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Understanding Macrophage Complexity in Metabolic Dysfunction-associated Steatotic Liver Disease: Transitioning from the M1/M2 Paradigm to Spatial Dynamics

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10 April 2024

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12 April 2024

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Abstract
Metabolic dysfunction-associated steatotic liver disease (MASLD) encompasses metabolic dysfunction-associated fatty liver (MASL) and metabolic dysfunction-associated steatohepatitis (MASH), with MASH posing a risk of progression to cirrhosis and hepatocellular carcinoma (HCC). The global prevalence of MASLD is estimated at approximately a quarter of the population, with significant healthcare costs and implications for liver transplantation. The pathogenesis of MASLD involves intrahepatic liver cells, extrahepatic components, and immunological aspects, particularly the involvement of macrophages. Hepatic macrophages are a crucial cellular component of the liver and play vital roles in liver function, contributing significantly to tissue homeostasis and swift responses during pathophysiological conditions. Recent advancements in technology have revealed the remarkable heterogeneity and plasticity of hepatic macrophage populations and their activation states in MASLD, challenging traditional classification methods like the M1/M2 paradigm and highlighting the coexistence of harmful and beneficial macrophage phenotypes that are dynamically regulated during MASLD progression. This complexity underscores the importance of considering macrophage heterogeneity in therapeutic targeting strategies, including their distinct ontogeny and functional phenotypes. This review provides an overview of macrophage involvement in MASLD progression, combining traditional paradigms with recent insights from single-cell analysis and spatial dynamics. It also addresses unresolved questions and potential therapeutic targets in this area.
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Subject: Medicine and Pharmacology  -   Gastroenterology and Hepatology

1. Introduction

Accumulation of excess fat in the liver exceeding 5% can result in steatotic liver disease (SLD). Among various etiologies of steatosis, individuals with minimal or no alcohol consumption were previously diagnosed with a condition known as nonalcoholic fatty liver disease (NAFLD). NAFLD is now referred to as metabolic dysfunction-associated steatotic liver disease (MASLD), encompassing patients with hepatic steatosis and at least one of five cardiometabolic risk factors [1]. MASLD includes two histological subtypes: metabolic dysfunction-associated fatty liver (MASL), a relatively mild form characterized by fat accumulation or steatosis in the liver, and metabolic dysfunction-associated steatohepatitis (MASH), a progressive form accompanied by steatosis, inflammation, hepatocyte death, and fibrosis. MASH has the potential to progress to cirrhosis and ultimately to hepatocellular carcinoma (HCC) [2,3]. Approximately a quarter of the global population is now thought to have MASLD, with the estimated global prevalence of MASH ranging from 3% to 5% [2,4,5]. Additionally, MASLD has emerged as a major cause of end-stage liver failure and is rapidly becoming the leading cause of liver transplantation in the United States [6]. Age, gender, ethnicity, and metabolic conditions like diabetes and obesity are recognized as major risk factors for MASL and MASH. In addition, genetic and environmental factors add to the complexity of MASLD [7,8,9]. The economic burden of MASLD is enormous, with the healthcare costs for MASLD patients significantly exceeding those for patients with similar comorbidities but without MASLD [10,11].
Researchers have extensively studied the pathogenesis and progression of MASLD, recognizing the involvement of both intrahepatic liver parenchymal and nonparenchymal cells and extrahepatic components in the disease development [12]. One conceptual framework used to explain the progression from MASL to MASH is the 'two-hit' hypothesis. This hypothesis proposes that dysregulated hepatic lipid accumulation constitutes the initial hit, while oxidative, metabolic, and cytokine stresses represent the second hit [13,14]. In addition, a 'three-hit' hypothesis suggests that in MASLD, oxidative stress diminishes hepatocyte proliferation, prompting alternative regeneration pathways involving hepatic progenitor cells, with fibrosis progression dependent on the efficiency of hepatocyte regeneration, thus implicating impaired progenitor cell proliferation and cell death as the 'third hit' in MASLD progression [15,16,17]. Recently, there has been significant attention on immunological aspects in MASLD progression, particularly the involvement of macrophages. Hepatic macrophages are key players in innate immunity and a crucial cellular component of the liver, with the ratio of hepatocytes to macrophages ranging from 5:1 to 2.5:1 [18]. They contribute significantly to liver homeostasis, injury, and repair, exhibiting diverse subpopulations that dynamically change in health and disease [19]. In MASLD, macrophages are pivotal in driving inflammation, fibrosis progression, and regression, thus influencing the disease's pathogenesis and progression [3,20,21].
Traditionally, macrophages have been classified based on the M1/M2 paradigm, but recent advancements in high-end technologies have rendered this classification outdated, particularly in the context of deciphering the liver macrophage landscape. Over the past 5 years, there has been rapid expansion in understanding the clinical and research implications of liver macrophage diversity in MASLD. Additionally, the spatial localization of individual cells is increasingly recognized as a crucial parameter defining their function, necessitating integration into advanced multidimensional analyses. In this review, we summarize the role of macrophages in various stages of MASLD, encompassing the traditional M1/M2 paradigm alongside recent insights from single-cell and spatially resolved analyses. Furthermore, we offer perspectives on unresolved questions and potential therapeutic targets of macrophages in this field.

2. General Overview of Macrophages

Macrophages, a distinct type of cells with unique ability to clear up foreign bodies through a process known as phagocytosis, were discovered by a Russian-born zoologist and microbiologist Ilya (Elie) Metchnikoff nearly 140 years ago [22]. Metchnikoff’s groundbreaking discovery earned him the Nobel Prize in Physiology and Medicine in 1908. Macrophages are integral components of the mononuclear phagocytic system (MPS), a classification introduced by Furth et al. to encompass phagocytic mononuclear cells, which consist of immature monocytic cells along with their bone-marrow precursors, peripheral blood monocytes, and tissue-resident macrophages [23]. Lineage studies examining gene expression profiles and recruitment dynamics of various tissue macrophages have uncovered distinct developmental origins of these cells. This includes embryonic-derived and monocyte-derived macrophages, and these lineages remain independent of each other throughout adulthood [23,24,25,26,27,28]. As the effector cells of innate immune system, macrophages play a crucial role as the body’s primary line of defense. They not only identify and eliminate pathogens but also engage in communication with a specialized defense mechanism known as adaptive or acquired immune system [29]. Macrophages are ubiquitous across various tissues in the body, serving crucial functions throughout an organism’s life, from developmental stages to maintaining homeostasis and influencing the pathophysiology of diseases.
Macrophages are motile and their migrations towards the site of infection and inflammation are critical for their role as effector cells in innate immunity. They express a diverse array of surface receptors facilitating interaction with the host-derived and foreign ligands. These receptors enable macrophages to sense their environment and perform various functions. Typical examples include phosphatidylserine recognition receptors for apoptotic cells removal, complement receptors for clearing opsonized necrotic cells and altered-self molecules, and pattern recognition receptors (PRRs) such as toll-like receptors (TLRs), nod-like receptors (NLRs), sensors for intracellular DNA and RNA, c-type lectin, and scavenger receptors. This PRR family allows macrophages to recognize and bind directly to pathogens and their products, initiating processes like inflammasome activation [30,31].
Resident macrophages are crucial components of tissues, contributing to organ development and maintaining homeostasis. Their adapt to their environment, displaying specialized functions. Despite being fully differentiated; resident macrophages exhibit high plasticity and undergo phenotypic reprogramming in response to changing tissue microenvironment [25,32]. In a healthy state, resident macrophages balance response to foreign particles while minimizing tissue damage. They patrol tissues, phagocytose cellular debris, clean the surroundings, and facilitate tissue repair. Resident macrophages play diverse roles throughout an organism’s life span, including involvement in osteoclast and bone remodeling, erythropoiesis, brain development, and lung homeostasis during early developmental stages [33,34,35,36,37]. Their absence can lead to developmental abnormalities in organs like mammary glands, pancreas, and kidneys [38]. For instance, in mice lacking biologically functional colony stimulating factor 1 (CSF-1), the absence of osteoclasts, which are resident macrophages crucial for bone remodeling, leads to the development of osteoporosis—a condition characterized by weakened and brittle bones [39]. In specialized bone marrow areas called erythroblastic islands, macrophages support the production of red blood cells by maintaining cell interactions between erythroblasts and macrophages [40,41,42] and facilitate erythropoiesis by phagocytizing extruded erythroblast nuclei and supplying iron to erythroid progenitors [43,44]. Microglia, the tissue resident macrophages of the brain and spinal cord, play essential roles in central nervous system (CNS) development, homeostasis, and diseases [45,46]. Alveolar macrophages, found in lungs, help maintain lung function by clearing inhaled dust [47].

3. Hepatic Macrophages: Type, Origin, and Function

Hepatic macrophages consist of two major types: resident macrophages and infiltrated monocyte-derived macrophages (MDM) that rapidly emerge during injury. The resident hepatic macrophages were initially discovered by Karl Wilhelm von Kupffer, a Baltic German anatomist, and are now known as Kupffer cells (KCs) [48]. Comprising approximately 15% of the total liver cell population, KCs represent 80-90% of all tissue-resident macrophages in the body [49]. Within the liver lobules, 43% of Kupffer cells are distributed in the periportal area, 28% in the midzonal, and 29% in the centrilobular area [50]. KCs are located within the lumen of liver sinusoids with proximity to the sinusoidal endothelial cells that form the blood vessel walls. Despite being considered fixed resident cells, evidence shows that they can exhibit some degree of mobility along sinusoidal walls, either with or against the direction of blood flow [51].
Differential expressions of cell surface markers enable the distinction between KCs and MDMs. In mice, KCs are identified as CD11blow, F4/80high and Clec4F+, while MDMs are defined as CD11b+, F4/80intermediate, Ly6C+, and colony stimulating factor 1 receptor (CSF1R)+ [52,53,54,55]. Bone marrow progenitor cells, defined as CX3CR1+CD117+Lin−, differentiate into MDMs, which can further be subdivided into Ly6Clow or Ly6Chigh MDMs in mouse model of liver diseases [56]. Fate mapping experiment revealed that CSF1R+ erythro-myeloid progenitors (EMPs), originating in the yolk-sac at embryonic day E8.5 and subsequently colonizing the fetal liver at E10.5, give rise to KCs during embryonic development [57,58]. In one-year-old mice, most KCs are of embryonic origin, with some derived from hematopoietic stem cells [58,59]. The average half-life of mouse KCs is 12.4 days, while in rats, their lifespan extends from several weeks to months [60,61]. KCs are self-renewing and can proliferate into mature cells, making their replenishment in steady state independent of MDMs [27,62]. The advent of single-cell RNA-Seq has facilitated the characterization of human hepatic macrophages, classifying them into three distinct subgroups: CD68+MARCO+ KCs, CD68+MARCO− macrophages, and CD14+ monocytes [63,64]. The CD68+MARCO+ KCs contribute to immunotolerance, while the latter two exhibit a proinflammatory.
KCs act as the liver’s primary defense against pathogens and antigens from the gastrointestinal tract. To maintain tissue homeostasis, the liver fosters an anti-inflammatory microenvironment, promoting immunological tolerance to prevent unnecessary immune responses against food-derived antigens and bacterial products from the portal vein. Among liver cells, KCs play a central role in scavenging circulating antigens. Their immunological tolerance function involved several mechanisms. KCs present antigens to promote the arrest of CD4 T-cells and expand interleukin-10-producing regulatory T cells, fostering tolerogenic immunity [65]. In addition, KCs express lower levels of Major Histocompatibility Complex (MHC) class II and other costimulatory molecules like B7-1 and B7-2, resulting in weaker T cell activation compared to dendritic cells [66]. Moreover, KCs generate prostaglandins like PGE2 and 15d-PGJ2 that can suppress dendritic cell-mediated activation antigen-specific T cells [66].

4. Macrophage Accumulation in MASLD: Insights from Animal Models and Human Studies

In diet-induced disease models of MASLD, the hepatic macrophage numbers increase significantly with the feeding period [67]. The infiltrated proinflammatory CD45+/CD11b+/F4/80intermediate MDMs population in the liver of obese mice nearly doubled compared to lean control [68]. Importantly, diet not only increases the infiltration of MDMs but also disrupts the balance of pro and anti-inflammatory macrophages in the liver [69]. Therefore, characterizing the heterogeneity of hepatic macrophage subpopulations is crucial for advancing our understanding of MASLD.
Resident KCs are the predominant hepatic macrophages in the healthy liver, but their numbers have been reported to be reduced in MASH and hepatocellular carcinoma (HCC) [70,71]. As KCs number depletes, MDMs infiltrate the liver [72]. Chemokines, small heparin-binding proteins regulating cell trafficking, play a crucial role in this process. Monocyte chemoattractant protein-1 (MCP-1/CCL2), a member of the C-C chemokine family and a potent chemoattractant for monocytes, controls the migration and infiltration of MDMs. While various cell types produce CCL2, monocyte/macrophages are the major source [73,74]. CCL2 exerts its effects through its cognate receptor CCR2, whose expression is restricted to certain cell types, including monocytes [75]. Studies in both ob/ob and HFD-fed obese mice models have shown a positive association between hepatic expression of both Ccl2 and Ccr2 and body weight. In study by Morinaga et al. [76], both KCs and MDMs were fluorescently labeled and evaluated in a high-fat diet-fed MASH mice model. The MDMs population in obese mice was approximately six times higher in number and more proinflammatory compared to MDMs from lean mice. This study demonstrated that KCs played a vital role in recruiting MDMs, evident from the significantly elevated expression of CCL2, while MDMs displayed significantly higher expression of CCR2, leading to the conclusion that MDMs contribute to the severity of inflammation and hepatic insulin resistance in MASLD [76]. This group also demonstrated that blocking of the infiltration of MDMs using CCR2 antagonists ameliorated steatohepatitis and fibrosis [77].
Similarly, an increase in macrophage numbers has been observed in liver samples from MASH patients. A retrospective study analyzing liver biopsies from young MASLD patients revealed elevated numbers of CD68+ KCs, with higher levels correlating with MASLD severity [78]. Another study showed that the presence of enlarged KCs with significantly elevated phagocytic activity in the hepatic sinusoids closely associates with transformed hepatic stellate cells and hepatic oval cells during MASH, signifies their strong involvement in disease progression [79]. In addition, a significant increase in the number of CCR2+ MDMs in human liver samples from MASH patients is strongly correlated with the severity of MASH and fibrosis [76]. Furthermore, an increase in portal macrophage numbers has been observed in individuals with MASL [80]. As the disease progression towards MASH, portal inflammatory infiltrates accompany elevated expression of proinflammatory cytokines IL1B and tumor necrosis factor (TNF) [80]. The presence of increased CD11c-positive macrophages surrounding hepatocytes containing large lipid droplets, forming aggregates known as hepatic crown-like structures, correlates with hepatocytes death and the development of fibrosis in human patients with MASH [81,82]. These aggregates are considered important sources of inflammation and fibrosis because of their intactness and close association with activated fibroblasts and collagen deposition.

5. Stimuli Trigger Hepatic Macrophage Activation during MASLD Development

Multiple stimuli, such as fatty acids, cholesterol, damage associated molecular patterns (DAMPs), and pathogen associated molecular patterns (PAMPs), trigger macrophage activation in MASLD. The heightened influx of fatty acids into the liver, along with the de novo lipogenesis, exacerbates oxidative stress and lipid peroxidation in MASLD, thereby aggravating liver inflammation and fibrosis and accelerating MASH progression [83]. When co-culture with macrophages, steatotic hepatocytes release pro-inflammatory cytokines such as TNFα, MCP-1, interleukin 6 (IL-6), and IL-18, which can activate the macrophages [84]. Saturated fatty acids, such as lauric acid (C12:0) and palmitic acis (C16:0), induce the expression of inflammatory markers, including cyclooxygenase-2 (COX-2), inducible nitric oxide synthase (iNOS), and IL-1α in Raw 264.7 mouse macrophage cell line through the activation of toll like receptor (TLR) 4 and NF-κB pathway [85]. On the contrary, unsaturated fatty acids inhibit the NF-κB pathway, thus impeding the saturated fatty acids-mediated COX-2 expression in Raw 264.7 cells. In addition, palmitic acid activates TLR2 in THP-1 human monocytic cell line, inducing inflammasome-mediated-IL-1β production, suggesting that dietary fat can activate monocytes and promote peripheral inflammation in MASLD [86]. Furthermore, Kim et al. has demonstrated that infiltrating macrophages, rather than resident macrophages, generate reactive oxygen species (ROS) in response to palmitate through the dynamin-mediated TLR4- NADPH oxidase 2 (NOX2) axis. Their study also showed that mice lacking Nox2 are resistant to high-fat diet-induced hepatic steatosis and insulin resistance [87].
The disruption of hepatic cholesterol homeostasis and the accumulation of free cholesterol in hepatocytes are linked to the pathogenesis of MASH [88]. Loannou et al. described the presence of free cholesterol in the hepatocytes of MASH patients and diet-induced MASH mice model. They proposed that the aggregation and activation of KCs in crown-like structures around lipid droplets containing cholesterol crystals are significant indicators of the progression from simple steatosis to MASH [89]. In addition, KCs within the crown-like structures stained positively for NLRP3 and activated caspase 1, indicating a mechanistic link between KC activation exposure to cholesterol [90].
In addition, oxidative damage to cellular proteins, lipids, and DNA in hepatocytes generates oxidation-specific epitopes, acting as DAMPs, which interact with macrophage-expressed PRRs such as CD36 and TLR4, thereby initiating various immune responses [91]. For instance, in vitro studies demonstrated that mouse KCs can engulf apoptotic bodies from UV-treated mouse hepatocytes, triggering the production of Fas ligand and TNFα [92]. Additionally, extracellular vesicles (EV) released from hepatocytes, particularly in response to palmitic acid treatment, contain factors like tumor necrosis factor-related apoptosis-inducing ligand [93]. These EVs induce the expression of IL-1β and IL-6 in mouse bone marrow-derived macrophages.
Furthermore, PAMPs such as lipopolysaccharide (LPS) and microbial nucleotides act as danger signals, recognized by PRRs on macrophages, triggering inflammatory responses via intracellular signaling pathways [94]. In MASLD, there is an increased influx of gut-derived microbial products into the liver due to changes in gut microorganisms and increased intestinal permeability. This leads to elevated TLRs-mediated immune signaling, contributing to liver inflammation and fibrogenesis [95]. In addition, triglycerides enhance LPS-mediated expression of proinflammatory mediators such as inducible iNOS, TNFα, IL-1β, and IL-6 in rat KCs, compared to LPS stimulation alone [96]. Inhibition of the NF-κB pathway significantly reduces the potentiating effect of triglycerides on iNOS expression by KCs [96]. Electron microscopic analysis of KCs from high-fat diet-fed mice reveals intracellular lipid droplet accumulation [97]. These fat-laden KCs generate significantly high levels of proinflammatory cytokines and chemokines in response to LPS compared to KCs from chow diet-fed mice.

6. Classical M1/M2 Macrophage Paradigm and MASLD Development

The dynamic heterogeneity and reprogramming of macrophages contribute significantly to the pathogenesis and progression of various liver diseases. An important aspect of this macrophage adaptability is evident in the differentiation of macrophages into either classically activated M1, characterized by a pro-inflammatory profile, or alternatively activated M2 macrophages, displaying an anti-inflammatory and pro-fibrogenic phenotypes (73,77). This classification is rooted in their origins from the Th1 strains (C57BL/6, B10D2) or Th2 strains (BALB/c, DBA/2), respectively. Macrophages activated in response to IFN-γ differentiate into M-1 like macrophages capable of generating nitric oxide (NO) to kill parasites. One the contrary, Th2 cytokines, including IL-4 and IL-10, suppress the activation of M-1 like macrophages, and these M2-like macrophages exhibit elevated arginine metabolism [98,99]. Although the M1/M2 classification oversimplifies the intricate in vivo response, it is widely recognized that the differentiation of macrophages into distinct pro-inflammatory or anti-inflammatory phenotypes profoundly influences host defense and the pathogenesis of various liver diseases.

6.1. Mechanisms Control Macrophage Polarization

6.1.1. TLR and NF-κB

Bacterial endotoxin LPS activates TLR4 expressed on macrophages, triggering proinflammatory reactions crucial for eliminating invading bacteria. Beyond LPS, endogenous DAMPs, including high mobility group box protein 1 (HMGB1) and hyaluronic acid, are released during tissue injury, activating TLR4 to facilitate tissue repair [100,101,102]. TLR4 activation leads to NF-κB activation through the myeloid differentiation factor 88 (MyD88)-dependent pathways or interferon regulatory factor (IRF) 3, promoting the expression of proinflammatory factors [103]. Jing et al., using Raw264.7 cells and primary peritoneal macrophages, demonstrated that berberine, a competitive inhibitor of TLR4, disrupts the TLR4/MyD88/NFκB signaling pathway, interfering with LPS-mediated proinflammatory M1 like macrophage polarization [104]. In a separate study, Xiang et al. showed that olean-28,13β-olide 2 (NZ) inhibited LPS-mediated generation of proinflammatory cytokines in Raw264.7 cells through the suppression of TLR-NF-κB signaling, downregulation of NLRP3 expression, and inhibition of caspase-1 activation [105]. Similarly, Lu et al. demonstrated that quercetin, a natural flavonoid compound, inhibits LPS-mediated M1 macrophage polarization via the NF-κB and IRF5 signaling [106]. Together, these findings suggest that the macrophage-specific TLR4/NF-κB signaling axis plays an important role in M1 like proinflammatory macrophage polarization.

6.1.2. Signal Transducer and Activator of Transcription (STAT)

The STAT family, comprising seven structurally similar and highly conserved members, including STAT1, STAT2, STAT3, STAT4, STAT5A, STAT5B, and STAT6, are recognized as important regulators of macrophage polarization [107,108]. Interferons and TLR signaling polarizes macrophages towards M1 like proinflammatory phenotype via STAT1 signaling, whereas IL-4 and IL-13 tilt macrophages towards M2 like anti-inflammatory phenotype through STAT6 signaling pathways [109]. In a study with J774 murine macrophages, Haydar et al. demonstrated that azithromycin promotes M2-like macrophage polarization by inhibiting STAT1 and NF-κB signaling pathways. Another study by He et al. showed that IL-4 skews macrophage towards M2 subtype through the JAK1/STAT6 pathway [110]. In addition, STAT3 plays a determinative role in M2 polarization, as the suppression of JAK3/STAT3 by miR-221-3p promotes the shift of macrophage polarization from M2 to M1 subtype [111].

6.1.3. Transforming Growth Factor Beta (TGF-β) and SMAD

Upon TGF-β binding to the TGF-β receptor complex, the receptor complex activation triggers Smads (Smad2 or 3)-mediated or non-Smad signaling pathways to regulate gene expression [112]. Growth differentiation factor 3 (GDF3), a member of the TGF-β superfamily, can modulate the inflammatory response by influencing macrophage inflammatory phenotypes. GDF3 phosphorylates and activates Smad2/Smad3, inhibiting NLRP3 expression in macrophages and directing macrophage polarization toward the M2 phenotype [113]. Similarly, it has been shown that quercetin-mediated inhibition of the TGF-β1-smad2/3 pathway constrains M2 polarization [106].

6.1.4. Peroxisome Proliferator-Activated Receptor γ

PPARγ belongs to PPAR nuclear receptors family and is mainly expressed in adipose tissue, hematopoietic cells, and the large intestine. As a transcription factor, PPARγ plays a crucial role in glucose and lipid metabolism, anti-inflammatory responses, and the regulation of oxidative stress [114]. Studies have underscored the important association between PPARγ activation and macrophage polarization [115,116]. PPARγ regulates inflammatory processes by directly interacting with NF-κB, resulting in NF-κB degradation [117]. Activation of PPAR-γ by a PPARγ agonist in Raw264.7 macrophages shifts lipid-mediated macrophages polarization from the M1 to M2 phenotype through the interaction between PPAR-γ and NF-κB p65 [118]. Similarly, PPARγ activation promotes native human monocytes toward an anti-inflammatory M2 phenotype [119]. Using mice with macrophage-specific deletion of PPARγ, it was shown that PPARγ is essential for the maturation of alternatively activated macrophage [120]. These results suggest that PPAR-γ is a master regulator M2 polarization of macrophages.

6.1.5. MicroRNAs (miRNAs) and Other Mechanisms

MicroRNAs (miRNAs) have garnered significant interest due to their pivotal roles in macrophages polarization by regulating various signaling pathways [121]. For example, miR-221-3p and MiR-1246 facilitate alternative macrophage polarization, with mechanistical involvement in modulating JAK3/STAT3 and NF-κB signaling pathways [111,122]. Exosomal vesicles derived from adipocytes delivered miR-34a into macrophages, repressing kruppel-like factor 4 (Klf4) expression and consequently inhibiting M2 like macrophage polarization [123]. Beyond these mechanisms, additional signaling pathways, including notch signaling, mammalian target of rapamycin (mTOR) signaling, PI3K/Akt signaling, and JNK/c-Myc signaling pathways, have been identified to play roles in macrophage polarization [124,125,126,127].

6.2. Macrophage Polarization in Early Stage of MASLD

Macrophages with pro-inflammatory phenotypes exacerbate early MASLD severity, while those with anti-inflammatory characteristics contribute beneficially to MASLD initiation and type 2 diabetes. An initial study using a methionine-choline-deficient (MCD) diet in C57BL/6 and Balb/c mice has demonstrated that C57BL/6 mice with M1 bias displayed elevated liver steatosis and lobular inflammation compared to the Balb/c mice with M2 bias [128]. Further research corroborated that high-fat diet-fed BALB/c mice displayed increased M2 KC polarization, leading to the apoptosis of M1 KCs and mitigating liver steatosis and hepatocyte death compared to C57BL6/J mice, with in vitro experiments demonstrating that M2 macrophages induce M1 apoptosis via IL10-mediated arginase activation [129]. Additional studies found that high-fat diet enriched in polyunsaturated fatty acids promotes the alternative M2 macrophage activation and improves metabolic disturbances [129,130]. The activation of M2 KCs by PPARδ promotes and ameliorates obesity-induced insulin resistance [131]. Histidine-rich glycoprotein (HRGP) is an α2-plasma glycoprotein and mainly produced by liver parenchymal cells in mammals. Liver-derived HRGP has been shown to promote the polarization of M1 macrophages and inhibit M2 polarization in both tumor and inflammatory environments [132]. In MCD diet or carbon tetrachloride (CCl4)-induced chronic liver injury model, macrophages polarization was tipped towards M2 in mice lacking HRGP, attenuating liver injury and fibrosis [133].

6.3. Macrophage Polarization in Advanced Stage of MASLD

Liver biopsies from patients with MASH reveal an increase in proinflammatory myeloperoxidase-positive KCs along with elevated expression of the proinflammatory marker IL-6. Interestingly, the expression of anti-inflammatory macrophage markers such as IL-10, dectin-1, and epidermal growth factor-like module containing mucin-like hormone receptor 1 (EMR1) are also induced in MASH, suggesting a reparative role of M2 macrophages following tissue injury, which may contribute to fibrosis development [81,134]. Type 2 immunity is characterized by increased levels of cytokines such as IL-4, IL-5, IL-9, and IL-13. Blocking anti-inflammatory type 2 TGFβ and IL-13 signaling has been shown to protect against high-fat diet-induced liver fibrosis in mice [135,136]. The scavenger receptor CD163 is considered as a marker for anti-inflammatory macrophages. Interestingly, targeting CD163 in KCs and other M2 macrophages with an anti-CD163-IgG-dexamethasone conjugate has been shown to improve MASH pathologies, including hepatic inflammation, hepatocyte ballooning, fibrosis, and glycogen deposition in a rat model of fructose-induced MASH [137]. These findings underscore the important role of M2 macrophage activation in MASLD progression.

7. Revealing the Dynamic Landscape of Hepatic Macrophages in MASLD: Heterogeneity and Plasticity

The widespread use of single-cell RNA sequencing (scRNA-seq) has greatly enhanced our comprehension of cellular diversity and changes in macrophage subpopulations under specific healthy or diseased conditions, surpassing the traditional M1/M2 macrophage paradigm. In the normal mouse liver, KCs are identified using markers such as F4/80, CLEC4F, and T cell immunoglobulin and mucin domain-containing 4 (Timd4) [55]. During the early stages of MASLD, KCs engage in lipid storage, compromising their ability for self-renewal [138]. Consequently, embryonic KCs are gradually lost and replaced by MDMs lacking Timd4 expression [139,140]. Mulder K et al. integrated 41 mononuclear phagocytes scRNA-seq datasets to compile a comprehensive monocytes-macrophage-focused compendium, revealing a diverse array of specialized cell subsets distributed across multiple tissues [141]. They identified three conserved macrophage populations across tissues, namely TREM2, IL4I1, and HES1, suggesting that TREM2 and IL4I1 macrophages could be predominantly derived from monocytes, whereas HES1 macrophages bear an embryonic signature [141]. TREM2 macrophages were initially studied in the context of brain disorders and neurodegeneration; however, recent evidence has revealed their presence not only in the brain but also in adipose tissue, liver, and different types of tumors, indicating a potential immunoregulation role in these contexts [142,143,144]. Several studies in mice have elucidated various roles of TREM2+ macrophages in liver disease, demonstrating protective functions for hepatocytes in MASLD and cholangiopathies [145,146], immunosuppressive roles in hepatocellular carcinoma, and contribution to supporting liver regeneration in both acute and chronic murine injury models [143,147]. In another study using scRNA-seq, researchers delineated the functional phenotypes of myeloid cells and liver macrophages throughout the progression of MASH, revealing significant alterations in both liver MDMs and their bone marrow precursors, as indicated by the downregulation of the inflammatory marker calprotectin [148].
Similarly, transcriptional profiles obtained from scRNA-seq analysis of parenchymal and non-parenchymal cells in human livers unveils distinct subsets of hepatic macrophages [63]. The first subset, CD68+MARCO- macrophages exhibit characteristics of pro-inflammatory macrophages with enriched expression of LYZ, CSTA, and CD74. The second subset, CD68+MARCO+ macrophages are identified as KCs and expressed genes associated with immune tolerance, including CD5L, MARCO, VSIG4, CD163, MAF, VCAM1, and KLF4. Furthermore, two distinct populations of MARCO+ KCs are distinguished by the expression of TIMD4, with a selective reduction of MARCO+ TIMD4- KCs observed in livers from cirrhosis patients [149].
These studies collectively suggest that macrophages exhibit a wider spectrum of phenotypic activation profiles during MASLD development than previously recognized. The integration of single-cell transcriptomics with advanced bioinformatics enables the prediction of novel cellular interactions and macrophage plasticity throughout MASLD progression.

8. Unraveling the Complexity of Hepatic Macrophages in MASLD: Insights into Spatial Dynamics

The localization of liver macrophages within hepatic lobules is closely intertwined with macrophage function. In the past decade, the technological advancements such as single-cell analysis and in situ expression measurements of landmark genes have significantly deepened our comprehension of liver macrophage populations during homeostasis and disease and revealed spatially specific responses that influence liver disease progression [150,151,152].
KCs are not restricted to blood vessels but extend into the perisinusoidal space of Disse, where they interact closely with hepatocytes and hepatic stellate cells (HSCs) [140]. Unlike KCs, MDMs are characterized by their smaller size and circulate through the sinusoids [140]. Their specific localization around the periportal area suggests that these cells may serve as primary responders to events such as bile duct leakage or the presence of pathogens in the portal vein. Hepatocyte zonation for metabolic functions is well-known, and recent studies indicate similar spatial variability in macrophages along the centrilobular–portal axis. For instance, during the weaning in mice, murine KCs tend to cluster around periportal regions, influenced by MYD88-dependent signaling activation in liver sinusoidal endothelial cells triggered by gut-derived bacteria, suggesting the significance of KC zonation in controlling pathogen dissemination [153]. In addition, spatial transcriptomics of healthy liver tissue unveils non-inflammatory macrophage genes and signatures in periportal regions and inflammatory counterparts closer to the central vein [154].
During liver injury, the hepatic macrophage landscape undergoes significant changes associated with disease stages. For instance, through conditional depletion of liver KCs, researchers demonstrate that Ly6C-high monocytes, when recruited, could replenish the KC pool with the recruited monocytes differentiating into F4/80+ monocyte-derived KCs and their subsequent CSF1R-dependent proliferation reaching the steady-state KC density by day 6 after depletion, a process occurring throughout the entire liver parenchyma without restriction to a particular hepatic zone [140]. In contrast, during MASLD development, researchers combining single-cell and -nucleus sequencing with spatial mapping have revealed distinct and evolutionarily conserved, spatially restricted hepatic macrophage niches, such as Gpnmb+Spp1+ lipid-associated macrophages (LAMs) in the centrilobular areas where steatosis occurs [155]. Additionally, they found that KC development crucially depends on their cross-talk with hepatic stellate cells via the ALK1-BMP9/10 axis [155]. Aligned with this finding, another study that utilized human liver samples form MASLD and primary sclerosing cholangitis patients combined RNA-seq and imaging cytometry, revealing intense aggregation of IBA1+CD16lowCD163low MDM-derived macrophage in nonparenchymal areas, exhibiting a distinct spatial proximity to CK19+ ductular cells in periportal areas [156]. Furthermore, they find that the loss of hepatocytes and increased ductular reaction tightly correlates with the accumulation of IBA1+CD163low MDMs, serving as a prominent immunological feature that underscores the progression of MASLD, primary sclerosing cholangitis, primary biliary cholangitis, and alcoholic hepatitis.

9. Targeting Macrophages for the Treatment of MASLD

Macrophages have emerged as important therapeutic targets in MASLD. The impact of Kupffer cells on steatosis, to some extent, is regulated by IL-1beta-mediated suppression of PPARα expression and activity, a master regulator of fatty acid oxidation in the liver [157,158]. Depletion of resident KCs in rats using gadolinium chloride has shown a protective role against diet-induced alterations in hepatic lipid metabolism and insulin sensitivity [159]. In monocytes/macrophages and KCs, the glucocorticoid receptor (GR) and glucocorticoid induced leucine zipper (GILZ) axis is involved in a variety of inflammatory processes, significantly contributing to the pathogenesis of diet-induced liver inflammation. Knockdown of Gilz renders KCs more susceptible to LPS, and transgenic mice overexpressing macrophage-specific Gilz significantly reduce obesity-induced liver inflammation [160].
Dexamethasone, a potent glucocorticoid widely used to treat diseases including multiple sclerosis, allergies, cerebral edema, inflammation, and shock, has been explored in the context of MASLD treatment [161]. Svendsen et al. demonstrated that a low dose of an anti-CD163-IgG-dexamethasone conjugate, specifically targeting CD163 receptors on KCs and alternatively activated macrophages, significantly reduces high-fructose diet-induced MASH like pathologies, including hepatocyte ballooning, hepatic inflammation, and fibrosis in rats, without apparent systemic side effects [137]. Galectin-3, a member of the endogenous lectin family with the ability to bind to terminal galactose residues in glycoproteins, has been extensively studied for its role in acute inflammation. It modulates immune cell adhesion and migration, cytokine production, phagocytosis, and immune cells survival [162,163,164,165]. Traber et al. investigated two galectin-3 binding carbohydrate drugs to treat MASH in a high-fat diet-fed mouse model. They reported that galectin-3 targeting drugs ameliorated MASH pathologies, suggesting a mechanistical involvement of macrophages [166].

10. Conclusions

Hepatic macrophages are a crucial cellular component of the liver and play vital roles in liver function, contributing significantly to tissue homeostasis and swift responses during pathophysiological conditions. While significant progress has been made in recent decades regarding the role of macrophages in MASLD, the functional activation of macrophages in the disease remains inadequately understood. Despite the identification of various proteins and signaling pathways implicated in macrophage activation, a comprehensive understanding of the coordinated regulatory mechanisms is still lacking. A more detailed examination of how macrophage phenotypes evolve over time could provide insights into their specific roles during distinct phases of the disease, from early steatosis to advanced fibrosis. Liver macrophages represent a heterogeneous population of phagocytes with specific adaptations in their phenotypes to the microenvironment. Therefore, fully capturing the functional contribution of a macrophage population to disease progression or regression in MASLD requires consideration of their spatial contextualization. Given the complexity of the hepatic macrophage landscape and the rapid technological advancements in macrophage biology studies, reaching a consensus on macrophage denominations to decipher the overarching functions of specific subpopulations would be advantageous. Further developments are expected to enable the acquisition of multiple omics data from a single sample on a large scale and with a high number of parameters, such as immunostaining combined with in situ messenger RNA sequencing. These advancements hold the potential to revolutionize our understanding of liver macrophage biology, ultimately facilitating the development of personalized and targeted interventions for MASLD and other liver diseases.

Author Contributions

F.A. performed the literature search and wrote the first draft. N.E., E.J. and Y.Z. edited the manuscript. All authors have read and agreed to the published version of the manuscript.

Funding

This research received funding from National Institutes of Health grants R01DK119131 and K22CA184146. The content is solely the responsibility of the authors and does not necessarily represent the official views of the National Institutes of Health.

Institutional Review Board Statement

Not applicable.

Informed Consent Statement

Not applicable.

Data Availability Statement

Not applicable.

Conflicts of Interest

The author declares no conflict of interest.

Abbreviations

ARG1: Arginase 1; BMDMs: Bone Marrow Derived Macrophages; CCL2: Chemokine (C-C Motif) Ligand 2; COL1A1: Collagen Type 1 Alpha 1; COL1A2: Collagen Type 1 Alpha 2; DAMP: Damage-Associated Molecular Patterns; HCC: Hepatocellular Carcinoma; HSC: Hepatic Stellate Cell; IFNg: Interferon Gamma; IL: Interleukin; KC: Kupffer cell; LPS: Lipopolysaccharide; MAPK: Mitogen-Activated Protein Kinase; MASH: Metabolic dysfunction-Associated Steatohepatitis; MASL: Metabolic dysfunction-Associated Steatotic Liver; MASLD: Metabolic dysfunction-Associated Steatotic Liver Disease; MCD: Methionine Choline Deficient; MDM: Monocyte Derived Macrophage; NAFL: Nonalcoholic Fatty Liver; NAFLD: Nonalcoholic Fatty Liver Disease; NASH: Nonalcoholic Steatohepatitis; NF-κB: Nuclear Factor Kappa B; NLRP3: NLR Family Pyrin Domain Containing 3; NOS2: Nitric Oxide Synthase 2; NPC: Non-parenchymal Cell; PA: Palmitic Acid; PPAR: Peroxisome Proliferator-Activated Receptor; ROS: Reactive Oxygen Species; SLD: Steatotic Liver Disease; T2DM: Type II Diabetes Mellitus; TGF-b: Transforming Growth Factor Beta; TLR: Toll-Like Receptor; TNFa: Tumor Necrosis Factor Alpha

References

  1. Rinella, M.E.; Sookoian, S. From NAFLD to MASLD: updated naming and diagnosis criteria for fatty liver disease. J Lipid Res 2024, 65, 100485. [Google Scholar] [CrossRef]
  2. Younossi, Z.; Tacke, F.; Arrese, M.; Chander Sharma, B.; Mostafa, I.; Bugianesi, E.; Wai-Sun Wong, V.; Yilmaz, Y.; George, J.; Fan, J.; et al. Global Perspectives on Nonalcoholic Fatty Liver Disease and Nonalcoholic Steatohepatitis. Hepatology 2019, 69, 2672–2682. [Google Scholar] [CrossRef]
  3. Schuppan, D.; Surabattula, R.; Wang, X.Y. Determinants of fibrosis progression and regression in NASH. J Hepatol 2018, 68, 238–250. [Google Scholar] [CrossRef]
  4. Fan, J.G.; Kim, S.U.; Wong, V.W. New trends on obesity and NAFLD in Asia. J Hepatol 2017, 67, 862–873. [Google Scholar] [CrossRef]
  5. Estes, C.; Anstee, Q.M.; Arias-Loste, M.T.; Bantel, H.; Bellentani, S.; Caballeria, J.; Colombo, M.; Craxi, A.; Crespo, J.; Day, C.P.; et al. Modeling NAFLD disease burden in China, France, Germany, Italy, Japan, Spain, United Kingdom, and United States for the period 2016-2030. J Hepatol 2018, 69, 896–904. [Google Scholar] [CrossRef]
  6. Wong, R.J.; Aguilar, M.; Cheung, R.; Perumpail, R.B.; Harrison, S.A.; Younossi, Z.M.; Ahmed, A. Nonalcoholic steatohepatitis is the second leading etiology of liver disease among adults awaiting liver transplantation in the United States. Gastroenterology 2015, 148, 547–555. [Google Scholar] [CrossRef]
  7. Williams, C.D.; Stengel, J.; Asike, M.I.; Torres, D.M.; Shaw, J.; Contreras, M.; Landt, C.L.; Harrison, S.A. Prevalence of nonalcoholic fatty liver disease and nonalcoholic steatohepatitis among a largely middle-aged population utilizing ultrasound and liver biopsy: a prospective study. Gastroenterology 2011, 140, 124–131. [Google Scholar] [CrossRef]
  8. Sayiner, M.; Koenig, A.; Henry, L.; Younossi, Z.M. Epidemiology of Nonalcoholic Fatty Liver Disease and Nonalcoholic Steatohepatitis in the United States and the Rest of the World. Clin Liver Dis 2016, 20, 205–214. [Google Scholar] [CrossRef]
  9. Walker, R.W.; Le, K.A.; Davis, J.; Alderete, T.L.; Cherry, R.; Lebel, S.; Goran, M.I. High rates of fructose malabsorption are associated with reduced liver fat in obese African Americans. J Am Coll Nutr 2012, 31, 369–374. [Google Scholar] [CrossRef]
  10. Younossi, Z.M.; Blissett, D.; Blissett, R.; Henry, L.; Stepanova, M.; Younossi, Y.; Racila, A.; Hunt, S.; Beckerman, R. The economic and clinical burden of nonalcoholic fatty liver disease in the United States and Europe. Hepatology 2016, 64, 1577–1586. [Google Scholar] [CrossRef]
  11. Younossi, Z.M.; Tampi, R.; Priyadarshini, M.; Nader, F.; Younossi, I.M.; Racila, A. Burden of Illness and Economic Model for Patients With Nonalcoholic Steatohepatitis in the United States. Hepatology 2019, 69, 564–572. [Google Scholar] [CrossRef]
  12. Magee, N.; Zou, A.; Zhang, Y. Pathogenesis of Nonalcoholic Steatohepatitis: Interactions between Liver Parenchymal and Nonparenchymal Cells. Biomed Res Int 2016, 2016, 5170402. [Google Scholar] [CrossRef]
  13. Day, C.P.; James, O.F. Steatohepatitis: a tale of two "hits"? Gastroenterology 1998, 114, 842–845. [Google Scholar] [CrossRef]
  14. Day, C.P. From fat to inflammation. Gastroenterology 2006, 130, 207–210. [Google Scholar] [CrossRef]
  15. Jou, J.; Choi, S.S.; Diehl, A.M. Mechanisms of disease progression in nonalcoholic fatty liver disease. Semin Liver Dis 2008, 28, 370–379. [Google Scholar] [CrossRef]
  16. Dowman, J.K.; Tomlinson, J.W.; Newsome, P.N. Pathogenesis of non-alcoholic fatty liver disease. QJM 2010, 103, 71–83. [Google Scholar] [CrossRef]
  17. Roskams, T.; Yang, S.Q.; Koteish, A.; Durnez, A.; DeVos, R.; Huang, X.; Achten, R.; Verslype, C.; Diehl, A.M. Oxidative stress and oval cell accumulation in mice and humans with alcoholic and nonalcoholic fatty liver disease. Am J Pathol 2003, 163, 1301–1311. [Google Scholar] [CrossRef]
  18. Lopez, B.G.; Tsai, M.S.; Baratta, J.L.; Longmuir, K.J.; Robertson, R.T. Characterization of Kupffer cells in livers of developing mice. Comp Hepatol 2011, 10, 2. [Google Scholar] [CrossRef]
  19. Tacke, F.; Zimmermann, H.W. Macrophage heterogeneity in liver injury and fibrosis. J Hepatol 2014, 60, 1090–1096. [Google Scholar] [CrossRef]
  20. Arrese, M.; Cabrera, D.; Kalergis, A.M.; Feldstein, A.E. Innate Immunity and Inflammation in NAFLD/NASH. Dig Dis Sci 2016, 61, 1294–1303. [Google Scholar] [CrossRef]
  21. Sutti, S.; Bruzzi, S.; Albano, E. The role of immune mechanisms in alcoholic and nonalcoholic steatohepatitis: a 2015 update. Expert Rev Gastroenterol Hepatol 2016, 10, 243–253. [Google Scholar] [CrossRef]
  22. Metchnikoff, Elie - Elie Metchnikoff.
  23. van Furth, R.; Cohn, Z.A.; Hirsch, J.G.; Humphrey, J.H.; Spector, W.G.; Langevoort, H.L. The mononuclear phagocyte system: a new classification of macrophages, monocytes, and their precursor cells. Bull World Health Organ 1972, 46, 845–852. [Google Scholar]
  24. Hume, D.A. The mononuclear phagocyte system. Curr Opin Immunol 2006, 18, 49–53. [Google Scholar] [CrossRef]
  25. Gautier, E.L.; Shay, T.; Miller, J.; Greter, M.; Jakubzick, C.; Ivanov, S.; Helft, J.; Chow, A.; Elpek, K.G.; Gordonov, S.; et al. Gene-expression profiles and transcriptional regulatory pathways that underlie the identity and diversity of mouse tissue macrophages. Nat Immunol 2012, 13, 1118–1128. [Google Scholar] [CrossRef]
  26. Wynn, T.A.; Chawla, A.; Pollard, J.W. Macrophage biology in development, homeostasis and disease. Nature 2013, 496, 445–455. [Google Scholar] [CrossRef]
  27. Schulz, C.; Gomez Perdiguero, E.; Chorro, L.; Szabo-Rogers, H.; Cagnard, N.; Kierdorf, K.; Prinz, M.; Wu, B.; Jacobsen, S.E.; Pollard, J.W.; et al. A lineage of myeloid cells independent of Myb and hematopoietic stem cells. Science 2012, 336, 86–90. [Google Scholar] [CrossRef]
  28. Yona, S.; Kim, K.W.; Wolf, Y.; Mildner, A.; Varol, D.; Breker, M.; Strauss-Ayali, D.; Viukov, S.; Guilliams, M.; Misharin, A.; et al. Fate mapping reveals origins and dynamics of monocytes and tissue macrophages under homeostasis. Immunity 2013, 38, 79–91. [Google Scholar] [CrossRef]
  29. Hirayama, D.; Iida, T.; Nakase, H. The Phagocytic Function of Macrophage-Enforcing Innate Immunity and Tissue Homeostasis. Int J Mol Sci 2017, 19. [Google Scholar] [CrossRef]
  30. Ley, K.; Pramod, A.B.; Croft, M.; Ravichandran, K.S.; Ting, J.P. How Mouse Macrophages Sense What Is Going On. Front Immunol 2016, 7, 204. [Google Scholar] [CrossRef]
  31. Gordon, S. Phagocytosis: The Legacy of Metchnikoff. Cell 2016, 166, 1065–1068. [Google Scholar] [CrossRef]
  32. Biswas, S.K.; Mantovani, A. Macrophage plasticity and interaction with lymphocyte subsets: cancer as a paradigm. Nat Immunol 2010, 11, 889–896. [Google Scholar] [CrossRef]
  33. Park, M.D.; Silvin, A.; Ginhoux, F.; Merad, M. Macrophages in health and disease. Cell 2022, 185, 4259–4279. [Google Scholar] [CrossRef]
  34. Cohen, M.; Giladi, A.; Gorki, A.D.; Solodkin, D.G.; Zada, M.; Hladik, A.; Miklosi, A.; Salame, T.M.; Halpern, K.B.; David, E.; et al. Lung Single-Cell Signaling Interaction Map Reveals Basophil Role in Macrophage Imprinting. Cell 2018, 175, 1031–1044. [Google Scholar] [CrossRef]
  35. Amit, I.; Winter, D.R.; Jung, S. The role of the local environment and epigenetics in shaping macrophage identity and their effect on tissue homeostasis. Nat Immunol 2016, 17, 18–25. [Google Scholar] [CrossRef]
  36. Gosselin, D.; Link, V.M.; Romanoski, C.E.; Fonseca, G.J.; Eichenfield, D.Z.; Spann, N.J.; Stender, J.D.; Chun, H.B.; Garner, H.; Geissmann, F.; et al. Environment drives selection and function of enhancers controlling tissue-specific macrophage identities. Cell 2014, 159, 1327–1340. [Google Scholar] [CrossRef]
  37. Lavin, Y.; Merad, M. Macrophages: gatekeepers of tissue integrity. Cancer Immunol Res 2013, 1, 201–209. [Google Scholar] [CrossRef]
  38. Pollard, J.W. Trophic macrophages in development and disease. Nat Rev Immunol 2009, 9, 259–270. [Google Scholar] [CrossRef]
  39. Wiktor-Jedrzejczak, W.; Bartocci, A.; Ferrante, A.W., Jr.; Ahmed-Ansari, A.; Sell, K.W.; Pollard, J.W.; Stanley, E.R. Total absence of colony-stimulating factor 1 in the macrophage-deficient osteopetrotic (op/op) mouse. Proc Natl Acad Sci U S A 1990, 87, 4828–4832. [Google Scholar] [CrossRef]
  40. Bessis, M. [Erythroblastic island, functional unity of bone marrow]. Rev Hematol 1958, 13, 8–11. [Google Scholar]
  41. Chasis, J.A.; Mohandas, N. Erythroblastic islands: niches for erythropoiesis. Blood 2008, 112, 470–478. [Google Scholar] [CrossRef]
  42. Sadahira, Y.; Yoshino, T.; Monobe, Y. Very late activation antigen 4-vascular cell adhesion molecule 1 interaction is involved in the formation of erythroblastic islands. J Exp Med 1995, 181, 411–415. [Google Scholar] [CrossRef]
  43. Bessis, M.C.; Breton-Gorius, J. Iron metabolism in the bone marrow as seen by electron microscopy: a critical review. Blood 1962, 19, 635–663. [Google Scholar] [CrossRef]
  44. Skutelsky, E.; Danon, D. On the expulsion of the erythroid nucleus and its phagocytosis. Anat Rec 1972, 173, 123–126. [Google Scholar] [CrossRef]
  45. Colonna, M.; Butovsky, O. Microglia Function in the Central Nervous System During Health and Neurodegeneration. Annu Rev Immunol 2017, 35, 441–468. [Google Scholar] [CrossRef]
  46. Erblich, B.; Zhu, L.; Etgen, A.M.; Dobrenis, K.; Pollard, J.W. Absence of colony stimulation factor-1 receptor results in loss of microglia, disrupted brain development and olfactory deficits. PLoS One 2011, 6, e26317. [Google Scholar] [CrossRef]
  47. Whitsett, J.A.; Wert, S.E.; Weaver, T.E. Alveolar surfactant homeostasis and the pathogenesis of pulmonary disease. Annu Rev Med 2010, 61, 105–119. [Google Scholar] [CrossRef]
  48. Wake, K. Karl Wilhelm Kupffer And His Contributions To Modern Hepatology. Comp Hepatol 2004, 3 Suppl 1, S2. [Google Scholar] [CrossRef]
  49. Bilzer, M.; Roggel, F.; Gerbes, A.L. Role of Kupffer cells in host defense and liver disease. Liver Int 2006, 26, 1175–1186. [Google Scholar] [CrossRef]
  50. Bouwens, L.; Baekeland, M.; De Zanger, R.; Wisse, E. Quantitation, tissue distribution and proliferation kinetics of Kupffer cells in normal rat liver. Hepatology 1986, 6, 718–722. [Google Scholar] [CrossRef]
  51. MacPhee, P.J.; Schmidt, E.E.; Groom, A.C. Evidence for Kupffer cell migration along liver sinusoids, from high-resolution in vivo microscopy. Am J Physiol 1992, 263, G17–23. [Google Scholar] [CrossRef]
  52. Wen, Y.; Lambrecht, J.; Ju, C.; Tacke, F. Hepatic macrophages in liver homeostasis and diseases-diversity, plasticity and therapeutic opportunities. Cell Mol Immunol 2021, 18, 45–56. [Google Scholar] [CrossRef]
  53. Krenkel, O.; Tacke, F. Liver macrophages in tissue homeostasis and disease. Nat Rev Immunol 2017, 17, 306–321. [Google Scholar] [CrossRef]
  54. Zigmond, E.; Samia-Grinberg, S.; Pasmanik-Chor, M.; Brazowski, E.; Shibolet, O.; Halpern, Z.; Varol, C. Infiltrating monocyte-derived macrophages and resident kupffer cells display different ontogeny and functions in acute liver injury. J Immunol 2014, 193, 344–353. [Google Scholar] [CrossRef]
  55. Scott, C.L.; Zheng, F.; De Baetselier, P.; Martens, L.; Saeys, Y.; De Prijck, S.; Lippens, S.; Abels, C.; Schoonooghe, S.; Raes, G.; et al. Bone marrow-derived monocytes give rise to self-renewing and fully differentiated Kupffer cells. Nat Commun 2016, 7, 10321. [Google Scholar] [CrossRef]
  56. Fogg, D.K.; Sibon, C.; Miled, C.; Jung, S.; Aucouturier, P.; Littman, D.R.; Cumano, A.; Geissmann, F. A clonogenic bone marrow progenitor specific for macrophages and dendritic cells. Science 2006, 311, 83–87. [Google Scholar] [CrossRef]
  57. Kim, K.W.; Zhang, N.; Choi, K.; Randolph, G.J. Homegrown Macrophages. Immunity 2016, 45, 468–470. [Google Scholar] [CrossRef]
  58. Hoeffel, G.; Chen, J.; Lavin, Y.; Low, D.; Almeida, F.F.; See, P.; Beaudin, A.E.; Lum, J.; Low, I.; Forsberg, E.C.; et al. C-Myb(+) erythro-myeloid progenitor-derived fetal monocytes give rise to adult tissue-resident macrophages. Immunity 2015, 42, 665–678. [Google Scholar] [CrossRef]
  59. Gomez Perdiguero, E.; Klapproth, K.; Schulz, C.; Busch, K.; Azzoni, E.; Crozet, L.; Garner, H.; Trouillet, C.; de Bruijn, M.F.; Geissmann, F.; et al. Tissue-resident macrophages originate from yolk-sac-derived erythro-myeloid progenitors. Nature 2015, 518, 547–551. [Google Scholar] [CrossRef]
  60. Bouwens, L.; Knook, D.L.; Wisse, E. Local proliferation and extrahepatic recruitment of liver macrophages (Kupffer cells) in partial-body irradiated rats. J Leukoc Biol 1986, 39, 687–697. [Google Scholar] [CrossRef]
  61. Wacker, H.H.; Radzun, H.J.; Parwaresch, M.R. Kinetics of Kupffer cells as shown by parabiosis and combined autoradiographic/immunohistochemical analysis. Virchows Arch B Cell Pathol Incl Mol Pathol 1986, 51, 71–78. [Google Scholar] [CrossRef]
  62. Yamamoto, T.; Naito, M.; Moriyama, H.; Umezu, H.; Matsuo, H.; Kiwada, H.; Arakawa, M. Repopulation of murine Kupffer cells after intravenous administration of liposome-encapsulated dichloromethylene diphosphonate. Am J Pathol 1996, 149, 1271–1286. [Google Scholar]
  63. MacParland, S.A.; Liu, J.C.; Ma, X.Z.; Innes, B.T.; Bartczak, A.M.; Gage, B.K.; Manuel, J.; Khuu, N.; Echeverri, J.; Linares, I.; et al. Single cell RNA sequencing of human liver reveals distinct intrahepatic macrophage populations. Nat Commun 2018, 9, 4383. [Google Scholar] [CrossRef]
  64. Zhao, J.; Zhang, S.; Liu, Y.; He, X.; Qu, M.; Xu, G.; Wang, H.; Huang, M.; Pan, J.; Liu, Z.; et al. Single-cell RNA sequencing reveals the heterogeneity of liver-resident immune cells in human. Cell Discov 2020, 6, 22. [Google Scholar] [CrossRef]
  65. Heymann, F.; Peusquens, J.; Ludwig-Portugall, I.; Kohlhepp, M.; Ergen, C.; Niemietz, P.; Martin, C.; van Rooijen, N.; Ochando, J.C.; Randolph, G.J.; et al. Liver inflammation abrogates immunological tolerance induced by Kupffer cells. Hepatology 2015, 62, 279–291. [Google Scholar] [CrossRef]
  66. You, Q.; Cheng, L.; Kedl, R.M.; Ju, C. Mechanism of T cell tolerance induction by murine hepatic Kupffer cells. Hepatology 2008, 48, 978–990. [Google Scholar] [CrossRef]
  67. Magee, N.; Ahamed, F.; Eppler, N.; Jones, E.; Ghosh, P.; He, L.; Zhang, Y. Hepatic transcriptome profiling reveals early signatures associated with disease transition from non-alcoholic steatosis to steatohepatitis. Liver Res 2022, 6, 238–250. [Google Scholar] [CrossRef]
  68. Obstfeld, A.E.; Sugaru, E.; Thearle, M.; Francisco, A.M.; Gayet, C.; Ginsberg, H.N.; Ables, E.V.; Ferrante, A.W., Jr. C-C chemokine receptor 2 (CCR2) regulates the hepatic recruitment of myeloid cells that promote obesity-induced hepatic steatosis. Diabetes 2010, 59, 916–925. [Google Scholar] [CrossRef]
  69. Ni, Y.; Nagashimada, M.; Zhuge, F.; Zhan, L.; Nagata, N.; Tsutsui, A.; Nakanuma, Y.; Kaneko, S.; Ota, T. Astaxanthin prevents and reverses diet-induced insulin resistance and steatohepatitis in mice: A comparison with vitamin E. Sci Rep 2015, 5, 17192. [Google Scholar] [CrossRef]
  70. Devisscher, L.; Scott, C.L.; Lefere, S.; Raevens, S.; Bogaerts, E.; Paridaens, A.; Verhelst, X.; Geerts, A.; Guilliams, M.; Van Vlierberghe, H. Non-alcoholic steatohepatitis induces transient changes within the liver macrophage pool. Cell Immunol 2017, 322, 74–83. [Google Scholar] [CrossRef]
  71. Lefere, S.; Degroote, H.; Van Vlierberghe, H.; Devisscher, L. Unveiling the depletion of Kupffer cells in experimental hepatocarcinogenesis through liver macrophage subtype-specific markers. J Hepatol 2019, 71, 631–633. [Google Scholar] [CrossRef]
  72. Tosello-Trampont, A.C.; Landes, S.G.; Nguyen, V.; Novobrantseva, T.I.; Hahn, Y.S. Kuppfer cells trigger nonalcoholic steatohepatitis development in diet-induced mouse model through tumor necrosis factor-alpha production. J Biol Chem 2012, 287, 40161–40172. [Google Scholar] [CrossRef]
  73. Yoshimura, T.; Robinson, E.A.; Tanaka, S.; Appella, E.; Leonard, E.J. Purification and amino acid analysis of two human monocyte chemoattractants produced by phytohemagglutinin-stimulated human blood mononuclear leukocytes. J Immunol 1989, 142, 1956–1962. [Google Scholar] [CrossRef]
  74. Deshmane, S.L.; Kremlev, S.; Amini, S.; Sawaya, B.E. Monocyte chemoattractant protein-1 (MCP-1): an overview. J Interferon Cytokine Res 2009, 29, 313–326. [Google Scholar] [CrossRef]
  75. Bartoli, C.; Civatte, M.; Pellissier, J.F.; Figarella-Branger, D. CCR2A and CCR2B, the two isoforms of the monocyte chemoattractant protein-1 receptor are up-regulated and expressed by different cell subsets in idiopathic inflammatory myopathies. Acta Neuropathol 2001, 102, 385–392. [Google Scholar] [CrossRef]
  76. Morinaga, H.; Mayoral, R.; Heinrichsdorff, J.; Osborn, O.; Franck, N.; Hah, N.; Walenta, E.; Bandyopadhyay, G.; Pessentheiner, A.R.; Chi, T.J.; et al. Characterization of distinct subpopulations of hepatic macrophages in HFD/obese mice. Diabetes 2015, 64, 1120–1130. [Google Scholar] [CrossRef]
  77. Krenkel, O.; Puengel, T.; Govaere, O.; Abdallah, A.T.; Mossanen, J.C.; Kohlhepp, M.; Liepelt, A.; Lefebvre, E.; Luedde, T.; Hellerbrand, C.; et al. Therapeutic inhibition of inflammatory monocyte recruitment reduces steatohepatitis and liver fibrosis. Hepatology 2018, 67, 1270–1283. [Google Scholar] [CrossRef]
  78. Park, J.W.; Jeong, G.; Kim, S.J.; Kim, M.K.; Park, S.M. Predictors reflecting the pathological severity of non-alcoholic fatty liver disease: comprehensive study of clinical and immunohistochemical findings in younger Asian patients. J Gastroenterol Hepatol 2007, 22, 491–497. [Google Scholar] [CrossRef]
  79. Lotowska, J.M.; Sobaniec-Lotowska, M.E.; Lebensztejn, D.M. The role of Kupffer cells in the morphogenesis of nonalcoholic steatohepatitis - ultrastructural findings. The first report in pediatric patients. Scand J Gastroenterol 2013, 48, 352–357. [Google Scholar] [CrossRef]
  80. Gadd, V.L.; Skoien, R.; Powell, E.E.; Fagan, K.J.; Winterford, C.; Horsfall, L.; Irvine, K.; Clouston, A.D. The portal inflammatory infiltrate and ductular reaction in human nonalcoholic fatty liver disease. Hepatology 2014, 59, 1393–1405. [Google Scholar] [CrossRef]
  81. Kazankov, K.; Jorgensen, S.M.D.; Thomsen, K.L.; Moller, H.J.; Vilstrup, H.; George, J.; Schuppan, D.; Gronbaek, H. The role of macrophages in nonalcoholic fatty liver disease and nonalcoholic steatohepatitis. Nat Rev Gastroenterol Hepatol 2019, 16, 145–159. [Google Scholar] [CrossRef]
  82. Itoh, M.; Kato, H.; Suganami, T.; Konuma, K.; Marumoto, Y.; Terai, S.; Sakugawa, H.; Kanai, S.; Hamaguchi, M.; Fukaishi, T.; et al. Hepatic crown-like structure: a unique histological feature in non-alcoholic steatohepatitis in mice and humans. PLoS One 2013, 8, e82163. [Google Scholar] [CrossRef] [PubMed]
  83. Neuschwander-Tetri, B.A. Hepatic lipotoxicity and the pathogenesis of nonalcoholic steatohepatitis: the central role of nontriglyceride fatty acid metabolites. Hepatology 2010, 52, 774–788. [Google Scholar] [CrossRef] [PubMed]
  84. Pan, X.; Wang, P.; Luo, J.; Wang, Z.; Song, Y.; Ye, J.; Hou, X. Adipogenic changes of hepatocytes in a high-fat diet-induced fatty liver mice model and non-alcoholic fatty liver disease patients. Endocrine 2015, 48, 834–847. [Google Scholar] [CrossRef] [PubMed]
  85. Lee, J.Y.; Sohn, K.H.; Rhee, S.H.; Hwang, D. Saturated fatty acids, but not unsaturated fatty acids, induce the expression of cyclooxygenase-2 mediated through Toll-like receptor 4. J Biol Chem 2001, 276, 16683–16689. [Google Scholar] [CrossRef] [PubMed]
  86. Snodgrass, R.G.; Huang, S.; Choi, I.W.; Rutledge, J.C.; Hwang, D.H. Inflammasome-mediated secretion of IL-1beta in human monocytes through TLR2 activation; modulation by dietary fatty acids. J Immunol 2013, 191, 4337–4347. [Google Scholar] [CrossRef] [PubMed]
  87. Kim, S.Y.; Jeong, J.M.; Kim, S.J.; Seo, W.; Kim, M.H.; Choi, W.M.; Yoo, W.; Lee, J.H.; Shim, Y.R.; Yi, H.S.; et al. Pro-inflammatory hepatic macrophages generate ROS through NADPH oxidase 2 via endocytosis of monomeric TLR4-MD2 complex. Nat Commun 2017, 8, 2247. [Google Scholar] [CrossRef] [PubMed]
  88. Musso, G.; Gambino, R.; Cassader, M. Cholesterol metabolism and the pathogenesis of non-alcoholic steatohepatitis. Prog Lipid Res 2013, 52, 175–191. [Google Scholar] [CrossRef] [PubMed]
  89. Ioannou, G.N.; Haigh, W.G.; Thorning, D.; Savard, C. Hepatic cholesterol crystals and crown-like structures distinguish NASH from simple steatosis. J Lipid Res 2013, 54, 1326–1334. [Google Scholar] [CrossRef] [PubMed]
  90. Ioannou, G.N.; Subramanian, S.; Chait, A.; Haigh, W.G.; Yeh, M.M.; Farrell, G.C.; Lee, S.P.; Savard, C. Cholesterol crystallization within hepatocyte lipid droplets and its role in murine NASH. J Lipid Res 2017, 58, 1067–1079. [Google Scholar] [CrossRef]
  91. Miller, Y.I.; Choi, S.H.; Wiesner, P.; Fang, L.; Harkewicz, R.; Hartvigsen, K.; Boullier, A.; Gonen, A.; Diehl, C.J.; Que, X.; et al. Oxidation-specific epitopes are danger-associated molecular patterns recognized by pattern recognition receptors of innate immunity. Circ Res 2011, 108, 235–248. [Google Scholar] [CrossRef]
  92. Canbay, A.; Feldstein, A.E.; Higuchi, H.; Werneburg, N.; Grambihler, A.; Bronk, S.F.; Gores, G.J. Kupffer cell engulfment of apoptotic bodies stimulates death ligand and cytokine expression. Hepatology 2003, 38, 1188–1198. [Google Scholar] [CrossRef] [PubMed]
  93. Hirsova, P.; Ibrahim, S.H.; Krishnan, A.; Verma, V.K.; Bronk, S.F.; Werneburg, N.W.; Charlton, M.R.; Shah, V.H.; Malhi, H.; Gores, G.J. Lipid-Induced Signaling Causes Release of Inflammatory Extracellular Vesicles From Hepatocytes. Gastroenterology 2016, 150, 956–967. [Google Scholar] [CrossRef] [PubMed]
  94. Mogensen, T.H. Pathogen recognition and inflammatory signaling in innate immune defenses. Clin Microbiol Rev 2009, 22, 240–273. [Google Scholar] [CrossRef] [PubMed]
  95. Frasinariu, O.E.; Ceccarelli, S.; Alisi, A.; Moraru, E.; Nobili, V. Gut-liver axis and fibrosis in nonalcoholic fatty liver disease: an input for novel therapies. Dig Liver Dis 2013, 45, 543–551. [Google Scholar] [CrossRef] [PubMed]
  96. Budick-Harmelin, N.; Dudas, J.; Demuth, J.; Madar, Z.; Ramadori, G.; Tirosh, O. Triglycerides potentiate the inflammatory response in rat Kupffer cells. Antioxid Redox Signal 2008, 10, 2009–2022. [Google Scholar] [CrossRef]
  97. Leroux, A.; Ferrere, G.; Godie, V.; Cailleux, F.; Renoud, M.L.; Gaudin, F.; Naveau, S.; Prevot, S.; Makhzami, S.; Perlemuter, G.; et al. Toxic lipids stored by Kupffer cells correlates with their pro-inflammatory phenotype at an early stage of steatohepatitis. J Hepatol 2012, 57, 141–149. [Google Scholar] [CrossRef]
  98. Mills, C.D.; Kincaid, K.; Alt, J.M.; Heilman, M.J.; Hill, A.M. M-1/M-2 macrophages and the Th1/Th2 paradigm. J Immunol 2000, 164, 6166–6173. [Google Scholar] [CrossRef] [PubMed]
  99. Mosmann, T.R.; Coffman, R.L. TH1 and TH2 cells: different patterns of lymphokine secretion lead to different functional properties. Annu Rev Immunol 1989, 7, 145–173. [Google Scholar] [CrossRef] [PubMed]
  100. Poltorak, A.; Smirnova, I.; He, X.; Liu, M.Y.; Van Huffel, C.; McNally, O.; Birdwell, D.; Alejos, E.; Silva, M.; Du, X.; et al. Genetic and physical mapping of the Lps locus: identification of the toll-4 receptor as a candidate gene in the critical region. Blood Cells Mol Dis 1998, 24, 340–355. [Google Scholar] [CrossRef]
  101. Yang, H.; Wang, H.; Ju, Z.; Ragab, A.A.; Lundback, P.; Long, W.; Valdes-Ferrer, S.I.; He, M.; Pribis, J.P.; Li, J.; et al. MD-2 is required for disulfide HMGB1-dependent TLR4 signaling. J Exp Med 2015, 212, 5–14. [Google Scholar] [CrossRef]
  102. Jiang, D.; Liang, J.; Fan, J.; Yu, S.; Chen, S.; Luo, Y.; Prestwich, G.D.; Mascarenhas, M.M.; Garg, H.G.; Quinn, D.A.; et al. Regulation of lung injury and repair by Toll-like receptors and hyaluronan. Nat Med 2005, 11, 1173–1179. [Google Scholar] [CrossRef]
  103. Chen, X.X.; Tang, L.; Fu, Y.M.; Wang, Y.; Han, Z.H.; Meng, J.G. Paralemmin-3 contributes to lipopolysaccharide-induced inflammatory response and is involved in lipopolysaccharide-Toll-like receptor-4 signaling in alveolar macrophages. Int J Mol Med 2017, 40, 1921–1931. [Google Scholar] [CrossRef]
  104. Gong, J.; Li, J.; Dong, H.; Chen, G.; Qin, X.; Hu, M.; Yuan, F.; Fang, K.; Wang, D.; Jiang, S.; et al. Inhibitory effects of berberine on proinflammatory M1 macrophage polarization through interfering with the interaction between TLR4 and MyD88. BMC Complement Altern Med 2019, 19, 314. [Google Scholar] [CrossRef]
  105. Xiang, P.; Chen, T.; Mou, Y.; Wu, H.; Xie, P.; Lu, G.; Gong, X.; Hu, Q.; Zhang, Y.; Ji, H. NZ suppresses TLR4/NF-kappaB signalings and NLRP3 inflammasome activation in LPS-induced RAW264.7 macrophages. Inflamm Res 2015, 64, 799–808. [Google Scholar] [CrossRef]
  106. Lu, H.; Wu, L.; Liu, L.; Ruan, Q.; Zhang, X.; Hong, W.; Wu, S.; Jin, G.; Bai, Y. Quercetin ameliorates kidney injury and fibrosis by modulating M1/M2 macrophage polarization. Biochem Pharmacol 2018, 154, 203–212. [Google Scholar] [CrossRef]
  107. Qi, Q.R.; Yang, Z.M. Regulation and function of signal transducer and activator of transcription 3. World J Biol Chem 2014, 5, 231–239. [Google Scholar] [CrossRef]
  108. Wang, N.; Liang, H.; Zen, K. Molecular mechanisms that influence the macrophage m1-m2 polarization balance. Front Immunol 2014, 5, 614. [Google Scholar] [CrossRef]
  109. Sica, A.; Mantovani, A. Macrophage plasticity and polarization: in vivo veritas. J Clin Invest 2012, 122, 787–795. [Google Scholar] [CrossRef]
  110. He, Y.; Gao, Y.; Zhang, Q.; Zhou, G.; Cao, F.; Yao, S. IL-4 Switches Microglia/macrophage M1/M2 Polarization and Alleviates Neurological Damage by Modulating the JAK1/STAT6 Pathway Following ICH. Neuroscience 2020, 437, 161–171. [Google Scholar] [CrossRef] [PubMed]
  111. Quero, L.; Tiaden, A.N.; Hanser, E.; Roux, J.; Laski, A.; Hall, J.; Kyburz, D. miR-221-3p Drives the Shift of M2-Macrophages to a Pro-Inflammatory Function by Suppressing JAK3/STAT3 Activation. Front Immunol 2019, 10, 3087. [Google Scholar] [CrossRef] [PubMed]
  112. Travis, M.A.; Sheppard, D. TGF-beta activation and function in immunity. Annu Rev Immunol 2014, 32, 51–82. [Google Scholar] [CrossRef] [PubMed]
  113. Wang, L.; Li, Y.; Wang, X.; Wang, P.; Essandoh, K.; Cui, S.; Huang, W.; Mu, X.; Liu, Z.; Wang, Y.; et al. GDF3 Protects Mice against Sepsis-Induced Cardiac Dysfunction and Mortality by Suppression of Macrophage Pro-Inflammatory Phenotype. Cells 2020, 9. [Google Scholar] [CrossRef] [PubMed]
  114. Marion-Letellier, R.; Savoye, G.; Ghosh, S. Fatty acids, eicosanoids and PPAR gamma. Eur J Pharmacol 2016, 785, 44–49. [Google Scholar] [CrossRef] [PubMed]
  115. Lehrke, M.; Lazar, M.A. The many faces of PPARgamma. Cell 2005, 123, 993–999. [Google Scholar] [CrossRef] [PubMed]
  116. Villanueva, C.J.; Tontonoz, P. Licensing PPARgamma to work in macrophages. Immunity 2010, 33, 647–649. [Google Scholar] [CrossRef] [PubMed]
  117. Hou, Y.; Moreau, F.; Chadee, K. PPARgamma is an E3 ligase that induces the degradation of NFkappaB/p65. Nat Commun 2012, 3, 1300. [Google Scholar] [CrossRef] [PubMed]
  118. Luo, W.; Xu, Q.; Wang, Q.; Wu, H.; Hua, J. Effect of modulation of PPAR-gamma activity on Kupffer cells M1/M2 polarization in the development of non-alcoholic fatty liver disease. Sci Rep 2017, 7, 44612. [Google Scholar] [CrossRef] [PubMed]
  119. Bouhlel, M.A.; Derudas, B.; Rigamonti, E.; Dievart, R.; Brozek, J.; Haulon, S.; Zawadzki, C.; Jude, B.; Torpier, G.; Marx, N.; et al. PPARgamma activation primes human monocytes into alternative M2 macrophages with anti-inflammatory properties. Cell Metab 2007, 6, 137–143. [Google Scholar] [CrossRef]
  120. Odegaard, J.I.; Ricardo-Gonzalez, R.R.; Goforth, M.H.; Morel, C.R.; Subramanian, V.; Mukundan, L.; Red Eagle, A.; Vats, D.; Brombacher, F.; Ferrante, A.W.; et al. Macrophage-specific PPARgamma controls alternative activation and improves insulin resistance. Nature 2007, 447, 1116–1120. [Google Scholar] [CrossRef]
  121. Wang, C.; Ma, C.; Gong, L.; Guo, Y.; Fu, K.; Zhang, Y.; Zhou, H.; Li, Y. Macrophage Polarization and Its Role in Liver Disease. Front Immunol 2021, 12, 803037. [Google Scholar] [CrossRef]
  122. Qian, M.; Wang, S.; Guo, X.; Wang, J.; Zhang, Z.; Qiu, W.; Gao, X.; Chen, Z.; Xu, J.; Zhao, R.; et al. Hypoxic glioma-derived exosomes deliver microRNA-1246 to induce M2 macrophage polarization by targeting TERF2IP via the STAT3 and NF-kappaB pathways. Oncogene 2020, 39, 428–442. [Google Scholar] [CrossRef]
  123. Pan, Y.; Hui, X.; Hoo, R.L.C.; Ye, D.; Chan, C.Y.C.; Feng, T.; Wang, Y.; Lam, K.S.L.; Xu, A. Adipocyte-secreted exosomal microRNA-34a inhibits M2 macrophage polarization to promote obesity-induced adipose inflammation. J Clin Invest 2019, 129, 834–849. [Google Scholar] [CrossRef]
  124. Singla, R.D.; Wang, J.; Singla, D.K. Regulation of Notch 1 signaling in THP-1 cells enhances M2 macrophage differentiation. Am J Physiol Heart Circ Physiol 2014, 307, H1634–1642. [Google Scholar] [CrossRef]
  125. Byles, V.; Covarrubias, A.J.; Ben-Sahra, I.; Lamming, D.W.; Sabatini, D.M.; Manning, B.D.; Horng, T. The TSC-mTOR pathway regulates macrophage polarization. Nat Commun 2013, 4, 2834. [Google Scholar] [CrossRef] [PubMed]
  126. Yu, T.; Gao, M.; Yang, P.; Liu, D.; Wang, D.; Song, F.; Zhang, X.; Liu, Y. Insulin promotes macrophage phenotype transition through PI3K/Akt and PPAR-gamma signaling during diabetic wound healing. J Cell Physiol 2019, 234, 4217–4231. [Google Scholar] [CrossRef]
  127. Zhang, Y.; Huang, T.; Jiang, L.; Gao, J.; Yu, D.; Ge, Y.; Lin, S. MCP-induced protein 1 attenuates sepsis-induced acute lung injury by modulating macrophage polarization via the JNK/c-Myc pathway. Int Immunopharmacol 2019, 75, 105741. [Google Scholar] [CrossRef] [PubMed]
  128. Maina, V.; Sutti, S.; Locatelli, I.; Vidali, M.; Mombello, C.; Bozzola, C.; Albano, E. Bias in macrophage activation pattern influences non-alcoholic steatohepatitis (NASH) in mice. Clin Sci (Lond) 2012, 122, 545–553. [Google Scholar] [CrossRef]
  129. Wan, J.; Benkdane, M.; Teixeira-Clerc, F.; Bonnafous, S.; Louvet, A.; Lafdil, F.; Pecker, F.; Tran, A.; Gual, P.; Mallat, A.; et al. M2 Kupffer cells promote M1 Kupffer cell apoptosis: a protective mechanism against alcoholic and nonalcoholic fatty liver disease. Hepatology 2014, 59, 130–142. [Google Scholar] [CrossRef] [PubMed]
  130. Papackova, Z.; Palenickova, E.; Dankova, H.; Zdychova, J.; Skop, V.; Kazdova, L.; Cahova, M. Kupffer cells ameliorate hepatic insulin resistance induced by high-fat diet rich in monounsaturated fatty acids: the evidence for the involvement of alternatively activated macrophages. Nutr Metab (Lond) 2012, 9, 22. [Google Scholar] [CrossRef]
  131. Odegaard, J.I.; Ricardo-Gonzalez, R.R.; Red Eagle, A.; Vats, D.; Morel, C.R.; Goforth, M.H.; Subramanian, V.; Mukundan, L.; Ferrante, A.W.; Chawla, A. Alternative M2 activation of Kupffer cells by PPARdelta ameliorates obesity-induced insulin resistance. Cell Metab 2008, 7, 496–507. [Google Scholar] [CrossRef]
  132. Rolny, C.; Mazzone, M.; Tugues, S.; Laoui, D.; Johansson, I.; Coulon, C.; Squadrito, M.L.; Segura, I.; Li, X.; Knevels, E.; et al. HRG inhibits tumor growth and metastasis by inducing macrophage polarization and vessel normalization through downregulation of PlGF. Cancer Cell 2011, 19, 31–44. [Google Scholar] [CrossRef]
  133. Bartneck, M.; Fech, V.; Ehling, J.; Govaere, O.; Warzecha, K.T.; Hittatiya, K.; Vucur, M.; Gautheron, J.; Luedde, T.; Trautwein, C.; et al. Histidine-rich glycoprotein promotes macrophage activation and inflammation in chronic liver disease. Hepatology 2016, 63, 1310–1324. [Google Scholar] [CrossRef]
  134. Rensen, S.S.; Slaats, Y.; Nijhuis, J.; Jans, A.; Bieghs, V.; Driessen, A.; Malle, E.; Greve, J.W.; Buurman, W.A. Increased hepatic myeloperoxidase activity in obese subjects with nonalcoholic steatohepatitis. Am J Pathol 2009, 175, 1473–1482. [Google Scholar] [CrossRef]
  135. Hart, K.M.; Fabre, T.; Sciurba, J.C.; Gieseck, R.L., 3rd; Borthwick, L.A.; Vannella, K.M.; Acciani, T.H.; de Queiroz Prado, R.; Thompson, R.W.; White, S.; et al. Type 2 immunity is protective in metabolic disease but exacerbates NAFLD collaboratively with TGF-beta. Sci Transl Med 2017, 9. [Google Scholar] [CrossRef] [PubMed]
  136. Nakayama, T.; Hirahara, K.; Onodera, A.; Endo, Y.; Hosokawa, H.; Shinoda, K.; Tumes, D.J.; Okamoto, Y. Th2 Cells in Health and Disease. Annu Rev Immunol 2017, 35, 53–84. [Google Scholar] [CrossRef] [PubMed]
  137. Svendsen, P.; Graversen, J.H.; Etzerodt, A.; Hager, H.; Roge, R.; Gronbaek, H.; Christensen, E.I.; Moller, H.J.; Vilstrup, H.; Moestrup, S.K. Antibody-Directed Glucocorticoid Targeting to CD163 in M2-type Macrophages Attenuates Fructose-Induced Liver Inflammatory Changes. Mol Ther Methods Clin Dev 2017, 4, 50–61. [Google Scholar] [CrossRef] [PubMed]
  138. Guillot, A.; Tacke, F. Liver Macrophages: Old Dogmas and New Insights. Hepatol Commun 2019, 3, 730–743. [Google Scholar] [CrossRef]
  139. Daemen, S.; Gainullina, A.; Kalugotla, G.; He, L.; Chan, M.M.; Beals, J.W.; Liss, K.H.; Klein, S.; Feldstein, A.E.; Finck, B.N.; et al. Dynamic Shifts in the Composition of Resident and Recruited Macrophages Influence Tissue Remodeling in NASH. Cell Rep 2021, 34, 108626. [Google Scholar] [CrossRef]
  140. Bonnardel, J.; T'Jonck, W.; Gaublomme, D.; Browaeys, R.; Scott, C.L.; Martens, L.; Vanneste, B.; De Prijck, S.; Nedospasov, S.A.; Kremer, A.; et al. Stellate Cells, Hepatocytes, and Endothelial Cells Imprint the Kupffer Cell Identity on Monocytes Colonizing the Liver Macrophage Niche. Immunity 2019, 51, 638–654. [Google Scholar] [CrossRef]
  141. Mulder, K.; Patel, A.A.; Kong, W.T.; Piot, C.; Halitzki, E.; Dunsmore, G.; Khalilnezhad, S.; Irac, S.E.; Dubuisson, A.; Chevrier, M.; et al. Cross-tissue single-cell landscape of human monocytes and macrophages in health and disease. Immunity 2021, 54, 1883–1900. [Google Scholar] [CrossRef]
  142. Khantakova, D.; Brioschi, S.; Molgora, M. Exploring the Impact of TREM2 in Tumor-Associated Macrophages. Vaccines (Basel) 2022, 10. [Google Scholar] [CrossRef]
  143. Zhou, L.; Wang, M.; Guo, H.; Hou, J.; Zhang, Y.; Li, M.; Wu, X.; Chen, X.; Wang, L. Integrated Analysis Highlights the Immunosuppressive Role of TREM2(+) Macrophages in Hepatocellular Carcinoma. Front Immunol 2022, 13, 848367. [Google Scholar] [CrossRef] [PubMed]
  144. Colonna, M. The biology of TREM receptors. Nat Rev Immunol 2023, 23, 580–594. [Google Scholar] [CrossRef]
  145. Labiano, I.; Agirre-Lizaso, A.; Olaizola, P.; Echebarria, A.; Huici-Izagirre, M.; Olaizola, I.; Esparza-Baquer, A.; Sharif, O.; Hijona, E.; Milkiewicz, P.; et al. TREM-2 plays a protective role in cholestasis by acting as a negative regulator of inflammation. J Hepatol 2022, 77, 991–1004. [Google Scholar] [CrossRef]
  146. Hendrikx, T.; Porsch, F.; Kiss, M.G.; Rajcic, D.; Papac-Milicevic, N.; Hoebinger, C.; Goederle, L.; Hladik, A.; Shaw, L.E.; Horstmann, H.; et al. Soluble TREM2 levels reflect the recruitment and expansion of TREM2(+) macrophages that localize to fibrotic areas and limit NASH. J Hepatol 2022, 77, 1373–1385. [Google Scholar] [CrossRef]
  147. Coelho, I.; Duarte, N.; Barros, A.; Macedo, M.P.; Penha-Goncalves, C. Trem-2 Promotes Emergence of Restorative Macrophages and Endothelial Cells During Recovery From Hepatic Tissue Damage. Front Immunol 2020, 11, 616044. [Google Scholar] [CrossRef]
  148. Krenkel, O.; Hundertmark, J.; Abdallah, A.T.; Kohlhepp, M.; Puengel, T.; Roth, T.; Branco, D.P.P.; Mossanen, J.C.; Luedde, T.; Trautwein, C.; et al. Myeloid cells in liver and bone marrow acquire a functionally distinct inflammatory phenotype during obesity-related steatohepatitis. Gut 2020, 69, 551–563. [Google Scholar] [CrossRef] [PubMed]
  149. Ramachandran, P.; Dobie, R.; Wilson-Kanamori, J.R.; Dora, E.F.; Henderson, B.E.P.; Luu, N.T.; Portman, J.R.; Matchett, K.P.; Brice, M.; Marwick, J.A.; et al. Resolving the fibrotic niche of human liver cirrhosis at single-cell level. Nature 2019, 575, 512–518. [Google Scholar] [CrossRef] [PubMed]
  150. Halpern, K.B.; Shenhav, R.; Matcovitch-Natan, O.; Toth, B.; Lemze, D.; Golan, M.; Massasa, E.E.; Baydatch, S.; Landen, S.; Moor, A.E.; et al. Single-cell spatial reconstruction reveals global division of labour in the mammalian liver. Nature 2017, 542, 352–356. [Google Scholar] [CrossRef]
  151. Satija, R.; Farrell, J.A.; Gennert, D.; Schier, A.F.; Regev, A. Spatial reconstruction of single-cell gene expression data. Nat Biotechnol 2015, 33, 495–502. [Google Scholar] [CrossRef]
  152. Guillot, A.; Tacke, F. Spatial dimension of macrophage heterogeneity in liver diseases. eGastroenterology 2023, 1, e000003. [Google Scholar] [CrossRef]
  153. Gola, A.; Dorrington, M.G.; Speranza, E.; Sala, C.; Shih, R.M.; Radtke, A.J.; Wong, H.S.; Baptista, A.P.; Hernandez, J.M.; Castellani, G.; et al. Commensal-driven immune zonation of the liver promotes host defence. Nature 2021, 589, 131–136. [Google Scholar] [CrossRef] [PubMed]
  154. Andrews, T.S.; Atif, J.; Liu, J.C.; Perciani, C.T.; Ma, X.Z.; Thoeni, C.; Slyper, M.; Eraslan, G.; Segerstolpe, A.; Manuel, J.; et al. Single-Cell, Single-Nucleus, and Spatial RNA Sequencing of the Human Liver Identifies Cholangiocyte and Mesenchymal Heterogeneity. Hepatol Commun 2022, 6, 821–840. [Google Scholar] [CrossRef] [PubMed]
  155. Guilliams, M.; Bonnardel, J.; Haest, B.; Vanderborght, B.; Wagner, C.; Remmerie, A.; Bujko, A.; Martens, L.; Thone, T.; Browaeys, R.; et al. Spatial proteogenomics reveals distinct and evolutionarily conserved hepatic macrophage niches. Cell 2022, 185, 379–396. [Google Scholar] [CrossRef] [PubMed]
  156. Guillot, A.; Winkler, M.; Silva Afonso, M.; Aggarwal, A.; Lopez, D.; Berger, H.; Kohlhepp, M.S.; Liu, H.; Ozdirik, B.; Eschrich, J.; et al. Mapping the hepatic immune landscape identifies monocytic macrophages as key drivers of steatohepatitis and cholangiopathy progression. Hepatology 2023, 78, 150–166. [Google Scholar] [CrossRef] [PubMed]
  157. Wang, Y.X. PPARs: diverse regulators in energy metabolism and metabolic diseases. Cell Res 2010, 20, 124–137. [Google Scholar] [CrossRef] [PubMed]
  158. Stienstra, R.; Saudale, F.; Duval, C.; Keshtkar, S.; Groener, J.E.; van Rooijen, N.; Staels, B.; Kersten, S.; Muller, M. Kupffer cells promote hepatic steatosis via interleukin-1beta-dependent suppression of peroxisome proliferator-activated receptor alpha activity. Hepatology 2010, 51, 511–522. [Google Scholar] [CrossRef] [PubMed]
  159. Huang, W.; Metlakunta, A.; Dedousis, N.; Zhang, P.; Sipula, I.; Dube, J.J.; Scott, D.K.; O'Doherty, R.M. Depletion of liver Kupffer cells prevents the development of diet-induced hepatic steatosis and insulin resistance. Diabetes 2010, 59, 347–357. [Google Scholar] [CrossRef] [PubMed]
  160. Kudo, H.; Takahara, T.; Yata, Y.; Kawai, K.; Zhang, W.; Sugiyama, T. Lipopolysaccharide triggered TNF-alpha-induced hepatocyte apoptosis in a murine non-alcoholic steatohepatitis model. J Hepatol 2009, 51, 168–175. [Google Scholar] [CrossRef]
  161. Johnson, D.B.; Lopez, M.J.; Kelley, B. Dexamethasone. In StatPearls; Treasure Island (FL) ineligible companies. Disclosure: Michael Lopez declares no relevant financial relationships with ineligible companies. Disclosure: Brendan Kelley declares no relevant financial relationships with ineligible companies, 2023. [Google Scholar]
  162. Hsu, D.K.; Yang, R.Y.; Pan, Z.; Yu, L.; Salomon, D.R.; Fung-Leung, W.P.; Liu, F.T. Targeted disruption of the galectin-3 gene results in attenuated peritoneal inflammatory responses. Am J Pathol 2000, 156, 1073–1083. [Google Scholar] [CrossRef]
  163. Liu, F.T.; Hsu, D.K.; Zuberi, R.I.; Kuwabara, I.; Chi, E.Y.; Henderson, W.R., Jr. Expression and function of galectin-3, a beta-galactoside-binding lectin, in human monocytes and macrophages. Am J Pathol 1995, 147, 1016–1028. [Google Scholar] [PubMed]
  164. Hughes, R.C. Galectins as modulators of cell adhesion. Biochimie 2001, 83, 667–676. [Google Scholar] [CrossRef] [PubMed]
  165. Di Lella, S.; Sundblad, V.; Cerliani, J.P.; Guardia, C.M.; Estrin, D.A.; Vasta, G.R.; Rabinovich, G.A. When galectins recognize glycans: from biochemistry to physiology and back again. Biochemistry 2011, 50, 7842–7857. [Google Scholar] [CrossRef]
  166. Traber, P.G.; Zomer, E. Therapy of experimental NASH and fibrosis with galectin inhibitors. PLoS One 2013, 8, e83481. [Google Scholar] [CrossRef] [PubMed]
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