B and T cells may play important roles in myocardial adaption to injury, albeit their small number in the heart. First, the number of B cells are increased in myocardium and pericardial adipose tissue after acute cardiac injury, Figure 2. Distribution of major nonmyocyte types in whereas decreasing the number of B cells the heart. Adapted from Alexander et al., 2016 (Ref. 67) with Pirfenidone or using B-cell deficient mouse model appears to protect the heart from adverse
LV remodeling and improves cardiac function (68-70). More importantly, B cells have been shown to develop autoantibodies that target many critical cardiac proteins such as troponin I,
+ + outer loop of the β1-adrenergic receptor, and Na /K ATPase (71). In addition, B cells can interact with TH-cells to increase the production of proinflammatory cytokines (72). On the other hand, adoptive transfer of CD3+ or CD4+ T cells alone from mice with heart failure could cause
20 LV dysfunction, fibrosis, and hypertrophy (73). In fact, activated T-cell populations (CD4+,
+ CD8 , TH, and Treg) expand in hearts of patients and mouse model of chronic heart failure, and these T cells produce high levels of pro-inflammatory cytokines such as IFNγ and TNF receptor
1 (73-75). Overall, although immune cells play critical roles in normal heart physiology as well as pathogenesis of cardiac damage, more investigation is needed to delineate the temporospatial regulation of different immune cells and its impact on cardiac inflammatory responses. This may eventually guide the development of new therapeutic strategies.
I.1.C. Functional Roles of Macrophage in the Heart
Macrophages are potentially the most important cell type of the cellular component in chronic inflammation due to its ability to release a large number of various biologically active products, such as neutral proteases, chemotactic factors, reactive oxygen metabolites, coagulation factors, growth-promoting factors, and cytokines (i.e. IL-6 and TNFα). These factors are critical for numerous cellular processes including growth of fibroblasts and blood vessels that are key components for wound healing (1, 2). In particular, as the most abundant leukocytes in the heart, the delicate balance of different subpopulations of macrophages is critical for effective damage control and proper tissue repair upon injury.
For the past half century, the general consensus for macrophages in the heart has been that bone marrow-derived hematopoietic stem cells (HSCs) give rise to circulating monocytes, which enter into tissues and differentiate to macrophages (76, 77). However, studies from the past decade using techniques such as fate-mapping and lineage tracing have revealed that tissue- resident macrophages are established during embryonic development, and they are able to maintain the population through self-renewal, which are distinct from recruited macrophages that
21 are differentiated from circulating monocytes (78-80). In normal unstressed condition of mouse and human hearts, the majority of cardiac macrophages are CCR2- and derived from embryonic precursors and continuously maintained through local proliferation, and they function as reparative macrophages that are associated with epithelial mesenchymal transition, coagulation, and myogenesis (79, 81). In addition, these CCR2- macrophages express many growth factors, extracellular matrix components, and conduction genes (i.e. IGF1, FGF13, GDF15, NRP1,
ECM1, SDC3, and SCN9A) (81). In contrast, CCR2+ macrophages represent the pro- inflammatory population that are maintained through differentiation and proliferation of infiltrated monocytes in the heart upon injury, they also can provoke collateral damage by recruiting neutrophils and Ly6ChighCCR2+ monocytes to the site of injury (82). Microarray analysis have revealed that CCR2+ macrophages express high levels of chemokine, chemokine receptors, and mediators of IL-1, IL-6, and NF-κB signaling pathways that augment inflammatory responses. Additionally, these macrophages exhibit higher expression of growth factors that promote fibrosis, hypertrophy, and extracellular matrix degradation (i.e. OSM,
MMP9, and TIMP1) (81).
It has been demonstrated that CCR2- macrophages, and both CCR2+ macrophages and monocytes, are all present in myocardium of patients with heart failure and mouse models of cardiac injury (i.e. permanent or reperfused myocardial infarction, diphtheria toxin cardiomyocytes ablation). Yet, the population of CCR2- macrophages gradually diminishes and is predominantly replaced by the CCR2+ infiltrating monocytes and macrophages (81-83). More importantly, the percentage and absolute number of CCR2+ macrophages positively correlates with adverse cardiac remodeling and LV dysfunction after mechanical unloading in cardiac tissue from patients with heart failure (81); and inhibition of monocyte recruitment to the heart
22 reduces adverse cardiac remodeling and ameliorates heart failure progression in myocardial infarction mouse model (83). Interestingly, under certain circumstances, namely during early stage of cardiac insult, inflammatory CCR2+ macrophages may acquire a reparative phenotype similar to resident macrophages, including upregulation of CX3CR1 expression, which may contribute to beneficial effects on cardiac function and LV remodeling (34). Nonetheless, the delicate balance and possible trans-differentiation of different subpopulation of macrophages needs to be further investigated.
Section 2: Human Secreted and Transmembrane Protein 1 (Sectm1) and Its Mouse
Homologs Secmt1a/Sectm1b
I.2.A. Discovery, Structure and Expression of Sectm1
Sectm1 was first discovered on human chromosome 17q25 in 1989 during attempts to investigate the transcription of CD7 (84). However, its sequence was not determined until 10 years later, when Russel et al. cloned Sectm1 cDNA from human erythroleukemic cells and characterized Sectm1 as a new member of the type 1a transmembrane protein family (85). The
Sectm1 gene locates ~5 kb upstream of the CD7 gene and spans ~14 kb, it encodes a 1.8kb mRNA that can be translated into a 248-amino-acid protein (85). Using immunofluorescent staining, Sectm1 was found in a perinuclear Golgi-like pattern and co-localized well with Golgi marker, Wheat Germ Agglutinin (WGA) (85). Similar to other type 1 transmembrane proteins,
Sectm1 is composed of an extracellular Ig-like N-terminus that can be cleaved and become the secreted form of Sectm1, a single conserved transmembrane domain, and a smaller intracellular
C-terminal domain (85). Although the crystal structure of Sectm1 protein has not been defined, a model for Sectm1 protein organization (Fig. 3) proposes a conserved region in the extracellular
23 N-terminal domain of 100 residues (spans residue Trp-33 to Val-133), which contained a highly conserved amino acid motif of G113Y115W117L119G121Q123 and 2 invariant cysteine residues (Cys-
38 and Cys-55). Unlike the disulfide bridge between 2 beta- sheets observed in classical Ig domains (Fig. 3A), an atypical disulfide bridge that connects a beta-strand and a loop may be Figure 3. Structure of classic Ig-domain (A) and predicted Sectm1a formed due to the close proximity Ig-like domain (B). Adapted from Laurent et al., 2015 (Ref. 86) of the 2 cysteines in amino acid sequence of Sectm1 protein (Fig. 3B) (86).
Sectm1 is widely expressed in various human organs including spleen, brain, GI track, lung and testis, with the highest expression in spleen. Within that organ, gene expression of
Sectm1 is found in epithelial cells and leukocytes of the myeloid lineage, with strongest expression in granulocytes but not detectable in lymphocytes (T and B cells, megakaryocytes)
(85, 87, 88). Additionally, different tumor cell lines also express Sectm1: a very low level of expression in colon carcinoma cell line (HT29) and cervical carcinoma cells (HeLa); whereas ovarian (OVCA 420) and breast (SKBR3 and ZR75-1) cancer cell lines showed much higher levels (85). Sectm1 gene expression in human thymic epithelial cells (TEs) increases in response to IFN-γ (88). Bioinformatics analysis revealed multiple putative binding sites for STAT1α/GAS,
STAT3, ISRE, and NF-κB, all transcription factors involved in the control of gene expression during inflammation, on Sectm1 promoter region (89). Likewise, IFN molecules upregulate
Sectm1 gene expression in human monocytes in a time dependent manner, reaching a maximum at 6 h and 12 h following IFNα, and IFNβ or IFNγ treatment, respectively (89). Interestingly,
24 treatment with LPS inhibited Sectm1 mRNA expression in human monocytes (89). Moreover, gene expression levels of Sectm1 was significantly higher in sepsis survivors when compared to non-survivors or patients with systemic inflammatory response syndrome (SIRS) (90).
Considering 1) chromosomal location and its neighboring genes; 2) the structure characteristics that resemble other Ig protein such as growth factor and cytokines; and 3) the expression profile in certain immune and cancer cells, Sectm1 is very likely to play important roles in hematopoietic and immune system processes.
I.2.B. Functional Roles of Sectm1 in Immunity
The first study investigating the functions of Sectm1 identified CD7 as a cognate of the
Sectm1 protein in 2000 (91). Stewart et al. generated a soluble version of Sectm1 by fusing an
Fc portion of human IgG1 to the extracellular domain of Sectm1 (Sectm1-Fc), and by using Flow
Cytometry and precipitation experiments, they detected high levels of binding of the Sectm1-Fc protein to both human T and NK cells through CD7. In addition, the authors cloned the mouse homolog of Sectm1, which was located on the chromosome 11 near the mouse CD7 gene, and it could bind to only mouse, but not human, CD7 protein (91). Since CD7 is primarily expressed on T and NK cells (92), they next sought to determine the effect of Sectm1 on T cell function.
Using the mouse Sectm1-Fc protein, they showed that Sectm1-Fc protein was able to suppress concanavalin A (ConA)-induced, but not anti-TcRα/β-induced, proliferation of mouse lymph node T cells in a dose-dependent manner (91). Similarly, Gordon et al. generated the recombinant human Sectm1 protein, which can bind strongly to soluble human CD7 (88).
Moreover, they found that Sectm1 gene was also expressed in human thymic epithelial cells
(TEs) and thymic fibroblasts, but not in human thymocytes; and the gene expression of Sectm1
25 in TEs could be upregulated by IFN-γ (88). In 2012, Tao et al. demonstrated that recombinant human Sectm1 protein can strongly induce CD4 and CD8 T cell proliferation and production of
IFN-γ and IL-2, and such effects synergized with anti-CD28 treatment (93). Interestingly, the authors found strong expression of Sectm1 gene in monocytes and immature monocyte-derived dendritic cells (imMoDCs) induced by IFN-γ in STAT1-dependent manner (93). Furthermore, treatment of KG1a (CD7+ acute myeloblastic leukemia cell line) with recombinant Sectm1-Fc protein suppresses the protein expression of GM-CSF through inhibition of PI3K-Akt pathway, suggesting that the binding of Sectm1 to CD7 may also act as a suppressor of specific cellular processes (94). In contrast, high levels of Sectm1 protein (both full length and soluble form cleaved from N-terminal) were detected in melanoma tissue and sera of metastatic patients; and binding of Sectm1 protein to CD7 could activate PI3K-Akt pathway in monocytes, resulting in significant increase in monocyte migration (95). Therefore, as a natural ligand for CD7, Sectm1 shows important potential in regulating immune cells (e.g. T and NK cell proliferation, monocyte recruitment, or macrophage activation) and cytokine secretion (e.g. IFNγ or IL-2).
I.2.C Mouse Homologs Sectm1a and Sectm1b
Stewart and colleagues first cloned the mouse homology of Sectm1 protein, which was shown to bind to only mouse, but not human CD7, and it could inhibit ConA-induced, but not anti-TcRα/β-induced, mouse T cell proliferation (91). This homolog was later designated as
Sectm1b by Duncan et al, as they had identified another Sectm1 homology in mouse genome,
Sectm1a (96). Alignment of the amino acid sequences showed greatest homology between human Sectm1 and mouse Sectm1a in their extracellular domain. Sectm1a and Sectm1b are ubiquitously expressed but with different tissue distribution, being the highest in the intestines
26 (96). Interestingly, Sectm1a could compete with Sectm1b for CD7 binding, yet they elicited opposite effects: Sectm1a enhanced CD4+ T cell proliferation and IL-2 production, whereas
Sectm1b inhibited TcR-mediated T cell activation (96). Surprisingly, actions of both Sectm1a and Sectm1b were CD7-independent. This might be attributed to the ability of Sectm1a proteins to bind to other receptors on cell surface such as glucocorticoid-induced TNFR-related protein
(GITR) (96).
Similar to the findings from previous studies showing that Sectm1 was an “early response gene” in monocytes (89), Hirofumi and his colleagues found that Sectm1 was upregulated during the early stage of pneumococcal pneumonia in patients, and type 1 IFN signaling (IFNα and
IFNβ) was necessary and sufficient to induce Sectm1 gene expression. This induction was mediated by signal transducer and activator of transcription 1 (STAT1) pathway and independent of NF-κB RelA signaling (87). The authors also found that recombinant mouse Sectm1a-Fc protein preferentially bound to myeloid cells, particularly Ly6Gbright and CD11bbright neutrophils, in infected mouse lungs. Importantly, recombinant Sectm1a protein did not bind to neutrophils in uninfected lungs. In addition, binding of Sectm1a recombinant protein to neutrophils augmented the expression of CXCL2, a neutrophil-attracting chemokine, which suggested that Sectm1a may act as a key player to sustain the positive feedback loop to recruit more neutrophils and amplify the inflammatory response during pneumococcal pneumonia (87).
Over all, these studies have clearly indicated that Sectm1 (and mouse homologs, Sectm1a and Sectm1b) is important regulator in immune responses by regulating T cell and neutrophil activation. Nonetheless, the functional roles of Sectm1a in macrophage function have yet to be defined.
27 Section 3: Role of Liver X Receptor in Macrophages
I.3.A. Overview of Anti-Inflammatory Functions of LXR
Nuclear receptors are intracellular transcription factors that are expressed in a wide variety of cells. They can interact with DNA directly to regulate many biological processes including cardiovascular function, immune response, and lipid metabolism of organism (97, 98). The Liver
X Receptors (LXRα and LXRβ, encoded by Nr1h3 and Nr1h2 genes, respectively) belong to the adopted orphan nuclear receptor family, and they have been identified as critical regulators linking inflammation and lipid metabolism (99). LXR genes were first cloned from mice, showing extensive sequence homology between the 2 isotypes (100). While LXRβ is ubiquitously expressed, LXRα is dominantly expressed in metabolic active tissue and cell types such as liver and macrophages (101, 102). LXRs bind to retinoid X receptor α (RXRα) to form obligate heterodimers that are able to recognize a specific DNA sequence called LXR response element (LXRE) (105). Endogenous ligands for LXRs include cholesterol derivatives (i.e. oxysterols), intermediate precursors in cholesterol biosynthesis pathway (i.e. desmosterol), and synthetic agonists that have been developed (e.g. GW3965 and T0901317) (106, 107). Upon ligand binding, LXRs undergo conformational change that causes the release of co-repressors (i.e.
SMRT and NCoR) and recruitment of co-activators (i.e. EP300 and ASC2), resulting in upregulation of target genes (104, 108, 109).
Well established functions of LXRs on lipid metabolism include: 1) facilitating cholesterol export from cells such as macrophages to high-density lipoprotein (HDL) particles through upregulation of ATP-binding cassette transporters (i.e. ABCA1, ABCG1) and ApoE (110-112); and 2) directly regulating genes involved in fatty acid synthesis such as fatty acid synthase
(FASN) and sterol regulatory element binding protein 1c (SREBP1c) (113, 114).
28 Apart from regulating cholesterol and fatty acid homeostasis, LXRs have robust anti- inflammatory activity (110, 115). For example, LXR activation prevents bacterial-induced apoptosis and promotes phagocytic responses on macrophages (116-118), whereas LXRα deficiency causes higher intracellular bacterial growth (119). In addition, LXRs are also beneficial against viral infection because of their important roles in regulation of fatty acid and cholesterol synthesis, which are two major metabolic pathways that support viral replication
(115). Thus, suppression of HIV replication upregulates the LXR target gene ABCA1, a membrane cholesterol transporter, in humanized mice (generated by transplantation of fetal liver derived CD34+ hematopoietic stem cells) (120); whereas pre-treating human macrophages with
T0901317 (synthetic LXR agonist) lowers the susceptibility of macrophages to HIV infection, possibly due to reduced lipid rafts and upregulation of ABCA1 (121).
LXR activation also reduces inflammatory responses in several macrophage cells. For example, pre-treatment of Kupffer cells (resident macrophages in liver) with a LXR agonist reduces the production of pro-inflammatory cytokines induced by LPS, and increases the levels of the anti-inflammatory cytokine IL-10 (122). Furthermore, LXRs downregulates the expression of proinflammatory cytokine IL-18 while upregulates IL-18BP, a potent IL-18 inhibitor, in murine BMDMs (123). Mechanistically, the anti-inflammatory effects of LXRs may be mediated by various mechanisms such as inhibition of NF-κB pathway or modification of plasma membrane composition to disrupt inflammatory signal transduction (99, 102, 122, 124, 125).
Furthermore, activation of toll-like receptors can inhibit LXR function (126), and animal model with LXR deficiency are more susceptible to LPS- or bacterial-induced illness, which can be rescued by LXR agonist, such as GW3965 (116, 118).
29 Lastly, LXRs modulate immune responses via affecting other immune cell types aside from macrophages. For instance, treatment with a LXR agonist in a mouse model of sterile peritonitis markedly reduced neutrophil recruitment mediated by downregulation of critical genes for leukocyte adhesion (i.e. ICAM1, VCAM1, and CCL5) and migration (i.e. IL-1β, CXCL1-3)
(127). In addition, LXR activation reduces neutrophil chemotactic and killing abilities in vitro. In keeping with this, mice treated with LXR agonist after CLP surgery (cecal ligation and puncture, a model of polymicrobial sepsis) exhibited reduced neutrophil infiltration at the infectious foci and increased systemic inflammation and mortality (128). Interestingly, neutrophils isolated from septic patients showed increased ABCA1 gene expression and impaired chemotactic response toward CXCL8 stimulation (128). Furthermore, LXR agonism stimulates cytotoxic T lymphocytes (CTLs) in mice and human, leading to robust suppression of tumor growth and progression (129). This beneficial effect is mediated through the activation of LXR-ApoE axis in circulating myeloid-derived suppressor cells (MDSCs), a type of immunosuppressive innate cell abundant in cancer patients (129).
I.3.B. Effects of LXR in Cardiac Function
Studies have shown that LXR activation provides protection in the pathogenesis of cardiac dysfunction and progression of heart failure, either by directly acting on the heart itself or through amelioration of concurrent co-morbidities. The direct effects of LXR on heart include alleviation of cardiac inflammation and preservation of cardiomyocyte viability and function. For instance, pre-treatment of H9c2 cells (an embryonic cardiomyocyte cell line, or myoblast) with the LXR agonist T0901317 effectively decreased ROS production, rescued mitochondrial membrane potential, and ultimately reduced apoptosis after high glucose challenge (130);
30 whereas pre-treatment with T0901317 in an CLP-induced acute myocardial injury model significantly ameliorated cardiac inflammation, reduced cardiomyocyte death, and improved cardiac function. The beneficial effects of T0901317 was mediated through SIRT1 (silencing information regulator 1) pathway, which enhanced FoxO1 (anti-oxidative response) and HSF1
(anti-ER stress) signaling while inhibiting NF-κB (anti-inflammation) and P53 (anti-apoptotic) pathways (131).
Given that LXRs are essential regulator of lipid metabolism and immune activity, the functional roles of LXRs have been extensively studied in various conditions that have a major impact on heart failure pathogenesis, including atherosclerosis, diabetes, and hypertension (110,
132).
LXR and atherosclerosis - The initial stage in the development of atherosclerosis involves the recruitment of macrophages to the arterial wall that are responsible for removing excess oxidized low-density lipoprotein (LDL). However, uncontrolled uptake of LDL by these macrophages will stimulate inflammatory responses and result in the formation of foam cells (133). As mentioned previously, LXRs are well-known for their ability to limit pathogenic accumulation of cholesterol by stimulating the reverse cholesterol transport (RCT) pathway, mediated mainly by upregulation of genes involved in cellular cholesterol efflux (i.e. ABCA1, ABCG1, and ApoE), plasma lipid transport (i.e. CETP and PLTP), entero-hepatic sterol absorption and excretion (i.e.
ABCG5 and ABCG8), and bile acid excretion (i.e. Cyp7a1) (110, 134). Therefore, this has sparked great interests in the therapeutic potential of LXR agonists to treat atherosclerotic cardiovascular diseases. Indeed, mice lacking both LXRα and LXRβ exhibit increased foam cell formation, further indicating the necessity of LXR signaling in maintaining cholesterol homeostasis (135). In keeping with this, administration of LXR agonists to Ldlr-/- and Apoe-/-
31 mice, which exhibit increased susceptibility to atherosclerosis, significantly slows the progression and even promotes the regression of the disease (135, 136). These results suggest that LXR activation in macrophages is necessary and sufficient to reduce atherosclerosis.
LXR and diabetes - Activation of LXR pathways with synthetic agonists on well-established mouse models of type 2 diabetes (db/db or high fat diet fed mice) provides remarkable anti- diabetic benefits, including lowering hyperglycemia and improving insulin sensitivity. Several mechanisms have been proposed: 1) by increasing insulin secretion from pancreatic β-cells (137,
138); 2) by upregulating of GLUT1 and GLUT4, which are major glucose transporters under basal and insulin-stimulated conditions (139, 140); 3) by increasing metabolic activity in brown adipose tissue (141); 4) by inhibiting of hepatic glucose production through downregulation of key gluconeogenic genes such as phosphoenolpyruvate carboxykinase (PEPCK) (139, 142); and
5) possibly through the anti-inflammatory functions of LXR, since chronic low-grade inflammation is the major contributor to the pathogenesis of diabetes (99, 102, 122-125).
However, the lipogenic effects of LXR agonists have limited their potential for clinical use on diabetes, because LXR activation can cause increased hepatic and muscle lipid accumulation, which would further aggravate the lipogenic pathology in diabetes (143). In addition, chronic activation of LXR pathway may cause lipotoxicity-induced pancreatic β-cell death, leading to further decline of insulin secretion (144).
LXR and hypertension - The renin-angiotensin-aldosterone system (RAAS) is the critical regulator of blood volume, fluid balance and systemic vascular resistance, and LXR has been implicated to control the expression of key genes involved in this pathway. Acute administration of LXR agonist T0901317 in vitro or in vivo regulates renin transcription (103, 145); whereas chronic activation of LXR inhibited renin, angiotensin-converting enzyme (ACE), and
32 angiotensin type 1 receptor (AT1R) in kidney and heart (146). Moreover, LXRα upregulation increased the gene expression of natriuretic peptides (ANP and BNP), which is another regulatory mechanism of RAAS (147). When coping with pathophysiological stimuli such as hypertension, cardiomyocytes undergo hypertrophic growth in order to maintain or enhance contractile function. Interestingly, there is significant increase of LXRα protein abundance in myocardium with pressure overload (148). In vitro and in vivo experiments using LXR agonists demonstrated decreased cardiomyocyte cell growth induced by different stimuli (i.e. Ang II or
LPS) leading to cardiac hypertrophy in WT mice (149, 150), whereas LXR-KO mice exhibit exacerbated hypertrophic responses (148, 150). Overall, LXR pathway has been demonstrated to affect many cardiometabolic traits, and beneficial effects may be patient-specific and disease state-oriented, and further investigation should focus on developing highly specific and selective ligands that minimize side effects (i.e. decreasing lipogenesis).
33 Chapter II: Materials and Methods
Section 1. Generation of Sectm1a-Knockout (KO) Mouse Models
All animal experiments conformed to the Guidelines for the Care and Use of Laboratory
Animals prepared by the National Academy of Sciences, published by the National Institute of
Health, and approved by the University of Cincinnati Animal Care and Use Committee. Mice and rats used in this study were maintained and bred in the Division of Laboratory Animal
Medical Services at the University of Cincinnati Medical Center. All animals were housed under a 12-hour light-dark cycle at constant temperature (23oC) and given regular chow diet (unless specified elsewhere).
The global Sectm1a-KO mouse model was generated using the CRISPR/ CAS9 system in mice of C57BL/6 background by the Division of Developmental Biology at Cincinnati
Children’s Hospital Medical Center. Two guideRNAs (gRNAs) targeting to Exon 3 were selected to inject with
Cas9 mRNA into one- cell embryos. The protospacer adjacent motifs (PAMs) of each Figure 4. CRISPR/CAS9 cassette for Sectm1a-KO mouse model gRNA targeting site are highlighted in bold. The spacer sequences are underlined. The cutting sites of Cas9 are indicated by arrows (Fig. 4). Because of the Cas9 activity, sequence of 88-bp between the gRNA targeting sites were deleted. At 21-day postpartum pups were tailed clipped
(~3mm) for genotyping to verify the absence of targeted gene sequence. DNA was extracted from tail using the Protease Plus in lysis buffer (Bimake, #B40015) following manufacturer’s
34 protocol. We performed routine genotyping by polymerase chain reaction (PCR) with the use of following primers: 5'-CATTCTCTCCATACAGGCTGG-3 (forward); 5'-
CTTGAACTTGGAGCTCCC AC-3 (reverse). The overall PCR mix contains tail DNA, primers, and 2xM-PCR OPTITM mix, which include optimized Taq DNA polymerase, dNTPs, MgCl2, and reaction buffer. The PCR protocol is detailed in Table 1 (below).
Table 1. PCR protocol for Sectm1a-KO genotyping
Segment Number of Temperature Duration Cycles 1 1 94oC 5 minutes
94oC 20 seconds
2 35 58oC 30 seconds
72oC 30 seconds
3 1 72oC 5 minutes
4 1 12 oC Until running gel electrophoresis
In addition, we performed quantitative real time PCR (qRT-PCR) using spleen tissue to further test whether Sectm1a gene was successfully knocked out and whether such a knockout affected Sectm1b expression. The primer sequences for measurement of Sectm1a and Sectm1b gene expression are listed in Table 2 (below).
Table 2. Primers for measurement of Sectm1a and Sectm1b gene expression
Gene Forward Reverse
Sectm1a 5’-CAGTGATGACCTGTAACATCTC-3’ 5’-CAAGTATATCCCTGTGTGGTCG-3’
Sectm1b 5'-GAGAAGCAGGTAAGAAGCTGGAG-3' 5'-CAGTTCACACCGAAGAACCC-3'
35 GAPDH 5’-TGCACCACCAACTGCTTAGC-3’ 5’-GGCATGGACTGTGGTCATGAG-3’
Section 2. Mouse Model of Acute Inflammation Induced by Endotoxemia
Male mice of 8-10 week of age were intraperitoneally (i.p.) injected with lipopoly- saccharides (LPS, from Escherichia Coli O111:B4, Sigma-Aldrich, #L2630) at a dose of 10 mg/kg of body weight (BW). Age- and sex-matched mice injected with sterile PBS were used as control. The survival rate of wild type (WT) and Sectm1a-KO mice after LPS injection were monitored every 6 h for a 72-hour (h) period. Serum samples were collected at different time points: 0 h, 3 h, 12 h, or 24 h after LPS injection for cytokine measurements using ELISA assays.
Heart tissues were collected at 12-hour time point for ELISA and Flow Cytometry analysis.
Cardiac function was assessed in mice by echocardiography 12 h after LPS injection. For LXR agonist experiment, WT and Sectm1a-KO mice were injected with the LXR agonist GW3965
(MedKoo, #522685, 30 mg/kg of BW, once daily, i.p.) for 3 days, DMSO-injected mice were used as control. Six hours after the last injection of GW3965, all mice received 1 injection of
LPS (10 mg/kg of BW), and cardiac function was measured by echocardiography at 12 h post-
LPS injection.
Section 3. High Fat Diet (HFD)-Induced Chronic Inflammation Model of Obesity
Starting at age of 5 or 6 weeks, male WT and Sectm1a-KO mice were given ad libitum access to HFD (Research Diet, #D12492, 60% kcal from fat, 20% kcal from protein, and 20% kcal from carbohydrate) for 18-24 weeks. Heart samples were collected for Flow Cytometry analysis after 5 weeks of HFD feeding. Echocardiography was performed to assess cardiac function when mice were fed with HFD for 20 weeks.
36 Section 4. Cell Isolation, Culture, and Treatments
II.4.A Culture of cell lines
L929 fibroblast cell line was purchased from American Type Culture Collection
(ATCC®CCL-1TM) and cultured in normal complete medium (DMEM supplemented with 15%
FBS, 1% penicillin/streptomycin), cell culture supernatant was collected after 10 days of culture and centrifuged at 750g for 10 min. Supernatant was then stored at -20oC until use. HEK293T cell line and mouse macrophage RAW264.7 cell line were purchased from ATCC and cultured
o in normal complete medium. All cells were grown at 37 C with 5% CO2 in fully humidified air.
II.4.B Isolation of bone marrow-derived macrophages (BMDMs)
Bone marrow-derived macrophages were isolated and cultured as described previously
(14, 151). WT and Sectm1a-KO mice were anesthetized by i.p. injection of Ketamine (90 mg/kg
BW) and Xylazine (20 mg/kg BW) followed by removal of both hind legs (tibias and femurs).
Skin and muscle were removed and bones were washed twice in ice-cold PBS. By using 25G needles filled with cold sterile wash medium (DMEM without calcium and magnesium), bone marrow was flushed out and filtered through 70 µm Nylon cell strainer. Then cells were centrifuged at 500g for 5 min at room temperature (RT), followed by resuspension of the cell pellet in red blood cell (RBC) lysis buffer for 5 min, and then centrifuged at 500g for 5 min again.
The resulting cell pellet was resuspended in complete BMDM medium (DMEM supplemented with 15% L929 cell culture supernatant, 10% FBS, 1% penicillin/streptomycin). Cells were allowed to grow and differentiate for 7 days before being used for experiments.
37 II.4.C Isolation of adult rat cardiomyocytes (ARCMs)
Adult rat cardiomyocytes were isolated from 6-week old male Sprague-Dawley rats
(purchased from The Jackson Laboratory) as described previously (152, 153). After rat was anesthetized by i.p. injection of Ketamine (80 mg/kg BW) and Xylazine (10 mg/kg BW), the heart was excised and perfused with modified Krebs-Henseleit buffer (KHB, includes: 118 mM
NaCl, 4.8 mM KCl, 25 mM Hepes, 1.25 mM K2PO4, 1.25 mM MgSO4, 11 mM glucose, 5 mM taurine, and 10 mM BDM, pH 7.4) for 5 min. The aorta was cannulated and the heart was mounted on a Langendorff apparatus, followed by perfusion with digestive solution (0.7 mg/ml collagenase type II, 0.2 mg/ml hyaluronidase, 0.1 % BSA, 25 µM CaCl) for 20 min. During perfusion, Ca concentration was gradually increased to 0.1 mM. Next, ventricular tissue was minced, pipette-dissociated, and filtered through a nylon mesh (200 µm). Cells were centrifuged and resuspended in ACCT medium (DMEM supplemented with 15% FBS, 2 mM L-carnitines, 5 mM creatine, 5 mM taurine, and 1% penicillin/streptomycin), then cells were counted and plated on laminin (10 µg/ml)-coated 6-well plates overnight.
II.4.D Construction and infection of Sectm1a adenovirus vector
The adenovirus expressing Sectm1a with a green fluorescent protein (GFP) probe
(Ad.Sectm1a) was constructed by SignaGen Laboratories by inserting full length Sectm1a cDNA into pCMV-shuttle vector, then cloned into an adenoviral backbone plasmid. Plated ARCMs in
6-well plates were infected by adenoviruses (Ad.Sectm1a or Ad.GFP) at 100 MOI for 2 h, then cells were cultured in normal complete DMEM medium for 24 h. Cardiomyocytes were then treated with LPS (50 ng/ml) for 3 h, and cells were the harvested for qRT-PCR analysis.
Cardiomyocytes infected with Ad.GFP were used as control. To overexpress Sectm1a in macrophages, BMDMs were allowed to grow for 5 days followed by infection with Ad.Sectm1a
38 adenovirus (Ad.GFP-infected BMDMs used as control) at 500 MOI for 48 h. Then BMDMs were treated with LPS (10 ng/ml) for 8 h. Cell culture supernatants were then collected for cytokine measurement using ELISA kits.
II.4.E Treatments on macrophages
RAW264.7 macrophages or BMDMs were plated in 6-well plates (3x105 cells/well seeding density) or 24-well plates (5x104 cells/well seeding density) and allowed to adhere for 24 h. After 2 washes with PBS, cells were treated with LPS (from Escherichia coli O111:B4,
#L4391, Sigma-Aldrich) or palmitate (Sigma-Aldrich, #P0500) at indicated concentrations and time points specified in Figures/Results. PBS or endotoxin-free BSA were used as control, respectively. Then, cell culture medium were collected for cytokine measurements, and cells were collected for gene expression, Western-blotting, or Flow Cytometry assays.
To investigate the effects of LXR agonist on Sectm1a-KO macrophage function, WT and
Sectm1a-KO BMDMs were first treated with LXR agonist GW3965 (Tocris, #2474, 2 µM) for
12 h, BMDMs treated with same volume of DMSO were used as control. These macrophages were then treated with LPS (10 ng/ml) for total 48 h, cell culture medium was collected at various time points for cytokine measurements using ELISA.
Section 5. In Vivo Assessment of Cardiac Function
Cardiac function was measured using trans-thoracic echocardiography with Vevo®2100 ultrasound imaging system (Visualsonics, Toronto Canada) equipped with a MS400 probe (30-
MHz centerline frequency) as described previously (154). Mice were anesthetized by isoflurane
(1.5-2%), images were obtained from parasternal long axis and short axis views at depth between
1 and 13 mm in M-mode. Left ventricular cavity size and wall thickness were analyzed with
39 Vevostrain software (Vevo 2100, v1.1.1 B1455), LV ejection fraction (EF) was calculated as:
[(LVDd3 – LVDs3)/LVDd3] ×100. All measurements were performed according to the American
Society for Echocardiography leading-edge technique standards, and averaged over at least three consecutive cardiac cycles. (LVDd, left ventricular diameter at diastole; LVDs, left ventricular diameter at systole).
Section 6. Gene Expression and Protein Measurements
II.6.A qRT-PCR experiments
Total RNA was extracted from cultured cells, whole blood, or tissue samples using the miRNeasy Mini kit (Qiagen, #1038703) in accordance with the manufacturer’s instructions. The quality and concentration of RNA was assessed by optical density using a NanoDrop 2000 system (ThermoFisher Scientific). cDNA was synthesized from 1.0 μg RNA using Superscript II
Reverse Transcriptase (Invitrogen, #8080044). Then qRT-PCR was performed in triplicate with the ABI PRISM 7900HT sequence detection system (ABI) using SYBR green (Denville
Scientific, #CC1731). Relative mRNA levels were calculated using the 2-ΔΔCt method using
Rps18 gene expression as internal control. Sequences of primers used for quantitative RT-PCR were obtained from literature and are listed in Table 3 (below):
Table 3. Primers used for qRT-PCR analysis
Gene Forward Reverse
RPS18 GCAATTATTCCCCATGAACG GGCCTCACTAAACCATCCAA
LXRα CCTGATGTTTCTCCTGACTC TGACTCCAACCCTATCCTTA
LXRβ ACCAGCCCAAAGTCACGC TTGGCAAAGTCCACAATCTCC
40 ABCA1 AGTGATAATCAAAGTCAAAGGCACAC AGCAACTTGGCACTAGTAACTCTG
ABCG1 TTCATCGTCCTGGGCATCTT CGGATTTTGTATCTGAGGACGAA
ApoE ACAGATCAGCTCGAGTGGCAAA ATCTTGCGCAGGTGTGTGGAGA
IL-1RN GCTCATTGCTGGGTACTTACAA CCAGACTTGGCACAAGACAGG
CD36 GATGACGTGGCAAAGAACAG TCCTCGGGGTCCTGAGTTAT
MMP12 AATGCTGCAGCCCCAAGGAAT CTGGGCAACTGGACAACTCAACTC
IL-6 CTGCAAGAGACTTCCATCCAG AGTGGTATAGACAGGTCTGTTGG
IL-1β GCAACTGTTCCTGAACTCAACT ATCTTTTGGGGTCCGTCAACT
II.6.B RNA-sequencing analysis
Directional RNA sequencing (RNA-seq) analyses were performed on WT and Sectm1a-
KO BMDMs (n=3 for each genotype) by the Genomics, Epigenomics and Sequencing Core
(GESC) at the University of Cincinnati. Total RNA was extracted as mentioned in previous section, and its total integrity was assessed by Bioanalyzer (Agilent, Santa Clara, CA). PolyA
RNA purification was performed using NEBNext Poly(A) mRNA Magnetic Isolation Module
(New England BioLabs, Ipswich, MA) with 1 µg of total RNA as input, and SMARTer Apollo
NGS library prep system (Takara, Mountain View, CA) was used for automated polyA RNA isolation. NEBNext Ultra II Directional RNA Library Prep kit (New England BioLabs) was used to prepare the library for RNA-seq. After indexing via PCR (12 cycles) enrichment, the amplified libraries and the negative control were cleaned by Pure XP beads for QC analysis, and the quality was determined by Bioanalyzer using DNA high sensitivity chip. To measure differential gene expression, individually indexed and compatible libraries were proportionally pooled (~25 million reads per sample) for clustering in cBot system (Illumina, San Diego, CA).
41 Libraries (15pM) were clustered onto a single read (SR) flow cell using Illumina TruSeq SR
Cluster kit v3, and sequenced to 51-bp using TruSeq SBS kit on Illumina HiSeq system.
II.6.C Western-blotting experiments
Total proteins were extracted from cultured BMDMs using the NP40 lysis buffer
(ThermoFisher Scientific, #FNN0021) supplemented with phenylmethylsulfonyl fluoride (PMSF, purchased from ThermoFisher Scientific, #36978, 0.1mM) and complete protease inhibitor cocktail (Roche Applied Science, #5892970001). Cell lysates were then centrifuged at 14000g for 15 min at 4oC. Protein concentration was determined using Biorad Protein Assay Reagent
(Biorad, #5000006). Protein samples were separated by 10%-12% SDS-PAGE and transferred to
0.2 µm nitrocellulose blotting membrane followed by incubation in blocking buffer (5% non-fat milk) for 1 h. The membrane was then incubated in primary antibody (listed in Table 4) at 4oC overnight. After washing with Tris-buffered saline (TBS, 100 mM Tris, 0.9% NaCl, pH7.4), the membrane was incubated in peroxidase-conjugated secondary antibody for 1 h at room temperature. After washing with TBS, binding of the primary antibody was detected by adding
HyGLO chemiluminescent detection reagent (Denville Scientifc, #E3212) or SuperSignal West
Femto Maximum Sensitivity Substrate (ThermoFisher Scientific, #34096). Western blot bands were quantified by MultiImage II (AlphaInnotech, USA) or ImageJ software.
Table 4. Antibodies used for Western Blotting experiments
Name Manufacturer Catalog #. Dilution
p65 Cell Signaling Tech. 8242 1:1000
Phosphorylated p65 (S536) Cell Signaling Tech. 3033s 1:1000
IkBα Santa Cruz SC371 1:1000
42 Phosphorylated IkBα (S32/36) Cell Signaling Tech. 9246s 1:1000
GAPDH Cell Signaling Tech. 97166S 1:1000
Section 7. Cytokine Measurement using ELISA Assays
Whole blood samples were collected by cardiac puncture with heparinized needles at indicated time points after LPS injection, and spun down at 4000 rpm for 10 min. Heart samples were homogenized in NP40 lysis buffer containing PMSF and protease inhibitor cocktail as described in Section 6, protein samples were then exacted and stored in -80oC. Cell culture supernatants from macrophages were harvested at different time points with different treatment schemes. The concentrations of TNFα, IL-1β, Il-6, and MCP-1 were determined in duplicates by commercial available ELISA kits [BioLegend, #430901 (TNFα), #432601 (IL-1β), #431301 (IL-
6), #432702 (MCP-1)], according to the manufacturer’s protocol.
Section 8. Immunofluorescent Staining
Bone marrow-derived macrophages from WT and Sectm1a-KO mice were seeded in 24- well plates (5x104 cells/well) for 24 h. These BMDMs were pretreated with GW3965 (2 µM) for
12 h followed by LPS treatment (10 ng/ml) for 30 min. Then cells were washed with PBS and fixed in 4% paraformaldehyde (PFA) for 20 min. Heart samples were harvested after perfusion with PBS and immediately fixed in 10% neutral buffered formalin (Sigma-Aldrich, #HT501128) at 4oC for at least 48 h, then they were embedded in paraffin and sliced at 5 µm of thickness. The sections were deparaffinized in xylene (ThermoFisher Scientific, #X3P-1GAL) and rehydrated through graded ethanol (100%, 95%, 70%, 50%). After rinsing with distilled water, heat-
43 mediated antigen retrieval with sodium citrate buffer (0.01M, pH6.0, 95 oC) was performed for
15 min. Next, BMDM samples or heart sections were incubated in blocking solution (PBS with
1% BSA and 0.3% triton) for 1 h at room temperature, followed by incubation with primary antibodies (listed in Table 5) at 4oC overnight. After washing with PBS, samples were then incubated with secondary antibodies for 1 h at room temperature. Then BMDM and heart samples were mounted with Antifade Mountant medium (Invitrogen, #P36962). Images were captured with Zeiss LSM710 LIVE Duo Confocal Microscope (Live Microscopy Core,
University of Cincinnati).
Table 5. Antibodies used for immunofluorescent staining
Name Manufacturer Catalog # Dilution
LXRα ThermoFisher PA1-330 1:100
F4/80 BioLegend 123102 1:50
α-actinin Sigma-Aldrich A7811 1:100
Section 9. Flow Cytometry
Methods for analyzing macrophages with Flow Cytometry were adopted from previous studies with modifications (14, 155, 156). Heart tissue were minced and digested in HBSS with
Collagenase IV (2 mg/ml, Worthington, #LS004188), Dispase II (1.2 U/ml, Sigma, #D4693) and
0.9 mM CaCl2, then incubated at 37°C for 45 min. with gentle agitation. Tissues were then passed through 40 µm cell strainer followed by centrifugation at 500 g at 4 °C for 5 min. The cell pellet was then resuspended in RBC lysis buffer and incubated at room temperature for 5 min.
Then cells were washed and resuspended in Flow Cytometry buffer (Ca/Mg-free HBSS with 1
44 mM EDTA, 25 mM HEPES and 1% FBS). To collect BMDMs for cell surface marker analysis, macrophages were first washed with PBS and scrapped off the plates, after centrifugation, cell pellets were then resuspended in Flow Cytometry buffer. Next, heart samples or BMDMs were incubated on ice with Fc-blocking solution (anti-CD16/32) for 10 min. After washing, cells were stained with primary antibodies (listed in Table 6) at 4°C for 30 min in dark. Then cells were washed twice, fixed in 0.1% PFA for 15 min. The compensation matrix was determined using
UltraComp eBeads (ThermoFisher Scientific, #01-2222-42). Appropriate fluorescence minus one
(FMO) and negative controls were used to set gates. Flow Cytometry was performed using
LSRII Analyzer (SHC Flow Cytometry Core, Cincinnati), and analyzed with FCSexpress software.
Table 6. Antibodies used for Flow Cytometry
Reagent Name Manufacturer Catalog # Dilution
LIVE/DEAD ThermoFisher L34962 1:1000
CD45.2 Alexa Fluor 488 BioLegend 109816 1:100
CD45.1 BV711 BioLegend 110739 1:100
Ly6G BV421 BioLegend 127628 1:100
CD11b APC-eFluor 780 eBioScience 47-0112-80 1:100
Ly6C APC eBioScience 17-5932-82 1:100
F4/80 BV510 Biolegend 123135 1:100
MHC-II PerCP-eFluor 710 eBioScience 46-5321-82 1:100
CD206 Alexa Fluor 700 Bio-Rad MCA2235A700 1:100
CCR2 PE-Cy7 Biolegend 150612 1:100
45 CD38 PE-Cyanine 7 eBioScience 25-0381-82 1:100
CD301 PE Biolegend 145704 1:100
Section 10. Co-Immunoprecipitation Assay
HEK 293T (purchased from ATCC) cells were simultaneously transfected with 2 plasmids encoding GFP-conjugated Sectm1a (Origene, #MG201838) and HA tag-conjugated
LXRα (SinoBiological, #MG57099-CY) using Effectene Transfection Reagent (Qiagen,
#301425), according to the manufacturer’s protocol. After 48 h of transfection, cell samples were collected to detect Sectm1a-LXR interaction using PieceTM Co-Immunoprecipitation Kit
(ThermoFisher, #26149) following manufacture’s instruction. Cell lysates were harvested, and protein concentration was determined as mentioned previously. Then a total of 500 µg of protein sample was incubated with 5 µg of primary antibody (listed in Table 7) at 4 oC overnight to form the immune complex. Then the complex lysate samples were mixed with protein A/G Plus
Agarose in the spin column that has been pretreated with resin, followed by incubation for 1 h with gentle end-over-end shaking. Then the samples were washed 3 times using lysis/wash buffer and eluted into collection tube, followed by SDS-PAGE analysis as described previously in the Western Blot section.
Table 7. Primary antibodies used for co-immunoprecipitation
Name Manufacturer Catalog #. Dilution
Anti-GFP Origene TA150041 1:100
Anti-HA tag Cell Signaling Tech. 3724S 1:100
46 Section 11. Bone Marrow Transplantation
Age- and Sex-matched C57BL6 WT mice expressing CD45.1 were used as recipients.
One week before irradiation, recipient mice were given ad libitum access to water supplemented with 0.2 mg/ml enrofloxacin. Then, utilizing a Xen-X (X-Strahl, Suwanee, GA) pre-clinical cabinet irradiator calibrated using NIST-traceable instruments at the Preclinical Imaging Core
(PIC) of University of Cincinnati, recipient mice were irradiated with 1200 cGy, split into 2 fractions (600 cGy each) 4 h apart, The instrument parameters were: 220 kVp, 13mA, Cu
Filtration, 0.67 mm of Cu Half Value Layer (CuHVL), 0.857 cGY/Sec delivered using a 100 mm x 100 mm collimator at extended distances. Within 24 h following irradiation, recipient mice were retro-orbitally injected with freshly isolated bone marrow cells from donor Sectm1a-KO mice and WT mice (expressing CD45.2), isolated as described earlier in Section II.4.B. Each recipient mouse received 106 cells suspended in 150-200 μl of PBS. All mice were provided with enrofloxacin-supplemented water since the week before irradiation until sacrifice. Body weight was measured weekly. 4 weeks after irradiation, recipient mice were injected with LPS (10 mg/kg BW, i.p.) followed by echocardiographic measurement of cardiac function 12 h post injection. Last, heart samples were harvest and processed for Flow Cytometry analysis immediately after echocardiography.
Section 12. Statistical Analysis
Data were expressed as means ± Standard Error of the Mean (SEM). Sample sizes were based on our previous experience with the procedures used. Graphpad Prism (version 6) software was used for statistical analysis. Comparison between 2 groups was determined by Student t test.
Differences among multiple groups were determined by one- or two-way ANOVA where
47 appropriate. The survival rates were constructed using the Kaplan–Meier method, and differences in mortality were compared using the log-rank-test. A p<0.05 was considered statistically significant.
48 Chapter III: Results
Section 1. Expression Profiles of Sectm1a in Different Organs and BMDMs with or without
LPS Challenge
We first confirmed that Sectm1a is highly expressed in multiple tissues such as spleen
(Fig. 5). Previously, Sectm1a has been implicated to play a role in regulating immune responses through regulating T cell, NK cells, neutrophils, and monocytes (87, 89, 91,95, 96), yet whether
Sectm1a plays any role in macrophages remains unknown. To this end, we sought to determine whether the expression of Sectm1a in macrophages was altered in response to inflammatory stimuli. BMDMs were treated with IFNγ alone or in combination with LPS for 4 h. IFNγ dramatically upregulated Sectm1a expression along with a modest but statistically significant increase in TNFα expression (Fig. 6A). Interestingly, when BMDMs were treated with IFNγ and
LPS together, Sectm1a expression was significantly decreased comparing to IFNγ alone, though it was still significantly higher than PBS control. Remarkably, the gene expression of TNFα was about 10-fold higher in BMDMs treated with IFNγ and LPS, when compared to IFNγ alone treatment (Fig. 6A). Next, we aimed to determine the effects of LPS alone on Sectm1a expression. After culturing BMDMs with increasing doses of LPS for 24 h, Sectm1a gene expression was significantly reduced when comparing to PBS control group (Fig. 6B). Gene expression levels of Sectm1b exhibited a degree of reduction that did not reach statistical significance (Fig. 6C). Interestingly, a time course analysis revealed that although treatment of
WT BMDMs with LPS for 24 h reduced Sectm1a gene expression, shorter exposures (i.e. 6 h) resulted in upregulated Sectm1a expression (Fig. 6D). Given that Sectm1a is highly abundant in the spleen and the blood (Fig. 5), we next measured its mRNA levels in whole blood and spleen
49 from WT mice after LPS injection. Sectm1a levels were dramatically reduced in whole blood but increased in spleen at 1 h and 3 h post-LPS injection. However, in the blood, gene expression of
Sectm1a increased at 24 h when comparing to earlier time points, though it was still significantly lower than control group (Fig. 7A). Nonetheless, gene expression profile of Sectm1a in spleen showed opposite trend: at 24 h, mRNA levels of Sectm1a was significantly reduced, which was even lower than the control group. (Fig. 7B). Put together, these data suggest that Sectm1a may be involved in the early activation of LPS-stimulated inflammatory responses.
50 Figure 5. Tissue distribution of Sectm1a in WT mice. Expression profile of Sectm1a in various organs of WT mice were determined using qRT-PCR (n=3).
51 Figure 6. Kinetics of LPS-stimulated gene expression of Sectm1a. (A) WT-BMDMs were treated with IFNγ alone (10 ng/ml), or together with LPS (10 ng/ml) for 4 h, gene expression of
Sectm1a and TNFα were determined by qRT-PCR (n=3). (B-C) Gene expression level of
Sectm1a (B) and Sectm1b (C) was measured in WT-BMDMs treated with indicated doses of
LPS for 24 h (n=3). (D) Sectm1a gene expression was determined in WT-BMDMs treated with
LPS (10 ng/ml) for indicated time points (n=3). (*, p<0.05; data are presented as Mean ± SEM;
1-way ANOVA with Dunnett's multiple comparisons test)
52 Figure 7. Expression of Sectm1a in whole blood and spleen of WT mice after LPS treatment. WT mice were i.p. injected with LPS (10 mg/kg of BW), Sectm1a mRNA levels in whole blood (A) and spleen (B) were determined at various time points with qRT-PCR (n=3-5).
(*, p<0.05; data are presented as Mean ± SEM; 1-way ANOVA with Dunnett’s multiple comparisons test)
53 Section 2. Sectm1a Deficiency Aggravates LPS-Induced Systemic Inflammation and
Mortality
To evaluate the role of endogenous Sectm1a, we generated a global (instead of a cell/tissue-specific) knockout (KO) mouse model using CRISPR-Cas9 technology. We validated that Sectm1a, but not Sectm1b, gene expression was effectively disrupted (Fig. 8A-B). The KO mice breed normally and do not exhibit gross behavioral abnormalities when compared to WT controls. To explore the potential role(s) of Sectm1a in inflammatory conditions, we first injected both WT and KO mice with LPS (10 mg/kg of body weight, i.p.), and measured the levels of proinflammatory cytokines in plasma using ELISA assays at 12 h post-LPS injection.
Loss of Sectm1a significantly increased plasma levels of IL-6, TNFα, and IL-1β, when compared to WT-LPS group (Fig. 9A-C). Next, we injected a separate cohort of mice with LPS (10 mg/kg of body weight, i.p.). The median survival for mice from WT-LPS group was 60 h (n=10-20) during 72 h of LPS treatment. In contrast, Sectm1a-KO mice exhibited a 40% significantly higher mortality rate with a median survival of 35 h (Fig. 10). Put together, these data indicate that Sectm1a may be a pivotal mediator in the regulation of LPS-induced inflammatory responses.
54 Figure 8. Validation of Sectm1a-KO mouse model. (A) Gene expression levels of Sectm1a and Sectm1b in spleen samples from WT and Sectm1a-KO mice measured by qRT-PCR (n=3).
(B) Gel electrophoresis results using products from qRT-PCR experiments to validate that
Sectm1a-KO model was successfully generated (n=3). (*, p<0.05; data are presented as Mean ±
SEM; Student’s t test)
55 Figure 9. Systemic inflammation is increased in Sectm1a-KO mice after LPS injection. (A-
C) serum cytokine levels (A: IL-6; B: TNFα; C: IL-1β) of WT and Sectm1a-KO mice were measured with ELISA 12 h after LPS (10 mg/kg) injection (n=6-9). (*, p<0.05; data are presented as Mean ± SEM; Student’s t test)
Figure 10. Sectm1a deficiency increases LPS-induced mortality. WT and Sectm1a-KO mice injected i.p. with LPS (10 mg/kg) were monitored for survival up to 72 h post treatment (n=10-
20). (*, p<0.05; log-rank test)
56 Section 3. Ablation of Sectm1a Leads to Exacerbated Cardiac Inflammation and
Dysfunction
We then sought to investigate the role of Sectm1a on cardiac function in the context of systemic inflammatory response initiated by bacterial endotoxin. To this end, we assessed cardiac function at 12 h post-LPS injection using echocardiography. Sectm1a-KO mice exhibited normal cardiac function similar to WT controls under basal conditions (PBS injection) (Fig.
11A-B and Table 8). However, cardiac dysfunction following LPS injection deteriorated significantly more in Sectm1a-KO mice , as evidenced by 38% reduction in fractional shortening, compared to WT controls (Fig.11B and Table 8). Of note, We have not observed any structural difference between WT and Sectm1a-KO mice after LPS injection (Table 8). Given that cardiac dysfunction may be at least partially attributed to increased infiltration of immune cells into the myocardium (158), we next went on to determine the types and numbers of immune cells in
LPS-treated mouse hearts, using Flow Cytometry analysis and immunofluorescent staining. The gating strategy is depicted in Figure 12. The number of neutrophils and macrophages were dramatically and significantly increased in hearts of Sectm1a-KO mice after LPS treatment, when compared to LPS-treated WT group (Fig.13A/B, C/G, and D/H). More interestingly, the macrophages from KO-LPS hearts displayed proinflammatory phenotype with higher expression of CCR2 and MHC-II but reduced levels of CD206 (Fig. 13E/I and F/J), compared to those cells from LPS-treated WT hearts.
Immunofluorescent staining of heart samples from LPS-treated WT and Sectm1a-KO mice suggested increased macrophage accumulation in the myocardium compared to PBS- treated counterparts (Fig. 14). Accordingly, levels of TNFα, IL-6, and IL-1β were significantly higher in the myocardium of Sectm1a-KO mice than those of WT mice at 12 h post LPS
57 injection (Fig. 15A-C). Altogether, these data suggest that Sectm1a may be needed to protect against inflammatory insult in the heart and thereby preserve cardiac function.
Figure 11. Knockout of Sectm1a exacerbated LPS-triggered cardiac dysfunction. (A-B) 12 h after injecting WT and Sectm1a-KO mice with LPS (10 mg/kg of BW, mice injected with PBS were used as control), cardiac function was determined by echocardiography (A), and fractional shortening (%) was calculated (B). (n=5-9) (*, p<0.05; data are presented as Mean ± SEM; 2- way ANOVA)
58 Table 8. Echocardiographic measurements of WT and Sectm1a-KO mic 12 h after LPS injection
59 Figure 12. Gating strategy for Flow Cytometry analysis of cardiac macrophages.
60 Figure 13. Ablation of Sectm1a enhances accumulation of inflammatory macrophages in hearts from LPS-treated mice. 12 h after LPS treatment, heart samples were collected from
WT and Sectm1a-KO mice for Flow Cytometry analysis. (A-B) Representative plots and quantification of neutrophils (Ly6G+) population in the hearts (n=4). (C-J) Representative Flow
Cytometry plots and quantification of cardiac macrophage marker expression showed more macrophage accumulation (F4/80+) with inflammatory phenotype (CCR2+, MHC-II+, CD206-) in the heart of KO mice 12 h after LPS injection (n=4). (*, p<0.05; data are presented as Mean ±
SEM; Student’s t test)
61 Figure 14. Impact of Sectm1a deficiency on macrophage infiltration in the heart upon LPS challenge. 12 h after LPS injection, heart samples from WT and Sectm1a KO mice were harvested and stained with markers for cardiomyocytes (α-actinin) and macrophage (F4/80),
DNA was stained with DAPI (blue).
62 Figure 15. Lack of Sectm1a increases cardiac cytokine levels after LPS injection. (A-C)
Cytokine levels (A: TNFα; B: IL-6; C: IL-1β) in the myocardium of WT and Sectm1a-KO mice after LPS treatment were measured by ELISA (n=3-4). (p<0.05, data are presented as Mean ±
SEM, Student’s t test).
63 Section 4. Lack of Sectm1a Augments LPS-Induced Inflammation via Skewing BMDMs toward Proinflammatory Phenotype
To define the functional role of Sectm1a in LPS-stimulated macrophages in a cell- autonomous manner, we isolated BMDMs from WT and Sectm1a-KO mice. Differentiation, proliferation and morphology was similar between BMDMs from WT and Sectm1a-KO mice
(Fig 16A-B). Also, Sectm1a deficiency did not alter the basal levels of inflammatory factors, such as TNFα, IL-1β, IL-6, and MCP-1 (Fig. 17A-D). However, in LPS-treated BMDMs, lack of
Sectm1a significantly augmented the secretion of these inflammatory factors at 24 h (Fig. 17A-
D). KO-BMDMs exhibited a small, yet statistically significant, increase in IL-6 levels at 12 h time point, when compared to WT-BMDMs (Fig. 17C).
In keeping with the cytokine profile, Flow Cytometry analysis revealed 31% higher but
24% lower in the levels of CD38 (proinflammatory marker) and CD206 (anti-inflammatory marker), respectively, in KO-BMDMs at 6 h post-LPS treatment (Fig. 18A-B). These data indicate that deletion of Sectm1a skews macrophages toward proinflammatory phenotype.
We then sought to examine whether Sectm1a deficiency could promote LPS-induced NF-
κB activation in macrophages and phosphorylation of the key subunit, p65. Our Western Blotting results showed that phosphorylation of p65 was further significantly increased by 48% in
Sectm1a-KO BMDMs 30 min after LPS exposure, when compared to WT-LPS controls (Fig.
19A-B). Moreover, we observed a significant increase in IkBα phosphorylation at S32/36, a key regulator of p65 activity, in KO-BMDMs after LPS treatment, when compared to WT-LPS group
(Fig. 19A-B).
64 Conversely, overexpression of Sectm1a in BMDMs via adenovirus significantly reduced
LPS-triggered phosphorylation of p65 and IkBα (Fig. 20A-D), when compared to Ad.GFP control group with LPS treatment. Subsequently, production of inflammatory cytokines was markedly reduced in BMDMs with Sectm1a overexpression (Fig. 21A-D). Intriguingly, overexpression of Sectm1a in adult rat cardiomyocytes using the same adenovirus, failed to regulate mRNA levels of cytokines (IL-6 and IL-1β) when compared to the control group (Fig.
22A-B), which implies that Sectm1a does not regulate inflammatory response in cardiomyocytes.
Overall, these data suggest that Sectm1a deficiency enhances inflammation in macrophages through the activation of NF-κB pathway.
65 Figure 16. Ablation of Sectm1a has no effect on macrophage maturation in vitro. (A-B)
BMDMs were isolated from WT and Sectm1a-KO mice and allowed to differentiate for 7 days,
Flow Cytometry experiments were performed to validate the purity of the differentiated macrophages. Representative images of mature BMDMs (A) and Flow Cytometry results (B) showed no differences on cell morphology and maturation. (Scale bar, 200 µm; *, p<0.05; data are presented as Mean ± SEM; Student’s t test)
66 Figure 17. Knockout of Sectm1a augments cytokine release from BMDMs. (A-D) After treating BMDMs with LPS (10 ng/ml), cytokine levels: TNFα (A), IL-1β (B), IL-6 (C), and
MCP-1 (D) from cell culture supernatant were measured using ELISA at 12- and 24-h time points (n=4-6). (*, p<0.05; data are presented as Mean ± SEM; 2-way ANOVA with Sidak’s multiple comparisons test)
67 Figure 18. Absence of Sectm1a skews BMDMs toward proinflammatory phenotype.
BMDMs were cultured for 7 days and stimulated with LPS for 6 h. (A-B) Representative Flow
Cytometry plots (A) and quantification of macrophage marker expression (B) revealed stronger inflammatory phenotype, as evidenced by increased CD38+ and lowered CD206+ expression, in
Sectm1a-KO BMDMs 6 h after LPS treatment (n=3). (*, p<0.05; data are presented as Mean ±
SEM; Student’s t test)
68 Figure 19. Sectm1a deficiency activates NF-κB pathway. (A-B) Western Blotting of phosphorylated p65 and IkBα in BMDMs with or without LPS stimulation (10 ng/ml, 30min.) n=3 dishes of BMDMs for isolation of proteins. (*, p<0.05; data are presented as Mean ± SEM;
2-way ANOVA with Sidak’s multiple comparisons test).
69 Figure 20. Overexpression of Sectm1a in BMDMs suppresses NF-κB pathway. (A-B)
Representative images of BMDMs infected with adenovirus encoding Sectm1a (or GFP as control), and qRT-PCR result validated that the overexpression of Sectm1a was successful (n=3).
(C-D) Western Blotting and quantification of phosphorylated p65 and IkBα in BMDMs treated with LPS after infection with adenovirus (n=3 dishes of BMDMs for isolation of proteins). (*, p<0.05, data are presented as Mean ± SEM, 2-way ANOVA with Sidak’s multiple comparisons test).
70 Figure 21. Overexpression of Sectm1a reduces cytokine production in BMDMs. (A-D)
Concentration of cytokines: TNFα (A), IL-1β (B), IL-6 (C), and MCP-1 (D) from cell culture supernatant were determined by ELISA (n=3-5). (*, p<0.05, data are presented as Mean ± SEM,
2-way ANOVA with Sidak’s multiple comparisons test).
71 Figure 22. Sectm1a upregulation does not affect cytokine gene expression in ARCMs. (A-B)
After infecting adult rat cardiomyocytes with adenovirus followed by LPS treatment, gene expression of IL-6 (A) and IL-1β (B) in cardiomyocytes were measured by qRT-PCR (n=3). (*, p<0.05, data are presented as Mean ± SEM, 2-way ANOVA).
72 Section 5. Gene Enrichment Analysis of Sectm1a-KO BMDMs
To gain insights on potential mechanisms underlying the aberrant inflammatory responses in Sectm1a-KO macrophages, we performed RNA sequencing analyses of the gene expression profile in BMDMs isolated from WT and Sectm1a-KO mice (Fig. 23A). Among the
714 upregulated and 746 downregulated genes in Sectm1a-KO BMDMs (Fig. 23B), 75 differentially expressed genes are involved in cytokine-cytokine receptor interaction and chemokine signaling pathways (Fig. 23C). Interestingly, many of the most significantly downregulated genes are directly or indirectly regulated by LXR signaling pathway, such as
ApoE, Plin2, IL-1RN, Cebpα, and ABCA1 (Fig. 24A-B). Further analysis revealed that 100
LXR-related genes were differentially expressed in Sectm1a-KO BMDMs (Fig. 24C). Consistent with the RNA-seq data, qRT-PCR validated significant decreases in the expression of several noted LXR-targeted genes: ApoE, ABCA1, ABCG1, CD36, and MMP12 (Fig. 24D).
More intriguingly, gene expression of LXRα itself was significantly reduced in KO-
BMDMs, when compared to WT-macrophages, while LXRβ levels exhibited no difference between two groups (Fig. 25A). In addition, gene network analysis identified that three genes
(IL-1RN, Cav1, S100a8) were involved in sepsis, cardiovascular diseases and LXR signaling cascade, and IL-1RN showed highest expression with most significant reduction (Fig. 25B-C).
Taken together, these data suggest that absence of Sectm1a impairs LXRα signal in macrophages.
73 Figure 23. RNA-seq analysis using WT and Sectm1a-KO BMDMs. (A-B) Heatmap (A) and volcano plot (B) of the overall gene expression alteration in BMDMs isolated from WT and
Sectm1a KO mice (n=3 per genotype). (C) Heatmap comparison of genes involved in cytokine- cytokine receptor interaction and chemokine signaling pathway.
74 Figure 24. Sectm1a deficiency downregulates LXRα-targeted genes. (A) Heatmap showing the top 20 most significantly downregulated genes in Sectm1a KO BMDMs. (B-C) Volcano plot
(B) and heat-map (C) of all LXR-related genes that were differentially expressed in Sectm1a KO
BMDMs. (D) expression of some common LXR-target genes in Sect1ma KO BMDMs were validated using qRT-PCR. n=3 for each genotype (*, p<0.05; data are presented as Mean ± SEM;
Student’s t test)
75 Figure 25. Deletion of Sectm1a affects LXRα, but not LXRβ pathway. (A) Expression of
LXRα and LXRβ as determined by RNA-seq analyses; (B) Venn diagram showing overlapped genes involved in sepsis, cardiovascular disease, and LXR-related signaling from our RNA-seq analyses. (C) Three genes overlapped among the 3 pathways mentioned in the Venn diagram (C)
(n=3 of each genotype). (*, p<0.05, data are presented as Mean ± SEM, Student’s t test).
76 Section 6. LXR Agonist Fails to Rescue LPS-Induced Inflammation and Cardiac
Dysfunction in Sectm1a-KO Model
To investigate the role of Sectm1a on LXRα signaling in BMDMs upon LPS challenge, we performed qRT-PCR to determine the gene expression levels of LXRα and its downstream targets. After treating BMDMs with LPS for 3 h, expression levels of LXRα and ABCG1 were significantly reduced in WT-LPS group, when compared with WT-PBS group (Fig. 26A).
Importantly, when comparing with WT-BMDMs after LPS treatment, the mRNA levels of
LXRα, ABCA1, ABCG1, and ApoE were further decreased in KO-LPS group (Fig. 26A). These data suggest that reduced baseline LXRα signaling cascade in macrophages of Sectm1a-KO mice may contribute to the aggravated inflammatory response following LPS administration. To gain insights on whether Sectm1a deficiency affects LXR signaling we next treated WT- and KO-
BMDMs with GW3965, a potent LXR agonist, 12 h prior to LPS stimulation, (Fig. 26B).
Treatment of GW3965 appeared to more markedly increase the nuclear translocation of LXRα in
WT than in Sectm1a-KO BMDMs when compared to their corresponding DMSO control groups
(Fig. 26C). Furthermore, the GW3965 treatment significantly attenuated the release of proinflammatory factors (i.e. TNFα, IL-6, IL-1β, and MCP-1) in WT BMDMs as early as 6 h after LPS treatment (Fig.27A-D). Nonetheless, this attenuation was not as strong in the cell culture supernatants of KO-GW3965 macrophages when compared to the WT-GW3965 group
(Fig. 27A-D).
Previous studies have reported the cardiac protective effects of GW3965 in db/db diabetic and ischemic/reperfused mouse models, which is mainly mediated through LXRα instead of
LXRβ subtype (161, 162). Consistent with these findings, LPS-induced cardiac dysfunction was improved with GW3965 treatment in WT-LPS group, as evidenced by a 29% increase in
77 fractional shortening (FS %) when compared to WT-DMSO group (Fig. 28A-B and Table 9). In contrast, the percentage of fractional shortening did not differ statistically between KO-DMSO and KO-GW3965 groups (Fig. 28A-B and Table 9).
To determine whether Sectm1a modulates LXRα signaling via direct physical interaction, we co-transfected HEK293T cells with 2 plasmids encoding GFP-conjugated Sectm1a and HA tag-conjugated LXRα proteins for 48 h. Cell samples were harvested and subjected to co- immunoprecipitation assay. Reciprocal immunoprecipitation suggests a potential physical interaction between Sectm1a and LXRɑ (Fig. 29A-B), rendering the possibility that Sectm1a may be a previous unidentified ligand/co-activator of LXRα. Thus, the anti-inflammatory benefits of Sectm1a, including the maintenance of cardiac function following LPS treatment, may be contributed by activating LXRα pathway in macrophages.
78 Figure 26. Lack of Sectm1a impairs LXRα translocation to nucleus after stimulation with agonist. (A) Gene expression of LXRα and target genes in WT and Sectm1a-KO BMDMs after
3 h of LPS (10ng/ml) treatment was measured using qRT-PCR (n=3). (B) Graphic scheme of treatment protocol. BMDMs from WT and Sectm1a-KO mice were first treated with LXR agonist, GW3965 (2µM, 12 h) followed by LPS stimulation (10 mg/ml, up to 48 h). (C)
Immunofluorescent staining of BMDMs with LXRα antibody after 12 h of GW3965 stimulation and 30 min of LPS treatment. DNA was stained with DAPI (blue). (Scale bar, 10µm; *, p<0.05; data are presented as Mean ± SEM; 2-way ANOVA with Sidak’s multiple comparisons test)
79 Figure 27. LXR agonist fails to rescue LPS-induced inflammation in Sectm1a-KO BMDMs.
(A-D) After treating BMDMs with GW3965 for 12 h, cytokine levels in cell culture supernatant were determined by ELISA at indicated time points post LPS treatment (n=4-5). (*, p<0.05 when comparing WT-DMSO to WT-GW groups; #, p<0.05 when comparing WT-GW to KO-GW groups; data are presented as Mean ± SEM; 2-way ANOVA with Sidak’s multiple comparisons test)
80 Figure 28. Administration of LXR agonist shows no effect on cardiac function in Sectm1a-
KO mice upon LPS injection. (A-B) WT and Sectm1a KO mice received 3 injection of
GW3965 (30 mg/kg of BW, once daily, DMSO used as control), 6 hr after last GW3965 injection, all mice received LPS injection (10 mg/kg) and underwent echocardiography measurement to assess cardiac function (n=4-7). (*, p<0.05; data are presented as Mean ± SEM;
2-way ANOVA with Holm-Sidak’s multiple comparisons test)
81 Table 9. Echocardiographic measurements of WT and Sectm1a-KO mice with GW3965 and LPS injection.
82 Figure 29. Sectm1a interacts with LXRα. HEK293T cells were simultaneously co-transfected with plasmids encoding GFP-conjugated Sectm1a and HA tag-conjugated LXRα for 48 h. (A)
Co-immunoprecipitation results showed that HA tag can be detected from cell lysate samples purified by GFP antibody; or vice versa (B).
83 Section 7. Sectm1a-KO Provokes HFD-Induced Inflammation and Cardiac Dysfunction
Given the pivotal roles of Sectm1a in regulating acute inflammatory response in mice with endotoxemia, we were interested in exploring whether Sectm1a was also involved in chronic inflammatory conditions (i.e., obesity). After treating WT-BMDMs with palmitate (to mimic hyperlipidemia condition in vitro) for 24 h, we observed a significant reduction (43%) in
Sectm1a gene expression only at 0.5 mM dose (Fig. 30A). Of interest, by using the same dose,
Sectm1a level was downregulated in RAW264.7 macrophages as early as 3 h after palmitate treatment (Fig. 17B). To determine whether Sectm1a may affect macrophage activation upon lipid stimulation, BMDMs isolated from WT and Sectm1a-KO mice were treated with 0.5 mM palmitate for 24 h. The results showed significantly higher levels of TNFα and IL-6 in KO- palmitate group, when compared to WT-palmitate control (Fig. 30C).
Consistent with aforementioned acute inflammation model, cardiac function was impaired in Sectm1a-KO mice, compared to WT controls, after 20-wk HFD feeding (Fig. 31A-B, and Table 10). . Consistently with impaired cardiac function, hearts of KO-HFD mice exhibited increased infiltration of monocytes (Ly6Chigh) and macrophages (F4/80+) with higher expression of CCR2, an inflammatory marker (Fig. 32A-C, E-G). Furthermore, cardiac macrophages isolated from KO-HFD mice exhibited reduced levels of anti-inflammatory marker, CD301,
(Fig.32D/H). Put together, these data indicate that Sectm1a expression provides protection against the deleterious effects of chronic HFD feeding on inflammatory responses associated with cardiac dysfunction.
84 Figure 30. Lack of Sectm1a promotes palmitate-induced macrophage activation. (A) WT
BMDMs were treated with indicated doses of palmitate for 24 h, and (B) RAW264.7 macrophages were treated with 0.5 mM palmitate for indicated time points, then gene expression of Sectm1a was measured with qRT-PCR (n=3). (C) WT and Sectm1a-KO BMDMs were treated with palmitate (0.5 mM, 24 h), and cytokine levels in cell culture supernatant were measured using ELISA (n=7-8). (*, p<0.05; data are presented as Mean ± SEM; Student’s t test and 1-way
ANOVA)
85 Figure 31. Sectm1a-KO mice show impaired cardiac function upon HFD feeding. (A-B)
Cardiac function was determined by echocardiography after WT and Sectm1a KO mice were fed with HFD for 20 wk (n=10 per group). FS, fractional shortening; LVID;s, left ventricular internal diameter at systole; LVID;d, left ventricular internal diameter at diastole (*, p<0.05; data are presented as Mean ± SEM; Student’s t test)
86 Table 10. Echocardiographic measurements of WT and Sectm1a- KO mice with 20 weeks of high fat diet feeding
87 Figure 32. Absence of Sectm1a leads to increased accumulation of proinflammatory macrophages in hearts of obese mice. (A-H) Representative Flow Cytometry plots and quantification of cardiac macrophage marker expression showed more monocytes (Ly6C+) and macrophage accumulation (F4/80+) with inflammatory phenotype (CCR2+, CD301-) in the heart of KO mice 5 wk after HFD feeding (n=6). (*, p<0.05; data are presented as Mean ± SEM;
Student’s t test)
88 Section 8. Sectm1a Deficiency-Mediated Cardiac Dysfunction is Mainly Ascribed to
Augmented Inflammation in the Heart
To further dissect whether expression on heart tissue or in cells of hematopoietic origin infiltrating the myocardium mediates the beneficial effect of Sectm1a on cardiac function following acute inflammation, we performed a bone marrow cell transplantation experiment.
This experiment involves the reciprocal transplantation of bone marrow cells from intact WT and
KO mice to recipient WT mice that were previously irradiated to eliminate their hematopoietic cell precursors (Fig. 33A). Of note, there was no difference on body weight before irradiation and 4 weeks after bone marrow cell injection (Fig. 33B), indicating equivalent recovery between the mice injected with WT and Sectm1a-KO cells. However, recipient mice injected with
Sectm1a-deficient cells displayed exacerbated cardiac dysfunction upon LPS challenge, evidenced by 31% decrease in fractional shortening (Fig. 34A-B, Table 11), which was consistent with our previous results showing worsened LPS-induced cardiac dysfunction in
Sectm1a-KO mic. To determine cardiac inflammation, we harvested heart samples immediately after echocardiography and performed Flow Cytometry to characterize different immune cell populations. The vast majority of lymphocytes in the heart of recipient mice were originated from transplanted bone marrow cells (CD45.2+), whereas the endogenous immune cells
(CD45.1+) were almost completely diminished (Fig. 35A). Surprisingly, the number of activated neutrophils (CD11b+Ly6G+) were similar in both recipient groups (Fig. 35). Considering that our previous data showed larger endogenous population of neutrophils in hearts of LPS-treated
Sectm1a-KO mice (Fig. 13A-B), this result suggests that deficiency of Sectm1a in other cell types in the heart may affect the recruitment of neutrophils.
89 Nonetheless, we also observed higher numbers of Ly6C- macrophages in recipient mice with WT-cell injection, whereas mice injected with Sectm1a-KO cells showed more inflammatory macrophages in the heart (Ly6Chigh) (Fig. 36A-B), with increased CCR2 expression (Fig. 37A-B). Together, these data indicate that the adverse effects of Sectm1a deficiency on cardiac dysfunction are mainly ascribed to increased immune response elicited by increased infiltration and activation of monocytes/macrophages in the heart.
90 Figure 33. Transplantation of WT and Sectm1a-KO bone marrow cells. (A) Schematic illustration of bone marrow cell transplantation experiment. (B) Body weight measurements before and 4-week after transplantation were recorded (n= 7). (Data are presented as Mean ±
SEM; Student’s t test)
91 Figure 34. Transplantation of Sectm1a-deficient bone marrow cells aggravated cardiac dysfunction after LPS injection. Freshly isolated bone marrow cells from WT and Sectm1a-
KO mice were injected into recipient mice within 24 h after irradiation. After 4 weeks of recovery, recipient mice were administered with LPS, and echocardiography was performed 12 h later to determine cardiac function. (A) Representative echocardiography images and (B) calculation of fraction shortening (n=7). (*, p<0.05; data are presented as Mean ± SEM;
Student’s t test)
92 Table 11. Echocardiographic measurements of LPS-treated recipient mice
after transplantation of bone marrow cells from WT (WT-LPS)
and Sectm1a-KO (KO-LPS) mice
93 Figure 35. Transplantation of bone marrow cells from Sectm1a-KO mice has no effect on cardiac neutrophil infiltration upon LPS stimulation. (A) CD45.1+ cells were almost entirely diminished after irradiation whereas CD45.2+ cells accumulated in the heart after transplantation;
(B) the number of Ly6G+ neutrophils in the heart was similar between mice injected with WT cells and Sectm1a-KO cells (n=5). (Data are presented as Mean ± SEM; Student’s t test)
94 A WT-LPS KO-LPS F 4 / 8 0
Ly6C Ly6C
B * 40 * 40 Ф Ф
35 M 30 M h i g 30 h 20
25 10 F F 4 4 / 8 / 8 0 20 0 0 + + L WT-LPS KO-LPS WT-LPS KO-LPS L y y 6 6 C C -
Figure 36. Transplantation of bone marrow cells from Sectm1a-KO mice increased monocyte-derived macrophage population in the heart upon LPS stimulation. (A-B) the number of F4/80+Ly6C- macrophages were higher in recipients with WT-cells injection; whereas accumulation of F4/80+Ly6Chigh macrophages was significantly increased in mice injected with
KO-cells (n=5). (*, p<0.05; data are presented as Mean ± SEM; Student’s t test)
95 Figure 37. Sectm1a-deficienct bone marrow cells give rise to inflammatory macrophages with increased CCR2 expression after LPS treatment. (A-B) recipient mice injected with
KO-cells showed significantly higher CCR2 signal in cardiac macrophages (n=5). (*, p<0.05; data are presented as Mean ± SEM; Student’s t test)
96 Chapter IV: Discussion
Section. 1 Dissertation Summary
In the present study, we have identified Sectm1a as a previously unrecognized regulator of inflammatory responses, specifically macrophage activation, both during endotoxemia (LPS- induced acute inflammation) and upon HFD feeding (similar to chronic low-grade inflammation observed in obese population). Consistent with previous evidence (87, 96), we observed the highest gene expression of sectm1a in spleen, an organ that harbors abundant immune cells from myeloid lineage. To determine whether Sectm1a is involved in macrophage function, we first measured the gene expression level of Sectm1a in BMDMs stimulated with different doses of
LPS. qRT-PCR results showed downregulation of Sectm1a in BMDM upon low dose of LPS treatment (10 ng/ml). This dose provided near maximal inhibition of Sectm1a expression and higher doses did not further suppress its expression. However, to our surprise, gene expression of
Sectm1a was dynamically changed over time in BMDMs treated with low dose of LPS. Sectm1a expression increased acutely following LPS treatment, reaching a maximum after 6 h.
Afterwards, the expression declined and reached levels lower than those of the PBS control group after 24 h. Similar results were notice in vivo when determining the expression levels of
Sectm1a in spleen at different time points post LPS administration. These results suggest that early Sectm1a upregulation may play a role as a compensatory mechanism to cope with augmented inflammatory stress. This early increase may be critical to prevent an onset of exaggerated inflammatory response. This postulate is consistent with a greater propensity for enhanced inflammatory responses and cardiac dysfunction exhibited by mice with global knockout of Sectm1a after LPS injection when compared to WT control groups, including a higher mortality rate at a 72-h survival experiment. In addition, we observed that BMDMs
97 isolated from Sectm1a-KO mice were skewed toward proinflammatory phenotype after LPS treatment with increased secretion of inflammatory cytokines. In line with the in vitro finding,
Flow Cytometry analysis revealed markedly increased infiltration of neutrophils, monocytes and proinflammatory macrophages in the heart of LPS-treated Sectm1a-KO mice, when compared to
WT-LPS hearts. In addition, deficiency of Sectm1a increased cytokine levels in the heart, which are key mediators of cardiac dysfunction.
In agreement with our loss-of-function experiments, overexpression of Sectm1a in
BMDMs using adenovirus (Ad.Sectm1a) resulted in significantly lowered production of inflammatory cytokines after LPS treatment. These findings further support the notion that the acute surge in Sectm1a expression is critical to prevent detrimental inflammatory responses.
Therefore, upregulation of Sectm1a at early time points of LPS treatment may serve as inhibitory mechanism to reduce inflammation. Because LPS injection can significantly increase IFNγ gene expression in spleen, with a peak expression at 6 h followed by gradual decrease over time (170), and IFNγ has been shown to regulate Sectm1 gene expression in monocytes (88). This is consistent with our observation that Sectm1a gene expression also peaks at 6 h and reduces at later time points. Interestingly, we did not detect such anti-inflammatory effects from Sectm1a overexpression in cardiomyocytes. This leads us to hypothesize that the protection from inflammation associated to Sectm1a requires its expression in macrophages rather than in cardiomyocytes. The results from our bone marrow transplantation experiments further validated that hypothesis: the effects of Sectm1a in cardiac function is mainly mediated through modulation of immune response by cells derived from the hematopoietic compartment.
Using high-throughput RNA-sequencing, we have identified that the LXRα signaling pathway was disrupted in macrophages due to lack of Sectm1a. Consistent with these findings,
98 loss of Sectm1a expression resulted in lesser suppression of cytokine release from macrophages and failed to rescue of cardiac function following treatment with the LXR agonist GW3965. To assess whether Sectm1a may act as LXRα ligand, we co-transfected plasmids of Sectm1a and
LXRα into HEK293T cells; and results from co-immunoprecipitation experiments suggested a potential interaction between Sectm1a and LXRα, thus activating the downstream signaling pathway. Nonetheless, more in depth experiments are required to demonstrate the binding of
Sectm1a to LXRɑ. Collectively, these findings uncover the critical role of Sectm1a in the regulation of endotoxin- and obesity-associated inflammation and cardiac dysfunction.
Section 2. The role of Sectm1a in regulating inflammatory response of macrophage
In this study, we found that Sectm1a was greatly downregulated in BMDMs at the later phase of LPS (24-48 h post-LPS) or palmitate treatments compared to vehicle-treated controls, yet the underlying mechanisms regulating Sectm1a expression remains unclear under both conditions. Our findings are consistent with previous studies showing that human Sectm1 is an early response gene to IFNγ in MM6 human monocytes, where its expression is increased at early time points (3, 6, 12 h) but decreases after 24 h; and the induction of Sectm1 expression by
IFNγ can be suppressed by LPS (89). Here we showed that Sectm1a expression was increased by more than 900-fold in WT BMDMs 4 h after IFNγ (10 ng/ml) treatment, but when BMDMs were treated with IFNγ and LPS, expression of Sectm1a was only increased by 20-fold. Interestingly,
TNFα mRNA levels were 10-fold higher in IFNγ+LPS group when comparing to IFNγ alone group, indicating that LPS-induced reduction of Sectm1a expression was able to trigger stronger inflammatory response. Indeed, Tsalik et al recently sequenced peripheral blood RNA of 129 representative subjects with systemic inflammatory response syndrome (SIRS) or sepsis,
99 including sepsis survivors and sepsis non-survivors, and revealed that Sectm1 was significantly higher in sepsis survivors, compared to non-survivor SIRS controls (90).
On the other hand, very little is known about Sectm1b so far. Previously, Sectm1b has been shown to act as an inhibitor of T cell activation, counteracting the action of Sectm1a (96).
However, given that Sectm1b shares less than 40% protein homology with either Sectm1a or human Sectm1 (96); the expression levels of Sectm1b in BMDMs showed no difference after
LPS treatment; and ablation of Sectm1a does not affect Sectm1b expression, we speculate that
Sectm1b might not participate in the regulation of macrophage activation. Collectively, these findings suggest that Sectm1a may act as an anti-inflammatory mediator, and its expression could be promptly reduced as inflammatory response progresses.
Following an insult by either intrinsic or extrinsic stimuli, the interplay of cardiomyocytes, cardiac fibroblasts, and innate immune cells determine an effective recovery or insufficient repair of damaged tissue. Macrophages, as the most abundant leukocytes in the heart, comprise
~7% of total nonmyocytes with great heterogeneity in origin (67). In particular, abundance and phenotype of macrophages are altered during inflammatory and reparative processes (67, 158).
Recent studies (78-81) reveal that, similar to mouse hearts, human cardiac macrophages can also be partitioned into distinct subsets depending on the expression of CCR2, a critical factor required for monocyte migration. As demonstrated by various approaches including genetic fate mapping, single-cell transcriptomics and parabiosis, cardiac CCR2- macrophages are a self- maintained resident population established early in development; whereas CCR2+ macrophages are derived from recruited monocytes and replenished through proliferation (81, 163). More importantly, CCR2+ macrophages are critical players to coordinate cardiac inflammation with marked increase of IL-1β; and the change of absolute number or percentage of CCR2+
100 macrophages is positively correlated with left ventricular systolic dysfunction following mechanic unloading (67). Consistently, our findings showed substantial increased of CCR2+ macrophages in the hearts of Sectm1a KO mice with LPS injection or HFD feeding, resulting in exacerbated inflammation and cardiac suppression.
Accordingly, Sectm1a-deficient macrophages released profoundly higher levels of cytokines (TNFα, IL-6, and IL1β), which are well-known inflammatory mediators that are ascribed to the pathogenesis of heart failure (164, 165). In particular, IL-6 was the most abundant cytokine in the myocardium of WT mouse after LPS injection, yet the concentration of IL-6 protein was 2-fold higher in Sectm1a-KO mouse hearts, when compared to WT-LPS group, which suggested that IL-6 may be the major contributor to cardiac dysfunction in Sectm1a-KO mice under inflammatory condition. In addition, Chomarat et al. previously reported that, upon interacting with monocytes, fibroblasts increased IL-6 secretion, which in turn upregulated the expression of macrophage colony-stimulating factor (M-CSF) receptor on monocytes. This resulted in differentiation of monocyte into macrophages instead of antigen presenting dendritic cells (166). Thus, in our Sectm1a KO mouse model, LPS treatment enhances IL-6 concentration in myocardium, which could contribute to further aggravation of the inflammatory response in heart by promoting monocyte differentiation into macrophages. Furthermore, besides affecting monocyte/macrophage recruitment and activation, IL-6 has been shown to be responsible for the induction of ICAM1 on cardiomyocytes during reperfusion, which would subsequently lead to increased infiltration of neutrophils, resulting in secondary damage associated with neutrophil activation (167-169). Interestingly, according to our RNA-sequencing results, the gene expression level of 5-LOX was reduced by 1.9 fold in Sectm1a-KO BMDMs, when comparing to WT BMDMs. Since 5-LOX is an essential mediator for lipoxin production, our results suggest
101 that Sectm1a deficiency causes significant impairment in the capacity to resolve inflammation in macrophages.
Section 3. Effects of Sectm1a on LXR signaling pathway
Along this line, as instructed by the RNA-Sequencing analyses, we provide further insight into Sectm1a-mediated regulation of macrophage activation through LXR signaling pathway. We observed that Sectm1a deficiency had dramatically increased inflammatory responses and aggravated cardiac dysfunction, when stimulated with LPS, which cannot be rescued by treatment with LXR agonist GW3965. Mechanistically, LXRα has been demonstrated to regulate NF-κB pathway, and thereby controls the downstream inflammatory responses in macrophages (124, 125). Consistently, when LXRα gene expression and activation was suppressed in WT BMDMs treated with LPS, we observed increased phosphorylation of p65 and
IkBα, indicating activated NF-κB pathway. Importantly, when comparing to WT-LPS group,
Sectm1a deficiency further suppressed LXRα signaling cascade, resulting in augmented phosphorylation of p65 and IkBα. More importantly, activation of NF-κB pathway could be alleviated by overexpressing Sectm1a in BMDMs and led to lower production of inflammatory cytokines. These data collectively suggest that Sectm1a could suppress inflammatory response by inhibiting NF-κB signaling through activation of LXRα pathway. Furthermore, subsequent gene network analysis identified that three genes (IL-1RN, Cav1, S100a8) were involved in sepsis, cardiovascular diseases and LXR signaling cascade, and IL-1RN showed highest expression with most significant reduction (Fig. 25). Future investigation on the effects of IL-
1RN might be beneficial to clarify the mechanism of Sectm1a.
102 Given that Sectm1a has a soluble form that is implicated to act as autocrine or paracrine, and previous study has predicted the IgG-like domain in the N-terminus of Secmt1a, we hypothesized that Sectm1a may bind to LXRα as a critical co-activator. Using plasmids encoding a GFP-conjugated Sectm1a protein and a HA tag-conjugated LXRα protein, we demonstrated that Sectm1a could bind to LXRα in vitro. Intriguingly, the expression level of Eid1 was significantly increased in BMDMs in the absence of Sectm1a. Eid1 has been known to inhibit the histone acetyltransferase activity of EP300, which is a key binding co-activator of LXR (104,
108, 109). Therefore, Sectm1a binding to LXRα may also activate a feed forward pathway to suppress the inhibitory mechanism, resulting in enhanced activation of LXRα signaling cascade.
Although RNA-seq results showed no change on LXRβ gene expression in Sectm1a-KO
BMDMs, possible interaction between Sectm1a and LXRβ may be worth of investigation.
Section 4. Limitations and future directions
Our in vitro experiments using adenovirus and in vivo bone marrow cell transplantation have demonstrated that the effects of Sectm1a on inflammation-associated cardiac dysfunction are primarily mediated through modulation of immune responses likely by macrophages.
However, since we have used a global knockout mouse model we are unable to completely ruled out potential contributions of Sectm1a deficiency on the heart itself, specifically on cardiomyocytes, fibroblasts, and endothelial cells to cardiac dysfunction. Strictly speaking this is plausible since all those cell types play critical roles in the pathophysiology of heart failure.
Furthermore, since LXRs is also expressed in these cell types, albeit in a much lower level comparing to that in macrophages (132), Sectm1a acting as LXRα co-activator may also manifest beneficial effects on maintaining or enhancing the function of these cells.
103 Secondly, here we have showed profound effects of Sectm1a on macrophage activation and the subsequent effects on cardiac function. However, we cannot completely rule out whether
Sectm1a regulates the recruitment of other immune cells, which in turn may contribute to the development of cardiac dysfunction. Increased infiltration of neutrophil was detected, this may due to increased CXCL2 expression in macrophages that causes a secondary influx of neutrophils. More in depth studies investigating the mechanisms whereby Sectm1a affects neutrophil function is warranted. Moreover, Sectm1a was originally known as co-stimulator for the proliferation and activation of T cells (96), which also have significant impact on cardiac function. Therefore, we cannot eliminate the possible interaction among Sectm1a and T cells, as well as the communication of T cells with macrophages and cardiomyocytes.
Lastly, one significant disadvantage of LXR agonism that limits its clinical potential is the adverse effects on lipogenesis (143). Although this falls outside the scope of this study, as a newly identified co-activator of LXRα, further investigation into the roles of Sectm1a in lipogenic pathways of LXRα is critically needed to determine the feasibility of Sectm1a to be used as a therapeutic target.
Section 5. Conclusion of the dissertation
In conclusion, our study presented here for the first time demonstrates that Sectm1a can protect against cardiac dysfunction triggered by acute or chronic inflammation. Specifically,
Sectm1a plays critical roles in modulating macrophage infiltration and activation in the heart, thus preventing excessive inflammatory responses. Mechanistically, the anti-inflammatory benefits of Sectm1a may be mediated through LXRɑ signaling pathway. Our data also shown the potential physical interaction between Sectm1a and LXRɑ, suggesting that Sectm1a may be a
104 previously unrecognized co-activator of LXRɑ. This leads to upregulation of genes that can modulate lipid homeostasis (eg, ABCA1, ABCG1) and inhibit inflammation (eg. NF-κB).
Therefore, approaches that enhance Sectm1a expression/activity would possess great therapeutic potential to treat inflammatory disease and its associated cardiac injury.
Figure 38. Graphic schema depicting the functional roles of Sectm1a on macrophage activation and subsequently cardiac injury.
105 References
1. James MA. Chapter II.2.2 - Inflammation, Wound Healing, and the Foreign-Body
Response. Biomaterials Science (Third Edition), Academic Press. 2013, Pages 503-512
2. Pahwa R, Goyal A, Bansal P, et al. Chronic Inflammation. [Updated 2020 Feb 14]. In:
StatPearls [Internet]. Treasure Island (FL): StatPearls Publishing; 2020 Jan-. Available
from: https://www.ncbi.nlm.nih.gov/books/NBK493173/
3. Lämmermann T, Afonso PV, Angermann BR, Wang JM, Kastenmüller W, Parent CA,
Germain RN. Neutrophil swarms require LTB4 and integrins at sites of cell death in
vivo. Nature. 2013; 498:371-5
4. Ridker PM, Everett BM, Pradhan A, MacFadyen JG, Solomon DH, Zaharris E, Mam V,
Hasan A, Rosenberg Y, Iturriaga E, Gupta M, Tsigoulis M, Verma S, Clearfield M,
Libby P, Goldhaber SZ, Seagle R, Ofori C, Saklayen M, Butman S, Singh N, Le May
M, Bertrand O, Johnston J, Paynter NP, Glynn RJ; CIRT Investigators. Low-Dose
Methotrexate for the Prevention of Atherosclerotic Events. N Engl J Med. 2019;
380:752-762
5. Ridker PM, Everett BM, Thuren T, MacFadyen JG, Chang WH, Ballantyne C, Fonseca
F, Nicolau J, Koenig W, Anker SD, Kastelein JJP, Cornel JH, Pais P, Pella D, Genest J,
Cifkova R, Lorenzatti A, Forster T, Kobalava Z, Vida-Simiti L, Flather M, Shimokawa
H, Ogawa H, Dellborg M, Rossi PRF, Troquay RPT, Libby P, Glynn RJ; CANTOS
Trial Group. Antiinflammatory Therapy with Canakinumab for Atherosclerotic Disease.
N Engl J Med. 2017; 377:1119-1131
106 6. InformedHealth.org [Internet]. Cologne, Germany: Institute for Quality and Efficiency
in Health Care (IQWiG); 2006-. What is an inflammation? 2010 Nov 23 [Updated 2018
Feb 22]. Available from: https://www.ncbi.nlm.nih.gov/books/NBK279298/
7. Morioka S, Maueröder C, Ravichandran KS. Living on the Edge: Efferocytosis at the
Interface of Homeostasis and Pathology. Immunity. 2019; 50:1149-1162
8. Kolaczkowska E, Kubes P. Neutrophil recruitment and function in health and
inflammation. Nat Rev Immunol. 2013; 13:159-75
9. Stortz JA, Raymond SL, Mira JC, Moldawer LL, Mohr AM, Efron PA. Murine Models
of Sepsis and Trauma: Can We Bridge the Gap? ILAR J. 2017; 58:90-105
10. Seemann S, Zohles F, Lupp A. Comprehensive comparison of three different animal
models for systemic inflammation. J Biomed Sci. 2017; 24:60
11. Wassenaar TM, Zimmermann K. Lipopolysaccharides in Food, Food Supplements, and
Probiotics: Should we be worried? Eur J Microbiol Immunol (Bp). 2018; 3:63–69
12. an Lier D, Geven C, Leijte GP, Pickkers P. Experimental human endotoxemia as a
model of systemic inflammation. Biochimie. 2019; 159:99-106
13. Recknagel P, Gonnert FA, Halilbasic E, Gajda M, Jbeily N, Lupp A, Rubio I, Claus RA,
Kortgen A, Trauner M, Singer M, Bauer M. Mechanisms and functional consequences
of liver failure substantially differ between endotoxaemia and faecal peritonitis in rats.
Liver Int. 2013; 33:283-93
14. Wang L, Li Y, Wang X, Wang P, Essandoh K, Cui S, Huang W, Mu X, Liu Z, Wang Y,
Peng T, Fan GC. GDF3 Protects Mice against Sepsis-Induced Cardiac Dysfunction and
Mortality by Suppression of Macrophage Pro-Inflammatory Phenotype. Cells. 2020; 9.
pii: E120
107 15. Jia L, Wang Y, Wang Y, Ma Y, Shen J, Fu Z, Wu Y, Su S, Zhang Y, Cai Z, Wang J,
Xiang M. Heme Oxygenase-1 in Macrophages Drives Septic Cardiac Dysfunction via
Suppressing Lysosomal Degradation of Inducible Nitric Oxide Synthase. Circ Res.
2018; 122:1532-1544
16. McInnes IB, Schett G. Pathogenetic insights from the treatment of rheumatoid arthritis.
Lancet. 2017; 389:2328-2337
17. Adamo L, Rocha-Resende C, Prabhu SD, Mann DL. Reappraising the role of
inflammation in heart failure. Nat Rev Cardiol. 2020
18. Avtanski D, Pavlov VA, Tracey KJ, Poretsky L. Characterization of inflammation and
insulin resistance in high-fat diet-induced male C57BL/6J mouse model of obesity.
Animal Model Exp Med. 2019; 2:252-258
19. Sansbury BE, Spite M. Resolution of Acute Inflammation and the Role of Resolvins in
Immunity, Thrombosis, and Vascular Biology. Circ Res. 2016; 119:113-30
20. Silvestre-Roig C, Braster Q, Ortega-Gomez A, Soehnlein O. Neutrophils as regulators
of cardiovascular inflammation. Nat Rev Cardiol. 2020
21. Headland SE, Norling LV. The resolution of inflammation: Principles and challenges.
Semin Immunol. 2015; 27:149-60
22. Wallace JL, Ianaro A, Flannigan KL, Cirino G. Gaseous mediators in resolution of
inflammation. Semin Immunol. 2015; 27:227-33
23. Schett G, Neurath MF. Resolution of chronic inflammatory disease: universal and
tissue-specific concepts. Nat Commun. 2018; 9:3261
24. Serhan CN. Pro-resolving lipid mediators are leads for resolution physiology. Nature.
2014; 510:92-101
108 25. Dalli J, Ramon S, Norris PC, Colas RA, Serhan CN. Novel proresolving and tissue-
regenerative resolvin and protectin sulfido-conjugated pathways. FASEB J. 2015;
29:2120-36
26. Basil MC, Levy BD. Specialized pro-resolving mediators: endogenous regulators of
infection and inflammation. Nat Rev Immunol. 2016; 16:51-67
27. Poon IK, Lucas CD, Rossi AG, Ravichandran KS. Apoptotic cell clearance: basic
biology and therapeutic potential. Nat Rev Immunol. 2014; 14:166-80
28. Serhan CN, Brain SD, Buckley CD, Gilroy DW, Haslett C, O'Neill LA, Perretti M,
Rossi AG, Wallace JL. Resolution of inflammation: state of the art, definitions and
terms. FASEB J. 2007; 21:325-32
29. Elliott MR, Koster KM, Murphy PS. Efferocytosis Signaling in the Regulation of
Macrophage Inflammatory Responses. J Immunol. 2017; 198:1387-1394
30. Serhan, C. N. & Sheppard, K. A. Lipoxin formation during human neutrophil-platelet
interactions. Evidence for the transformation of leukotriene A4 by platelet
12- lipoxygenase in vitro. J. Clin. Invest. 1990; 85:772–780
31. Levy BD, Romano M, Chapman HA, Reilly JJ, Drazen J, Serhan CN. Human alveolar
macrophages have 15-lipoxygenase and generate 15(S)-hydroxy-5,8,11-cis-13- trans-
eicosatetraenoic acid and lipoxins. J Clin Invest. 1993; 92:1572-9
32. Serhan, C. N., Hamberg, M. & Samuelsson, B. Lipoxins: novel series of biologically
active compounds formed from arachidonic acid in human leukocytes. Proc. Natl Acad.
Sci. USA. 1984; 81:5335–5339
33. Levine B, Kalman J, Mayer L, Fillit HM, Packer M. Elevated circulating levels of
tumor necrosis factor in severe chronic heart failure. N Engl J Med. 1990; 323:236-41
109 34. Vagnozzi RJ, Maillet M, Sargent MA, Khalil H, Johansen AKZ, Schwanekamp JA,
York AJ, Huang V, Nahrendorf M, Sadayappan S, Molkentin JD. An acute immune
response underlies the benefit of cardiac stem cell therapy. Nature. 2020; 577:405-409
35. Ellison GM, Vicinanza C, Smith AJ, Aquila I, Leone A, Waring CD, Henning BJ,
Stirparo GG, Papait R, Scarfò M, Agosti V, Viglietto G, Condorelli G, Indolfi C,
Ottolenghi S, Torella D, Nadal-Ginard B. Adult c-kit(pos) cardiac stem cells are
necessary and sufficient for functional cardiac regeneration and repair. Cell. 2013;
154:827-42
36. Liu Q, Yang R, Huang X, Zhang H, He L, Zhang L, Tian X, Nie Y, Hu S, Yan Y,
Zhang L, Qiao Z, Wang QD, Lui KO, Zhou B. Genetic lineage tracing identifies in situ
Kit-expressing cardiomyocytes. Cell Res. 2016; 26:119-30
37. van Berlo JH, Kanisicak O, Maillet M, Vagnozzi RJ, Karch J, Lin SC, Middleton RC,
Marbán E, Molkentin JD. c-kit+ cells minimally contribute cardiomyocytes to the heart.
Nature. 2014; 509:337-41
38. verett BM, Cornel JH, Lainscak M, Anker SD, Abbate A, Thuren T, Libby P, Glynn RJ,
Ridker PM. Anti-Inflammatory Therapy With Canakinumab for the Prevention of
Hospitalization for Heart Failure. Circulation. 2019; 139:1289-1299
39. Dick SA, Epelman S. Chronic Heart Failure and Inflammation: What Do We Really
Know? Circ Res. 2016; 119:159-76
40. Mann DL. The emerging role of innate immunity in the heart and vascular system: for
whom the cell tolls. Circ Res. 2011; 108:1133-45
41. Mackey D, McFall AJ. MAMPs and MIMPs: proposed classifications for inducers of
innate immunity. Mol Microbiol. 2006; 61:1365-71
110 42. Mann DL. Innate immunity and the failing heart: the cytokine hypothesis revisited.
Circ Res. 2015; 116:1254-68
43. Santosh k, Subir M. Role of cytokines in heart failure. J Pharmacol Rep. 2017. 2:123
44. Yokoyama T, Vaca L, Rossen RD, Durante W, Hazarika P, Mann DL. Cellular basis
for the negative inotropic effects of tumor necrosis factor-alpha in the adult mammalian
heart. J Clin Invest. 1993; 92:2303-12
45. Yan AT, Yan RT, Cushman M, Redheuil A, Tracy RP, Arnett DK, Rosen BD,
McClelland RL, Bluemke DA, Lima JA. Relationship of interleukin-6 with regional
and global left-ventricular function in asymptomatic individuals without clinical
cardiovascular disease: insights from the Multi-Ethnic Study of Atherosclerosis. Eur
Heart J. 2010; 31:875-82
46. Duncan DJ, Hopkins PM, Harrison SM. Negative inotropic effects of tumour necrosis
factor-alpha and interleukin-1beta are ameliorated by alfentanil in rat ventricular
myocytes. Br J Pharmacol. 2007; 150:720-6
47. Long CS. The role of interleukin-1 in the failing heart. Heart Fail Rev. 2001; 6:81-94
48. Tsai CT, Wu CK, Lee JK, Chang SN, Kuo YM, Wang YC, Lai LP, Chiang FT, Hwang
JJ, Lin JL. TNF-α down-regulates sarcoplasmic reticulum Ca²⁺ ATPase expression and
leads to left ventricular diastolic dysfunction through binding of NF-κB to promoter
response element. Cardiovasc Res. 2015; 105:318-29
49. Fauconnier J, Meli AC, Thireau J, Roberge S, Shan J, Sassi Y, Reiken SR, Rauzier JM,
Marchand A, Chauvier D, Cassan C, Crozier C, Bideaux P, Lompré AM, Jacotot E,
Marks AR, Lacampagne A. Ryanodine receptor leak mediated by caspase-8 activation
111 leads to left ventricular injury after myocardial ischemia-reperfusion. Proc Natl Acad
Sci U S A. 2011; 108:13258-63
50. Krown KA, Page MT, Nguyen C, Zechner D, Gutierrez V, Comstock KL, Glembotski
CC, Quintana PJ, Sabbadini RA. Tumor necrosis factor alpha-induced apoptosis in
cardiac myocytes. Involvement of the sphingolipid signaling cascade in cardiac cell
death. J Clin Invest. 1996; 98:2854-65
51. Schulz R, Panas DL, Catena R, Moncada S, Olley PM, Lopaschuk GD. The role of
nitric oxide in cardiac depression induced by interleukin-1 beta and tumour necrosis
factor-alpha. Br J Pharmacol. 1995; 114:27-34
52. Shen Y, Qin J, Bu P. Pathways involved in interleukin-1β-mediated murine
cardiomyocyte apoptosis. Tex Heart Inst J. 2015; 42:109-16
53. Ing DJ, Zang J, Dzau VJ, Webster KA, Bishopric NH. Modulation of cytokine-induced
cardiac myocyte apoptosis by nitric oxide, Bak, and Bcl-x. Circ Res. 1999; 84:21-33
54. Rose-John S. IL-6 trans-signaling via the soluble IL-6 receptor: importance for the pro-
inflammatory activities of IL-6. Int J Biol Sci. 2012; 8:1237-47
55. Markousis-Mavrogenis G, Tromp J, Ouwerkerk W, Devalaraja M, Anker SD, Cleland
JG, Dickstein K, Filippatos GS, van der Harst P, Lang CC, Metra M, Ng LL,
Ponikowski P, Samani NJ, Zannad F, Zwinderman AH, Hillege HL, van Veldhuisen DJ,
Kakkar R, Voors AA, van der Meer P. The clinical significance of interleukin-6 in heart
failure: results from the BIOSTAT-CHF study. Eur J Heart Fail. 2019; 21:965-973
56. Fontes JA, Rose NR, Čiháková D. The varying faces of IL-6: From cardiac protection
to cardiac failure. Cytokine. 2015; 74:62-8
112 57. Yu X, Kennedy RH, Liu SJ. JAK2/STAT3, not ERK1/2, mediates interleukin-6-
induced activation of inducible nitric-oxide synthase and decrease in contractility of
adult ventricular myocytes. J Biol Chem. 2003; 278:16304-9
58. Prabhu SD. Cytokine-induced modulation of cardiac function. Circ Res. 2004;
95:1140-53
59. Yamauchi-Takihara K, Kishimoto T. Cytokines and their receptors in cardiovascular
diseases--role of gp130 signalling pathway in cardiac myocyte growth and maintenance.
Int J Exp Pathol. 2000; 81:1-16
60. Hirota H, Yoshida K, Kishimoto T, Taga T. Continuous activation of gp130, a signal-
transducing receptor component for interleukin 6-related cytokines, causes myocardial
hypertrophy in mice. Proc Natl Acad Sci U S A. 1995; 92:4862-6
61. Girbl T, Lenn T, Perez L, Rolas L, Barkaway A, Thiriot A, Del Fresno C, Lynam E,
Hub E, Thelen M, Graham G, Alon R, Sancho D, von Andrian UH, Voisin MB, Rot A,
Nourshargh S. Distinct Compartmentalization of the Chemokines CXCL1 and CXCL2
and the Atypical Receptor ACKR1 Determine Discrete Stages of Neutrophil Diapedesis.
Immunity. 2018; 49:1062-1076.e6
62. Wang L, Zhang YL, Lin QY, Liu Y, Guan XM, Ma XL, Cao HJ, Liu Y, Bai J, Xia YL,
Du J, Li HH. CXCL1-CXCR2 axis mediates angiotensin II-induced cardiac
hypertrophy and remodelling through regulation of monocyte infiltration. Eur Heart J.
2018; 39:1818-1831
63. Aukrust P, Ueland T, Müller F, Andreassen AK, Nordøy I, Aas H, Kjekshus J,
Simonsen S, Frøland SS, Gullestad L. Elevated circulating levels of C-C chemokines in
patients with congestive heart failure. Circulation. 1998; 97:1136-43
113 64. França CN, Izar MCO, Hortêncio MNS, do Amaral JB, Ferreira CES, Tuleta ID,
Fonseca FAH. Monocyte subtypes and the CCR2 chemokine receptor in cardiovascular
disease. Clin Sci (Lond). 2017; 131:1215-1224
65. Prabhu SD, Frangogiannis NG. The Biological Basis for Cardiac Repair After
Myocardial Infarction: From Inflammation to Fibrosis. Circ Res. 2016; 119:91-112
66. Swirski FK, Nahrendorf M. Cardioimmunology: the immune system in cardiac
homeostasis and disease. Nat Rev Immunol. 2018; 18:733-744
67. Pinto AR, Ilinykh A, Ivey MJ, Kuwabara JT, D'Antoni ML, Debuque R, Chandran A,
Wang L, Arora K, Rosenthal NA, Tallquist MD. Revisiting Cardiac Cellular
Composition. Circ Res. 2016; 118:400-9
68. Zouggari Y, Ait-Oufella H, Bonnin P, Simon T, Sage AP, Guérin C, Vilar J, Caligiuri
G, Tsiantoulas D, Laurans L, Dumeau E, Kotti S, Bruneval P, Charo IF, Binder CJ,
Danchin N, Tedgui A, Tedder TF, Silvestre JS, Mallat Z. B lymphocytes trigger
monocyte mobilization and impair heart function after acute myocardial infarction. Nat
Med. 2013; 19:1273-80
69. Adamo L, Staloch LJ, Rocha-Resende C, Matkovich SJ, Jiang W, Bajpai G,
Weinheimer CJ, Kovacs A, Schilling JD, Barger PM, Bhattacharya D, Mann DL.
Modulation of subsets of cardiac B lymphocytes improves cardiac function after acute
injury. JCI Insight. 2018; 3:120137
70. Horckmans M, Bianchini M, Santovito D, Megens RTA, Springael JY, Negri I, Vacca
M, Di Eusanio M, Moschetta A, Weber C, Duchene J, Steffens S. Pericardial Adipose
Tissue Regulates Granulopoiesis, Fibrosis, and Cardiac Function After Myocardial
Infarction. Circulation. 2018; 137:948-960
114 71. Kaya Z, Leib C, Katus HA. Autoantibodies in heart failure and cardiac dysfunction.
Circ Res. 2012; 110:145-58
72. Vazquez MI, Catalan-Dibene J, Zlotnik A. B cells responses and cytokine production
are regulated by their immune microenvironment. Cytokine. 2015; 74:318-26
73. Yndestad A, Holm AM, Müller F, Simonsen S, Frøland SS, Gullestad L, Aukrust P.
Enhanced expression of inflammatory cytokines and activation markers in T-cells from
patients with chronic heart failure. Cardiovasc Res. 2003; 60:141-6
74. Bansal SS, Ismahil MA, Goel M, Patel B, Hamid T, Rokosh G, Prabhu SD. Activated T
Lymphocytes are Essential Drivers of Pathological Remodeling in Ischemic Heart
Failure. Circ Heart Fail. 2017; 10:e003688
75. Bansal SS, Ismahil MA, Goel M, Zhou G, Rokosh G, Hamid T, Prabhu SD.
Dysfunctional and Proinflammatory Regulatory T-Lymphocytes Are Essential for
Adverse Cardiac Remodeling in Ischemic Cardiomyopathy. Circulation. 2019;
139:206-221
76. van Furth R, Cohn ZA. The origin and kinetics of mononuclear phagocytes. J Exp Med.
1968; 128:415-35
77. Epelman S, Liu PP, Mann DL. Role of innate and adaptive immune mechanisms in
cardiac injury and repair. Nat Rev Immunol. 2015; 15:117-29
78. Sieweke MH, Allen JE. Beyond stem cells: self-renewal of differentiated macrophages.
Science. 2013; 342:1242974
79. Epelman S, Lavine KJ, Beaudin AE, Sojka DK, Carrero JA, Calderon B, Brija T,
Gautier EL, Ivanov S, Satpathy AT, Schilling JD, Schwendener R, Sergin I, Razani B,
Forsberg EC, Yokoyama WM, Unanue ER, Colonna M, Randolph GJ, Mann DL.
115 Embryonic and adult-derived resident cardiac macrophages are maintained through
distinct mechanisms at steady state and during inflammation. Immunity. 2014; 40:91-
104
80. Lavine KJ, Epelman S, Uchida K, Weber KJ, Nichols CG, Schilling JD, Ornitz DM,
Randolph GJ, Mann DL. Distinct macrophage lineages contribute to disparate patterns
of cardiac recovery and remodeling in the neonatal and adult heart. Proc Natl Acad Sci
U S A. 2014; 111:16029-34
81. Bajpai G, Schneider C, Wong N, Bredemeyer A, Hulsmans M, Nahrendorf M,
Epelman S, Kreisel D, Liu Y, Itoh A, Shankar TS, Selzman CH, Drakos SG, Lavine KJ.
The human heart contains distinct macrophage subsets with divergent origins and
functions. Nat Med. 2018; 24:1234-1245
82. Bajpai G, Bredemeyer A, Li W, Zaitsev K, Koenig AL, Lokshina I, Mohan J, Ivey B,
Hsiao HM, Weinheimer C, Kovacs A, Epelman S, Artyomov M, Kreisel D, Lavine KJ.
Tissue Resident CCR2- and CCR2+ Cardiac Macrophages Differentially Orchestrate
Monocyte Recruitment and Fate Specification Following Myocardial Injury. Circ Res.
2019; 124:263-278
83. Sager HB, Hulsmans M, Lavine KJ, Moreira MB, Heidt T, Courties G, Sun Y,
Iwamoto Y, Tricot B, Khan OF, Dahlman JE, Borodovsky A, Fitzgerald K, Anderson
DG, Weissleder R, Libby P, Swirski FK, Nahrendorf M. Proliferation and Recruitment
Contribute to Myocardial Macrophage Expansion in Chronic Heart Failure. Circ Res.
2016; 119:853-64
116 84. Haynes, B. F., Denning, S. M., Singer, K. H., and Kurtzburg, J. Ontogeny of T cell
precursors: A model for the initial stages of human T cell development. Immunol.
Today. 1989. 10: 87–91
85. Slentz-Kesler KA, Hale LP, Kaufman RE. Identification and characterization of K12
(SECTM1), a novel human gene that encodes a Golgi-associated protein with
transmembrane and secreted isoforms. Genomics. 1998; 47:327-40
86. Bianchetti L, Tarabay Y, Lecompte O, Stote R, Poch O, Dejaegere A, Viville S. Tex19
and Sectm1 concordant molecular phylogenies support co-evolution of both eutherian-
specific genes. BMC Evol Biol. 2015. 15:222
87. Kamata H, Yamamoto K, Wasserman GA, Zabinski MC, Yuen CK, Lung WY, Gower
AC, Belkina AC, Ramirez MI, Deng JC, Quinton LJ, Jones MR, Mizgerd JP. Epithelial
Cell-Derived Secreted and Transmembrane 1a Signals to Activated Neutrophils during
Pneumococcal Pneumonia. Am J Respir Cell Mol Biol. 2016; 55:407-18
88. Lam GK, Liao HX, Xue Y, Alam SM, Scearce RM, Kaufman RE, Sempowski GD,
Haynes BF. Expression of the CD7 ligand K-12 in human thymic epithelial
cells:regulation by IFN-gamma. J Clin Immunol. 2005.25:41-9
89. Huyton T, Göttmann W, Bade-Döding C, Paine A, Blasczyk R. The T/NK cell co-
stimulatory molecule SECTM1 is an IFN "early response gene" that is negatively
regulated by LPS in human monocytic cells. Biochim Biophys Acta. 2011; 1810:1294-
301
90. Tsalik EL, Langley RJ, Dinwiddie DL, Miller NA, Yoo B, van Velkinburgh JC, Smith
LD, Thiffault I, Jaehne AK, Valente AM, Henao R, Yuan X, Glickman SW, Rice BJ,
McClain MT, Carin L, Corey GR, Ginsburg GS, Cairns CB, Otero RM, Fowler VG Jr,
117 Rivers EP, Woods CW, Kingsmore SF. An integrated transcriptome and expressed
variant analysis of sepsis survival and death. Genome Med. 2014; 6:111
91. Lyman SD, Escobar S, Rousseau AM, Armstrong A, Fanslow WC. Identification of
CD7 as a cognate of the human K12 (SECTM1) protein. J Biol Chem. 2000;275:3431-7
92. Sempowski GD, Lee DM, Kaufman RE, Haynes BF. Structure and function of the CD7
molecule. Crit Rev Immunol. 1999; 19:331-48
93. Wang T, Huang C, Lopez-Coral A, Slentz-Kesler KA, Xiao M, Wherry EJ, Kaufman
RE. K12/SECTM1, an interferon-γ regulated molecule, synergizes with CD28 to
costimulate human T cell proliferation. J Leukoc Biol. 2012; 91:449-59
94. Bade-Döding C, Göttmann W, Baigger A, Farren M, Lee KP, Blasczyk R, Huyton T.
Autocrine GM-CSF transcription in the leukemic progenitor cell line KG1a is mediated
by the transcription factor ETS1 and is negatively regulated through SECTM1 mediated
ligation of CD7. Biochim Biophys Acta. 2014; 1840:1004-13
95. Wang T, Ge Y, Xiao M, Lopez-Coral A, Li L, Roesch A, Huang C, Alexander P, Vogt
T, Xu X, Hwang WT, Lieu M, Belser E, Liu R, Somasundaram R, Herlyn M, Kaufman
RE. SECTM1 produced by tumor cells attracts human monocytes via CD7-mediated
activation of the PI3K pathway. J Invest Dermatol. 2014; 134:1108-1118
96. Howie D, Garcia Rueda H, Brown MH, Waldmann H. Secreted and transmembrane 1A
is a novel co-stimulatory ligand. PLoS One. 2013; 8:e73610
97. Huang P, Chandra V, Rastinejad F. Structural overview of the nuclear receptor
superfamily: insights into physiology and therapeutics. Annu Rev Physiol. 2010;
72:247-72
118 98. Wang K, Wan YJ. Nuclear receptors and inflammatory diseases. Exp Biol Med
(Maywood). 2008;233:496-506
99. Joseph SB, Castrillo A, Laffitte BA, Mangelsdorf DJ, Tontonoz P. Reciprocal
regulation of inflammation and lipid metabolism by liver X receptors. Nat Med. 2003; 9:
213-9
100. Apfel R, Benbrook D, Lernhardt E, Ortiz MA, Salbert G, Pfahl M. A novel orphan
receptor specific for a subset of thyroid hormone-responsive elements and its
interaction with the retinoid/thyroid hormone receptor subfamily. Mol Cell Biol.
1994;14:7025-35
101. Per-Arne S, Daniel A.H, Margareta J, Mikael C.O.E, Lillemor M. H, Bertil G.O,
Johannes H, Olov W, Bjorn C, Bjorn F, Lena M.S.C. Identification of genes
predominantly expressed in human macrophages. Atherosclerosis. 2004; 2:287-290
102. Botez G, Piraino G, Hake PW, Ledford JR, O'Connor M, Cook JA, Zingarelli B. Age-
dependent therapeutic effects of liver X receptor-α activation in murine polymicrobial
sepsis. Innate Immun. 2015; 21:609-18
103. Morello F, de Boer RA, Steffensen KR, Gnecchi M, Chisholm JW, Boomsma F,
Anderson LM, Lawn RM, Gustafsson JA, Lopez-Ilasaca M, Pratt RE, Dzau VJ. Liver
X receptors alpha and beta regulate renin expression in vivo. J Clin Invest. 2005;
115:1913-22
104. Wang B, Tontonoz P. Liver X receptors in lipid signalling and membrane homeostasis.
Nat Rev Endocrinol. 2018; 14:452-463
119 105. Willy PJ, Umesono K, Ong ES, Evans RM, Heyman RA, Mangelsdorf DJ. LXR, a
nuclear receptor that defines a distinct retinoid response pathway. Genes Dev. 1995;
9:1033-45
106. Yang C, McDonald JG, Patel A, Zhang Y, Umetani M, Xu F, Westover EJ, Covey DF,
Mangelsdorf DJ, Cohen JC, Hobbs HH. Sterol intermediates from cholesterol
biosynthetic pathway as liver X receptor ligands. J Biol Chem. 2006; 281:27816-26
107. Muse ED, Yu S, Edillor CR, Tao J, Spann NJ, Troutman TD, Seidman JS, Henke A,
Roland JT, Ozeki KA, Thompson BM, McDonald JG, Bahadorani J, Tsimikas S,
Grossman TR, Tremblay MS, Glass CK. Cell-specific discrimination of desmosterol
and desmosterol mimetics confers selective regulation of LXR and SREBP in
macrophages. Proc Natl Acad Sci U S A. 2018; 115:E4680-E4689
108. Huuskonen J, Fielding PE, Fielding CJ. Role of p160 coactivator complex in the
activation of liver X receptor. Arterioscler Thromb Vasc Biol. 2004; 24:703-8
109. Liang N, Jakobsson T, Fan R, Treuter E. The Nuclear Receptor-Co-repressor Complex
in Control of Liver Metabolism and Disease. Front Endocrinol (Lausanne). 2019;
10:411
110. Schulman IG. Liver X receptors link lipid metabolism and inflammation. FEBS Lett.
2017; 591:2978-2991
111. Chawla A, Boisvert WA, Lee CH, Laffitte BA, Barak Y, Joseph SB, Liao D, Nagy L,
Edwards PA, Curtiss LK, Evans RM, Tontonoz P. A PPAR gamma-LXR-ABCA1
pathway in macrophages is involved in cholesterol efflux and atherogenesis. Mol Cell.
2001; 7:161-71
120 112. Repa JJ, Turley SD, Lobaccaro JA, Medina J, Li L, Lustig K, Shan B, Heyman RA,
Dietschy JM, Mangelsdorf DJ. Regulation of absorption and ABC1-mediated efflux of
cholesterol by RXR heterodimers. Science. 2000; 289:1524-9
113. Repa JJ, Liang G, Ou J, Bashmakov Y, Lobaccaro JM, Shimomura I, Shan B, Brown
MS, Goldstein JL, Mangelsdorf DJ. Regulation of mouse sterol regulatory element-
binding protein-1c gene (SREBP-1c) by oxysterol receptors, LXRalpha and LXRbeta.
Genes Dev. 2000; 14:2819-30
114. Schultz JR, Tu H, Luk A, Repa JJ, Medina JC, Li L, Schwendner S, Wang S, Thoolen
M, Mangelsdorf DJ, Lustig KD, Shan B. Role of LXRs in control of lipogenesis. Genes
Dev. 2000; 14:2831-8
115. Leopold Wager CM, Arnett E, Schlesinger LS. Macrophage nuclear receptors:
Emerging key players in infectious diseases. PLoS Pathog. 2019; 15:e1007585
116. Joseph SB, Bradley MN, Castrillo A, Bruhn KW, Mak PA, Pei L, Hogenesch J,
O'connell RM, Cheng G, Saez E, Miller JF, Tontonoz P. LXR-dependent gene
expression is important for macrophage survival and the innate immune response. Cell.
2004; 119:299-309
117. Valledor AF, Hsu LC, Ogawa S, Sawka-Verhelle D, Karin M, Glass CK. Activation of
liver X receptors and retinoid X receptors prevents bacterial-induced macrophage
apoptosis. Proc Natl Acad Sci U S A. 2004; 101:17813-8
118. Matalonga J, Glaria E, Bresque M, Escande C, Carbó JM, Kiefer K, Vicente R, León
TE, Beceiro S, Pascual-García M, Serret J, Sanjurjo L, Morón-Ros S, Riera A, Paytubi
S, Juarez A, Sotillo F, Lindbom L, Caelles C, Sarrias MR, Sancho J, Castrillo A, Chini
EN, Valledor AF. The Nuclear Receptor LXR Limits Bacterial Infection of Host
121 Macrophages through a Mechanism that Impacts Cellular NAD Metabolism. Cell Rep.
2017; 18:1241-1255
119. Korf H, Vander Beken S, Romano M, Steffensen KR, Stijlemans B, Gustafsson JA,
Grooten J, Huygen K. Liver X receptors contribute to the protective immune response
against Mycobacterium tuberculosis in mice. J Clin Invest. 2009; 119:1626-37
120. Dubrovsky L, Van Duyne R, Senina S, Guendel I, Pushkarsky T, Sviridov D,
Kashanchi F, Bukrinsky M. Liver X receptor agonist inhibits HIV-1 replication and
prevents HIV-induced reduction of plasma HDL in humanized mouse model of HIV
infection. Biochem Biophys Res Commun. 2012; 419:95-8.
121. Ramezani A, Dubrovsky L, Pushkarsky T, Sviridov D, Karandish S, Raj DS, Fitzgerald
ML, Bukrinsky M. Stimulation of Liver X Receptor Has Potent Anti-HIV Effects in a
Humanized Mouse Model of HIV Infection. J Pharmacol Exp Ther. 2015; 354:376-83
122. Miao CM, He K, Li PZ, Liu ZJ, Zhu XW, Ou ZB, Ruan XZ, Gong JP, Liu CA. LXRα
represses LPS-induced inflammatory responses by competing with IRF3 for GRIP1 in
Kupffer cells. Int Immunopharmacol. 2016; 35:272-279
123. Pourcet B, Gage MC, León TE, Waddington KE, Pello OM, Steffensen KR, Castrillo A,
Valledor AF, Pineda-Torra I. The nuclear receptor LXR modulates interleukin-18
levels in macrophages through multiple mechanisms. Sci Rep. 2016; 6:25481
124. Wu S, Yin R, Ernest R, et al. Liver X receptors are negative regulators of cardiac
hypertrophy via suppressing NF-kappaB signalling. Cardiovasc Res. 2009; 84:19–126
125. Ito A, Hong C, Rong X, Zhu X, Tarling EJ, Hedde PN, Gratton E, Parks J, Tontonoz P.
LXRs link metabolism to inflammation through Abca1-dependent regulation of
membrane composition and TLR signaling. Elife. 2015; 4:e08009
122 126. Castrillo A, Joseph SB, Vaidya SA, Haberland M, Fogelman AM, Cheng G, Tontonoz
P. Crosstalk between LXR and toll-like receptor signaling mediates bacterial and viral
antagonism of cholesterol metabolism. Mol Cell. 2003; 12:805-16
127. Thomas DG, Doran AC, Fotakis P, Westerterp M, Antonson P, Jiang H, Jiang XC,
Gustafsson JÅ, Tabas I, Tall AR. LXR Suppresses Inflammatory Gene Expression and
Neutrophil Migration through cis-Repression and Cholesterol Efflux. Cell Rep. 2018;
25:3774-3785.e4
128. Souto FO, Castanheira FVS, Trevelin SC, Lima BHF, Cebinelli GCM, Turato WM,
Auxiliadora-Martins M, Basile-Filho A, Alves-Filho JC, Cunha f67FQ. Liver X
receptor activation impairs neutrophil functions and aggravates sepsis. J Infect Dis.
2019. pii: jiz635
129. Tavazoie MF, Pollack I, Tanqueco R, Ostendorf BN, Reis BS, Gonsalves FC, Kurth I,
Andreu-Agullo C, Derbyshire ML, Posada J, Takeda S, Tafreshian KN, Rowinsky E,
Szarek M, Waltzman RJ, Mcmillan EA, Zhao C, Mita M, Mita A, Chmielowski B,
Postow MA, Ribas A, Mucida D, Tavazoie SF. LXR/ApoE Activation Restricts Innate
Immune Suppression in Cancer. Cell. 2018; 172:825-840.e18
130. Cheng Y, Feng Y, Zhu M, Yan B, Fu S, Guo J, Hu J, Song X, Guo S, Liu G. Synthetic
liver X receptor agonist T0901317 attenuates high glucose-induced oxidative stress,
mitochondrial damage and apoptosis in cardiomyocytes. Acta Histochem. 2014;
116:214-21
131. Han D, Li X, Li S, Su T, Fan L, Fan WS, Qiao HY, Chen JW, Fan MM, Li XJ, Wang
YB, Ma S, Qiu Y, Tian ZH, Cao F. Reduced silent information regulator 1 signaling
123 exacerbates sepsis-induced myocardial injury and mitigates the protective effect of a
liver X receptor agonist. Free Radic Biol Med. 2017; 113:291-303
132. Cannon MV, van Gilst WH, de Boer RA. Emerging role of liver X receptors in cardiac
pathophysiology and heart failure. Basic Res Cardiol. 2016; 111:3
133. Moore KJ, Tabas I. Macrophages in the pathogenesis of atherosclerosis. Cell. 2011;
145:341-55
134. Repa JJ, Mangelsdorf DJ. The liver X receptor gene team: potential new players in
atherosclerosis. Nat Med. 2002; 8:1243-8
135. Joseph SB, McKilligin E, Pei L, Watson MA, Collins AR, Laffitte BA, Chen M, Noh G,
Goodman J, Hagger GN, Tran J, Tippin TK, Wang X, Lusis AJ, Hsueh WA, Law RE,
Collins JL, Willson TM, Tontonoz P. Synthetic LXR ligand inhibits the development of
atherosclerosis in mice. Proc Natl Acad Sci U S A. 2002; 99:7604-9
136. Bischoff ED, Daige CL, Petrowski M, Dedman H, Pattison J, Juliano J, Li AC,
Schulman IG. Non-redundant roles for LXRalpha and LXRbeta in atherosclerosis
susceptibility in low density lipoprotein receptor knockout mice. J Lipid Res. 2010;
51:900-6
137. Helleboid-Chapman A, Helleboid S, Jakel H, Timmerman C, Sergheraert C, Pattou F,
Fruchart-Najib J, Fruchart JC. Glucose regulates LXRalpha subcellular localization and
function in rat pancreatic beta-cells. Cell Res. 2006; 16:661-70
138. Ogihara T, Chuang JC, Vestermark GL, Garmey JC, Ketchum RJ, Huang X, Brayman
KL, Thorner MO, Repa JJ, Mirmira RG, Evans-Molina C. Liver X receptor agonists
augment human islet function through activation of anaplerotic pathways and
glycerolipid/free fatty acid cycling. J Biol Chem. 2010; 285:5392-404
124 139. Laffitte BA, Chao LC, Li J, Walczak R, Hummasti S, Joseph SB, Castrillo A, Wilpitz
DC, Mangelsdorf DJ, Collins JL, Saez E, Tontonoz P. Activation of liver X receptor
improves glucose tolerance through coordinate regulation of glucose metabolism in
liver and adipose tissue. Proc Natl Acad Sci U S A. 2003; 100:5419-24
140. Steffensen KR, Gustafsson JA. Putative metabolic effects of the liver X receptor (LXR).
Diabetes. 2004; 53 Suppl 1:S36-42
141. Korach-André M, Archer A, Barros RP, Parini P, Gustafsson JÅ. Both liver-X receptor
(LXR) isoforms control energy expenditure by regulating brown adipose tissue activity.
Proc Natl Acad Sci U S A. 2011; 108:403-8
142. Cao G, Liang Y, Broderick CL, Oldham BA, Beyer TP, Schmidt RJ, Zhang Y,
Stayrook KR, Suen C, Otto KA, Miller AR, Dai J, Foxworthy P, Gao H, Ryan TP,
Jiang XC, Burris TP, Eacho PI, Etgen GJ. Antidiabetic action of a liver x receptor
agonist mediated by inhibition of hepatic gluconeogenesis. J Biol Chem. 2003;
278:1131-6. Epub 2002 Oct 31
143. Kase ET, Wensaas AJ, Aas V, Højlund K, Levin K, Thoresen GH, Beck-Nielsen H,
Rustan AC, Gaster M. Skeletal muscle lipid accumulation in type 2 diabetes may
involve the liver X receptor pathway. Diabetes. 2005; 54:1108-15
144. Choe SS, Choi AH, Lee JW, Kim KH, Chung JJ, Park J, Lee KM, Park KG, Lee IK,
Kim JB. Chronic activation of liver X receptor induces beta-cell apoptosis through
hyperactivation of lipogenesis: liver X receptor-mediated lipotoxicity in pancreatic
beta-cells. Diabetes. 2007; 56:1534-43
125 145. Tamura K, Chen YE, Horiuchi M, Chen Q, Daviet L, Yang Z, Lopez-Ilasaca M, Mu H,
Pratt RE, Dzau VJ. LXRalpha functions as a cAMP-responsive transcriptional regulator
of gene expression. Proc Natl Acad Sci U S A. 2000; 97:8513-8
146. Kuipers I, van der Harst P, Kuipers F, van Genne L, Goris M, Lehtonen JY, van
Veldhuisen DJ, van Gilst WH, de Boer RA. Activation of liver X receptor-alpha
reduces activation of the renal and cardiac renin-angiotensin-aldosterone system. Lab
Invest. 2010; 90:630-6
147. Cannon MV, Silljé HH, Sijbesma JW, Khan MA, Steffensen KR, van Gilst WH, de
Boer RA. LXRα improves myocardial glucose tolerance and reduces cardiac
hypertrophy in a mouse model of obesity-induced type 2 diabetes. Diabetologia. 2016;
59:634-43
148. Wu S, Yin R, Ernest R, Li Y, Zhelyabovska O, Luo J, Yang Y, Yang Q. Liver X
receptors are negative regulators of cardiac hypertrophy via suppressing NF-kappaB
signalling. Cardiovasc Res. 2009; 84:119-26
149. Cannon MV, Yu H, Candido WM, Dokter MM, Lindstedt EL, Silljé HH, van Gilst WH,
de Boer RA. The liver X receptor agonist AZ876 protects against pathological cardiac
hypertrophy and fibrosis without lipogenic side effects. Eur J Heart Fail. 2015; 17:273-
82
150. Kuipers I, Li J, Vreeswijk-Baudoin I, Koster J, van der Harst P, Silljé HH, Kuipers F,
van Veldhuisen DJ, van Gilst WH, de Boer RA. Activation of liver X receptors with
T0901317 attenuates cardiac hypertrophy in vivo. Eur J Heart Fail. 2010; 12:1042-50
151. Trouplin V, Boucherit N, Gorvel L, Conti F, Mottola G, Ghigo E. Bone marrow-
derived macrophage production. J Vis Exp. 2013; 81:e50966
126 152. Qian J, Vafiadaki E, Florea SM, Singh VP, Song W, Lam CK, Wang Y, Yuan Q,
Pritchard TJ, Cai W, Haghighi K, Rodriguez P, Wang HS, Sanoudou D, Fan GC,
Kranias EG. Small heat shock protein 20 interacts with protein phofsphatase-1 and
enhances sarcoplasmic reticulum calcium cycling. Circ Res. 2011; 108:1429-1438
153. Qin D, Wang X, Li Y, Wang L, Wang R, Peng J, Essandoh K, Mu X, Peng T, Han Q,
Yu KJ, Fan GC. MicroRNA-223-5p and -3p Cooperatively Suppress Necroptosis in
Ischemic/Reperfused Hearts. J Biol Chem. 2016; 291:20247–20259
154. Koch SE, Gao X, Haar L, Jiang M, Lasko VM, Robbins N, Cai W, Brokamp C, Varma
P, Tranter M, Liu Y, Ren X, Lorenz JN, Wang HS, Jones WK, Rubinstein J.
Probenecid: novel use as a non-injurious positive inotrope acting via cardiac TRPV2
stimulation. J Mol Cell Cardiol. 2012; 53:134-44
155. Yu YR, O'Koren EG, Hotten DF, Kan MJ, Kopin D, Nelson ER, Que L, Gunn MD. A
Protocol for the Comprehensive Flow Cytometric Analysis of Immune Cells in Normal
and Inflamed Murine Non-Lymphoid Tissues. PLoS One. 2016;11:e0150606
156. Peng J, Li Y, Wang X, Deng S, Holland J, Yates E, Chen J, Gu H, Essandoh K, Mu X,
Wang B, McNamara RK, Peng T, Jegga AG, Liu T, Nakamura T, Huang K, Perez-
Tilve D, Fan GC. An Hsp20-FBXO4 Axis Regulates Adipocyte Function through
Modulating PPARγ Ubiquitination. Cell Rep. 2018; 23:3607-3620
157. Liu YC, Yu MM, Shou ST, Chai YF. Sepsis-induced cardiomyopathy: mechanisms and
treatments. Front Immunol. 2017; 8:1021
158. Ma Y, Mouton AJ, Lindsey ML. Cardiac macrophage biology in the steady-state heart,
the aging heart, and following myocardial infarction. Transl Res. 2018; 191:15-28
127 159. Christian F, Smith EL, Carmody RJ. The Regulation of NF-κB Subunits by
Phosphorylation. Cells. 2016; 5:e12
160. Wang Y, Liu J, Kong Q, Cheng H, Tu F, Yu P, Liu Y, Zhang X, Li C, Li Y, Min X, Du
S, Ding Z, Liu L. Cardiomyocyte-specific deficiency of HSPB1 worsens cardiac
dysfunction by activating NFκB-mediated leucocyte recruitment after myocardial
infarction. Cardiovasc Res. 2019; 115:154–167
161. He Q, Pu J, Yuan A, Yao T, Ying X, Zhao Y, Xu L, Tong H, He B. Liver X receptor
agonist treatment attenuates cardiac dysfunction in type 2 diabetic db/db mice.
Cardiovasc Diabetol. 2014; 13:149
162. He Q, Pu J, Yuan A, Lau WB, Gao E, Koch WJ, Ma XL, He B. Activation of liver-X-
receptor α but not liver-X-receptor β protects against myocardial ischemia/reperfusion
injury. Circ Heart Fail. 2014; 7:1032-1041
163. Andreadou I, Cabrera-Fuentes HA, Devaux Y, Frangogiannis NG, Frantz S, Guzik T,
Liehn EA, Gomes CPC, Schulz R, Hausenloy DJ. Immune cells as targets for
cardioprotection: new players and novel therapeutic opportunities. Cardiovasc Res.
2019; 115:1117-1130
164. Zhang Y, Huang Z, Li H. Insights into innate immune signalling in controlling cardiac
remodelling. Cardiovasc Res. 2017; 113:1538-1550
165. Zuurbier CJ, Abbate A, Cabrera-Fuentes HA, Cohen MV, Collino M, De Kleijn DPV,
Downey JM, Pagliaro P, Preissner KT, Takahashi M, Davidson SM. Innate immunity
as a target for acute cardioprotection. Cardiovasc Res. 2019; 115:1131-1142
166. Chomarat P, Banchereau J, Davoust J, Palucka AK. IL-6 switches the differentiation of
monocytes from dendritic cells to macrophages. Nat Immunol. 2000; 1:510-4
128 167. Kukielka GL, Smith CW, Manning AM, Youker KA, Michael LH, Entman ML.
Induction of interleukin-6 synthesis in the myocardium. Potential role in
postreperfusion inflammatory injury. Circulation. 1995; 92:1866-75
168. Smith CW, Entman ML, Lane CL, Beaudet AL, Ty TI, Youker K, Hawkins HK,
Anderson DC. Adherence of neutrophils to canine cardiac myocytes in vitro is
dependent on intercellular adhesion molecule-1. J Clin Invest. 1991; 88:1216-23
169. Kukielka GL, Hawkins HK, Michael L, Manning AM, Youker K, Lane C, Entman ML,
Smith CW, Anderson DC. Regulation of intercellular adhesion molecule-1 (ICAM-1)
in ischemic and reperfused canine myocardium. J Clin Invest. 1993; 92:1504-16
170. Varma TK, Lin CY, Toliver-Kinsky TE, Sherwood ER. Endotoxin-induced gamma
interferon production: contributing cell types and key regulatory factors. Clin Diagn
Lab Immunol. 2002; 9:530-43.
129