Next Article in Journal
Effective Treatment of Patients Experiencing Primary, Acute HIV Infection Decreases Exhausted/Activated CD4+ T Cells and CD8+ T Memory Stem Cells
Next Article in Special Issue
The Stress-Responsive microRNA-34a Alters Insulin Signaling and Actions in Adipocytes through Induction of the Tyrosine Phosphatase PTP1B
Previous Article in Journal
Bone Disease in Multiple Myeloma: Biologic and Clinical Implications
Previous Article in Special Issue
The Endocrine Adipose Organ: A System Playing a Central Role in COVID-19
 
 
Font Type:
Arial Georgia Verdana
Font Size:
Aa Aa Aa
Line Spacing:
Column Width:
Background:
Review

Importance of the Microenvironment and Mechanosensing in Adipose Tissue Biology

1
Nutrition and Obesities: Systemic Approaches Research Group (Nutri-Omics), Sorbonne Université, INSERM, F-75013 Paris, France
2
Labex Inflamex, Université Sorbonne Paris Nord, INSERM, F-93000 Bobigny, France
3
Department of Medicine (H7), Karolinska Institutet Hospital, C2-94, 14186 Stockholm, Sweden
4
Laboratoire de Biologie du Développement (UMR 7622), IBPS, Sorbonne Université, F-75005 Paris, France
5
Assistance Publique Hôpitaux de Paris, Nutrition Department, CRNH Ile-de-France, Pitié-Salpêtrière Hospital, F-75013 Paris, France
*
Author to whom correspondence should be addressed.
Cells 2022, 11(15), 2310; https://doi.org/10.3390/cells11152310
Submission received: 16 June 2022 / Revised: 19 July 2022 / Accepted: 23 July 2022 / Published: 27 July 2022
(This article belongs to the Special Issue The Role of Adipose Tissue in Metabolic Diseases and Beyond)

Abstract

:
The expansion of adipose tissue is an adaptive mechanism that increases nutrient buffering capacity in response to an overall positive energy balance. Over the course of expansion, the adipose microenvironment undergoes continual remodeling to maintain its structural and functional integrity. However, in the long run, adipose tissue remodeling, typically characterized by adipocyte hypertrophy, immune cells infiltration, fibrosis and changes in vascular architecture, generates mechanical stress on adipose cells. This mechanical stimulus is then transduced into a biochemical signal that alters adipose function through mechanotransduction. In this review, we describe the physical changes occurring during adipose tissue remodeling, and how they regulate adipose cell physiology and promote obesity-associated dysfunction in adipose tissue.

1. Introduction

The dramatic rising incidence of obesity has increased the urgent need to understand underlying mechanisms of obesity and its link with numerous co-morbidities. Obesity is now one of the greatest public health issues affecting ~13% of adults worldwide [1]. Obesity leads to the development of complications such as type-2 diabetes, non-alcoholic fatty liver disease, cardiovascular diseases, neurodegenerative disorders, greater predisposition to serious infections (including COVID-19) and cancers [2,3,4].
Adipose tissue (AT) expansion and its altered function is the cornerstone of obesity-related complication development. AT is a highly dynamic organ that modifies its size, cellular composition, and function, upon nutritional and hormonal challenges (Appendix A). In obesity, white adipose tissue (WAT) can expand by 10 times through hypertrophy (enlarged adipocyte size) and hyperplasia (increased adipocyte number), whereas upon fasting or cold exposure, WAT shrinks by lipolysis to supply fatty acids to peripheral organs [5,6]. In addition to their storage capacity, adipocytes are able to sense the energy state and secrete factors called adipokines (including lipokines) to coordinate, and to regulate, whole body metabolism [7]. In obesity, adipocytes are exposed to different forms of stresses such as hypoxic, oxidative, metabolic, inflammatory and mechanical stressors [8,9,10]. However, the precise triggers for adipocyte dysfunction and their coordinated mode of action still remain uncertain. Nevertheless, there is little doubt that adipocyte hypertrophy is a major factor of adipocyte dysfunction, suggesting that this stimulus requires careful attention. It is indeed known that the fat cell size correlates with the severity of insulin resistance and type 2 diabetes mellitus as well as altered responses to weight loss [11,12,13,14,15].
AT expansion is a complex biological process consisting of major coordinated and intertwined events: (i) increase in the size of adipocytes triggered by excessive lipogenesis and triglyceride storage, (ii) infiltration of immune cells in the parenchyma and around adipocytes, (iii) remodeling of the extracellular matrix (ECM), (iv) formation of new adipocytes by adipogenesis from progenitor cells (Appendix B), and (v) angiogenesis. Adipocytes adapt to a dynamic niche as well as a microenvironment invaded by foreign cells in its stromal vascular fraction (SVF). However, the adaptation potential is limited in time and space, whereby the inability of WAT to adequately expand to meet the energy storage demands results in AT dysfunction, ectopic lipid deposition, and insulin resistance [16]. Whereas ECM is recognized as a critical component of the AT microenvironment [9,10], the resulting altered tissue mechanics in obesity need to be better studied, since this phenomenon can be a potential driver of AT dysfunction. This also requires deeper understanding of adipocyte mechanotransduction. Indeed, during AT expansion, tightly controlled tissue homeostasis is lost, and dysregulation of the ECM may alter the mechano-reciprocity between adipocytes, SVF cells and the ECM to create an iatrogenic feedback loop.
In this review, we discuss the emerging role of mechanosensitive pathways in regulating AT function and the potential repercussions of altered adipocyte mechanical microenvironment in the initiation of AT-related diseases.

2. Major Alterations of Expanded Adipose Tissue

2.1. Hypertrophy vs. Hyperplasia

In adulthood, AT size is fairly constant under homeostatic conditions, but it is highly sensitive to dietary challenges (e.g., high caloric diet). In a given fat depot, the number of adipocytes is determined early in life and is mostly stable through adulthood [17,18,19,20]. Since mature adipocytes are post-mitotic cells, new cells are derived from adipose stem cells (ASC) located in the SVF (Appendix B) [10,17,21]. In young adult mice, the rate of adipogenesis has been estimated to 10–15% per month [19,20], and retrospective human studies have estimated the turnover rate to 10% per year [18]. However, the turnover rate of adipocytes gradually declines with age due to defective regenerative ability of the ASCs and the senescence of the microenvironment [20,22].
In response to caloric excess, mature adipocytes possess impressive hypertrophic potential, being able to increase in size to several hundreds of micrometers in diameter [23]. Moreover, fat mass might also expand by hyperplasia [19,20,24,25,26]. For instance, it has been shown that visceral WAT expansion in males occurs through both adipocyte hyperplasia and hypertrophy in both mice [24,27] and humans [28]. In male mice, subcutaneous WAT does not exhibit relevant hyperplasia in response to obesogenic stimuli [24,25,27,29] while in female mice, high-fat diet (HFD) induces hyperplasia in both visceral and subcutaneous WAT [27]. Thus, hyperplasia is induced in a sex hormone-dependent manner in WAT depots [27]. In 1956, Jean Vague was the first to show the importance of regional WAT distribution in relation to various metabolic diseases [30]. He observed that individuals with obesity who preferentially expand visceral WAT (i.e., “apple-shaped obesity”) are at greater risk for metabolic disorders than those who accumulate subcutaneous WAT (i.e., “pear-shaped obesity”) [30]. Recent studies suggest that hypertrophy in the subcutaneous WAT is linked to insulin resistance while visceral WAT hypertrophy also correlates with dyslipidemia [13]. Moreover, while adipocyte number in subcutaneous WAT protects against metabolic complications, an increase in the number of fat cells in visceral depots does not associate with metabolic state [31].

2.2. Inflammation and Fibrosis

In subjects with obesity, WAT expansion is characterized by local inflammation followed by ECM deposition. AT secretome is usually characterized by lower adiponectin and increased levels of leptin, free fatty acids, tumor necrosis factor α (TNFα), interleukin-6 (IL-6), IL-8, monocyte chemoattractant protein-1 (MCP1), and acute-phase serum amyloid A proteins [7,32]. Some inflammatory mediators are produced by hypertrophic adipocytes [33] but tissue-resident immune cells are also major contributors to the AT secretome in obesity [8,34]. Immune cells indeed dramatically increase in abundance during WAT expansion [35,36]. AT-derived immune cells display distinct phenotypes from their circulating counterparts [37,38]. AT is indeed enriched with an important diversity of immune cells including macrophages forming “crown-like” structures around hypertrophic adipocytes [39,40], regulatory T and B cells [41,42,43], lymphocytes T gamma delta and natural killer T (NKT) cells that drive thermogenesis [44,45,46], memory T cells [47], cytotoxic innate lymphoid cells [48], dendritic cells [39,49], neutrophils [50], as well as, ILC2 cells [51,52,53,54,55]. This network of immune cells is tailored to support AT homeostasis, enabling it to adapt to environmental and nutritional factors [8,56,57,58]. In obesity, the immune system of AT is dysregulated, driving sterile inflammation with altered remodeling that can progress towards ECM deposition and fibrosis accumulation [59,60] (Appendix C). Moreover, basal membrane components such as collagen IV, nidogen and proteoglycans are increased in obese AT and associate with insulin-resistance markers in subjects with obesity [61,62].
Progression towards pathological AT remodeling is often accompanied by decreased ECM flexibility possibly due to pericellular collagen deposition and enhanced ECM crosslinking by lysyl oxidase (LOX) enzymes [63]. This promotes the formation of collagen bundles that stiffen the tissue and constrain AT expansion [64]. The production of ECM proteins is also regulated by a variety of AT cells including adipocyte progenitors, adipocytes, fibroblasts, and myofibroblasts [9,65]. AT fibrosis is an aggravating factor for metabolic condition [10,66,67]. For instance, Col6a1 knockout mouse model [66] that results in a complete loss of collagen VI fibers within the ECM in AT is accompanied by uncontrolled enlargement of the adipocytes in obesogenic condition. Although massively obese, these transgenic mice exhibit an improved metabolic and inflammatory profile [66]. In subjects with obesity, collagen VI expression levels are linked to WAT remodeling and insulin resistance [68,69]. In accordance, it was shown that subjects with obesity and insulin-resistance have increased subcutaneous and omental AT fibrosis, compared with insulin-sensitive obese subjects [61,70]. Notably, a histological study reported that pericellular fibrosis in omental AT was not associated with significant adipocyte hypertrophy [61]. It was hypothesized that pericellular fibrosis in omental AT restricts adipocyte hypertrophy which may, in turn, preserve adipocyte function and systemic metabolism [71]. However, omental mature adipocytes from insulin-resistant obese individuals display features of enhanced inflammation, oxidative stress and endoplasmic reticulum stress, compared with normoglycemia obese individuals [72].
Altogether, the AT microenvironment is a complex system composed of a myriad of cells including adipocytes, endothelial and immune cells, ASC, and fibroblasts. These cells interact with their surroundings via both mono- and heterotypic cell–cell, or cell–ECM interaction [73]. AT cells receive and integrate information from their microenvironment impacting on their behavior, including the control of expansion/shrinking and remodeling upon nutritional or environmental stimuli.

3. Cell–Cell Cross-Talk in Adipose Tissue

Cells of multicellular organisms need to communicate with each other to maintain tissue function and homeostasis. Intercellular crosstalk often involves direct physical cell–cell contact between adjacent cells via specific cell surface receptors and/or via soluble factors such as cytokines, chemokines, growth factors, neurotransmitters, and extracellular vesicles.

3.1. Connexins and Neuronal Fibers

Direct cell–cell contacts between adipocytes are mediated by connexin 43 gap junctions [74,75,76]. A gap junction is composed of two hemichannels (e.g., connexons), each of which is a hexamer of connexin subunits forming an aqueous channel with a molecular weight cut-off of approximately 1 kDa [77,78]. The connexin gap is large enough to enable the passage of ions, cAMP, and other small molecules and metabolites [79]. Scherer and colleagues demonstrated that connexin 43 facilitates the propagation of sympathetic neuronal signals across adipocytes, to enable beiging [76]. Sympathetic nerve fibers release noradrenaline implicated in mediating adipocyte lipolysis and thermogenesis [80]. However, as WAT is not well innervated compared with brown adipose tissue (BAT) [81], the gap junctions are critical to maximize the physical interactions between white adipocytes allowing an effective propagation of neuronal signals in response to stimuli. Moreover, sympathetic fibers show close apposition to >90% of adipocytes [82] and can form neuroadipocyte junctions in mouse WAT [83]. Surgical or pharmacological destruction of sympathetic fibers inhibits the leptin-stimulated lipolytic response of WAT [83]. However, the mechanism of neural regulation in AT remains elusive. Adipocytes produce neurotrophic factors including nerve growth factor [84], S-100 protein β-chain [85,86] and neuregulin 4 [87] which control proliferation, survival, migration and differentiation of neurons. Cold exposure in mice or fasting increase the expression of these neurotrophic factors that promote densification of the sympathetic axonal tree in AT [81,82,88,89]. In obese condition, AT sympathetic innervation is significantly reduced as shown in Ob/Ob compared with lean mice and subcutaneous WAT from subjects with obesity [90]. Reduced sympathetic innervation is associated with attenuated catecholamine-induced lipolysis, promoting WAT expansion and reducing the ability to withstand starvation and thermogenesis [91]. Immunohistochemical analysis of selective neuron markers, such as tyrosine hydroxylase, revealed sympathetic nerve fibers in close anatomical association with the vasculature in AT [80,92].

3.2. Vasculature

The vasculature supplies the AT microenvironment with the necessary oxygen, nutrients, hormones, and growth factors, and removes metabolic waste products. Vasculature is known to play an important role in the regulation of triglyceride storage. Lipoprotein lipase (LPL), which is mainly synthesized by the adipocyte, is an extracellular enzyme that acts on the endothelial surface and hydrolyzes the triglycerides of chylomicrons to fatty acids [93,94], that are then up taken by FAT/CD36 (fatty acid translocase), the principal adipocyte fatty acid transporter [95,96]. The vasculature contributes to the maintenance of adipocyte turnover [97]. Several populations of ASCs have been identified in the mural compartment of vascular structures [97,98,99,100]. Furthermore, a study using irradiation followed by bone marrow transplantation suggests that over 85% of macrophages are recruited from blood monocytes into AT in a CSF1-dependent manner [35]. Interestingly, sympathetic activation promotes angiogenesis via upregulation of VEGF-A expression in rat BAT [101,102]. Adipocytes secrete VEGF which acts on the VEGFR2 receptor expressed on the endothelium to stimulate angiogenesis [103]. Induction of VEGF expression in adipocytes increases adipose vasculature, improves AT function and reduces insulin resistance and glucose intolerance in HFD fed mice [104,105]. Whether these results obtained in mouse models can be translated to human is still under debate [106].

3.3. Immune Cells

Resident AT immune cells are necessary for adipocyte biological function. In the lean condition, the immune cells are mostly anti-inflammatory, comprising alternatively activated (M2) macrophages which maintain their polarization through type 2 innate lymphoid cells (ILC2), T regulatory cells (Tregs), invariant natural killer T cells (iNKT cells), natural killer cells, and eosinophils [106]. Macrophages are responsible for many housekeeping processes, such as the removal of dead cells, ECM remodeling, regulation of catecholamine availability, modulation of angiogenesis and differentiation of adipocyte precursors [83,107,108]. At an early stage, the inflammatory reactions are necessary for AT expansion. However, in obesity, the adipose immune system shifts toward a chronic pro-inflammatory state and, at an early stage, the proinflammatory reaction is necessary for AT expansion. The maintenance of chronic inflammation though leads to major perturbations of AT homeostasis [8]. The triggers of inflammation are still to be better understood whereas hypertrophic adipocytes release proinflammatory factors such as TNFα, IL6 and CCL2, that promote AT macrophage infiltration [35,36,109]. The rise of the number of macrophages is not only due to the recruitment and differentiation of CCR2-dependent blood monocytes. Aouadi and collaborators demonstrated that AT macrophages (ATMs) can locally proliferate in at least a partially CCL2-dependent manner in the visceral AT of mice [110]. Functionally, obesity reduces the endocytic capability of vascular-associated ATMs [111]. Furthermore, ATMs undergo “unconventional” metabolic activation [112] rather than a classically activated (M1) or alternatively activated (M2) macrophage polarization [113]. Studies using single-cell RNA sequencing (scRNA-seq) analyses revealed that the transcriptomic profiles of ATMs are more heterogeneous in obese AT than anticipated [39,40]. A population of lipid-associated macrophages (LAMs) that accumulate at high numbers in obesity has been recently described. LAMs are characterized by the expression of the lipid sensor TREM2 that drives gene expression programs involved in phagocytosis, lipid catabolism, and energy metabolism [40]. Impairment of this process in obese mice with a global deficiency of Trem2 inhibited the downstream molecular LAM program, leading to adipocyte hypertrophy associated with worsened metabolic condition [40]. Metabolic adaptation is a key component of macrophage plasticity and polarization [114]. Interestingly, it was demonstrated that murine macrophages acquire mitochondria from adipocytes in vivo [115]. This adipocyte-to-macrophage transfer of mitochondria defined a distinct tissue macrophage subpopulation found to be reduced in obesity. Interestingly, the inhibition of this transfer resulted in decreased energy expenditure and increased weight gain and glucose intolerance [115]. Whereas the adipocyte-to-macrophage transfer of mitochondria illustrates again the importance of cell crosstalk in AT, the mechanisms involved are still unclear. Previous studies have identified extracellular vesicles as a potential vehicle for intercellular transfer of mitochondria [116,117].

4. Mechanical Regulation of Adipocyte

4.1. Physical Constraints to and from Adipocytes

Few studies have explored the mechanical properties of adipose cells and how this may impact on their functions. Adipocytes are characterized by a unique large spherical lipid droplet which is surrounded by a thin rim of cytoplasm containing a crescent shaped nucleus and sparse organelles, mainly elongated mitochondria [118,119]. Upon intense lipolysis, smooth endoplasmic reticulum is visualized in tight contact with the lipid droplets [118,119]. During caloric excess promoting adipogenesis, the accretion of lipid droplet requires major rearrangement of the cytoskeleton [120,121] also involving changes in the mechanical properties of white adipocytes [73]. For example, adipogenesis is associated with a progressive cell stiffness increasing from 300–900 Pa in 3T3-L1 preadipocytes to 2 kPa in differentiated 3T3-L1 cells [122]. The stiffness of the lipid droplets is 2.5 to 8.3 times greater than the effective stiffness of the surrounding cytoplasm [123]. This suggests that mechanical properties of adipocytes vary according to lipid droplet size.
Moreover, it can be estimated that physical tension applied from the interior to the adipocyte membrane is generated from the lipid droplet [124]. Membrane caveolae are invaginated structures specialized in the response to mechanical tension at the cell surface membrane [125] and are particularly abundant in mature adipocytes [126,127]. It has been shown that these structures are involved in buffering lipid storage of fat cells [128]. Of interest, the importance of caveolae in the adipose cell biology is illustrated by the severe lipoatrophic syndrome that develops in patients with caveolin gene mutations [129].
Thus, in obesity, adipocyte hypertrophy is associated with increased AT stiffness [130,131,132]. Stiffness of hypertrophic adipocytes may impose mechanical constraints on the surrounding microenvironment perturbing local cell biology and eventually promoting ECM remodeling (Figure 1). On the other hand, AT fibrosis limits adipocyte hypertrophy as a biophysical constraint; e.g., adipocytes trapped in fibrotic bundles are smaller in size compared with the other adipocytes outside the bundles from the same tissue [133]. Interestingly, in a diet-induced obesity rat model, surgery-induced weight loss was related to an improvement of the biomechanical properties of visceral AT compared with “sham surgery” in obese rats. These changes in biomechanical properties were associated with enhancement of angiogenesis, reduction in adipocyte size and downregulation of the expression of ECM components [134]. Furthermore, modelling the physical constraints applied to adipocytes in ex vivo systems showed that the mechanical compression leads to major adipocyte dysfunction characterized by increased production of inflammatory adipokines as well as dysregulated lipolysis and perturbed insulin sensitivity in mature adipocytes [135].

4.2. Environmental Stiffness and Regulation of Adipogenesis

From a cellular perspective, a major source of knowledge comes from cell culture models involving fibroblast-like cells that differentiate to form adipocytes in response to hormonal cocktails. Observations indicate a high degree of intracellular and extracellular structural remodeling during adipogenesis [136]. Along with the biochemical properties of the progenitors’ microenvironment in vivo [137], mechanical cues impact on both stem cell proliferation and differentiation [138]. The transmission of extracellular mechanical force to the inner part of the cell, the nucleus, is achieved by a dynamic regulation of membrane and cytoskeleton integrity and tension [139]. Adipogenesis is achieved by controlling substrate mechanics, nanotopography (referring to the specific surface features generated at the nanoscale) and cell spreading [140,141]. For example, in two-dimensional (2D) cell culture models, adipogenesis requires cell confluence (e.g., cells are physically in contact with each other) [142,143]. Upon confluence, contact inhibition will arrest preadipocyte growth. In vitro adipogenesis is induced by hormonal stimulation, where preadipocytes re-enter the cell cycle, arrest proliferation and undergo terminal adipocyte differentiation [144,145]. Furthermore, three-dimensional (3D) cell culture within a methylcellulose gel promotes adipogenesis by constraining the shape of the cell similar to what is observed in confluent cells [144]. Mesenchymal stem cells plated on a small surface leading to a rounded morphology tend to undergo adipogenesis, whereas single cells grown on larger surfaces allowing cell spreading tend to undergo osteogenesis [136,141,146,147,148]. As cellular shape is influenced by mechanical properties and forces exerted on the cell, it is thus not surprising that mesenchymal stem cell fate is determined by substrate mechanical cues [138]. In mesenchymal stem cells, adipogenesis is favored by a soft microenvironment [140,149]. Thus, culturing ASCs on gels that mimic the native stiffness of AT (2 kPa) encourages adipogenesis, even without exogenous adipogenic hormonal stimuli [150]. When human mesenchymal stem cells are seeded on polyacrylamide gels with low stiffness, they are more prone to become adipocytes than cells grown on stiffer gels [150]. When microenvironment stiffness increases, the ASC spread out and lose their rounded morphology. Moreover, cyclic stretching or vibration impedes adipocyte differentiation, but static stretching promotes it [122,123,151]. Altogether, cell spreading and mechanical cues can constrain adipogenesis by modifying cell shape [136].

4.3. Influence of Extracellular Matrix and Osmotic Microenvironment on Adipose Cells

ECM is a major determinant of AT mechanical properties. Knockout of genes that encode collagens [66] or collagenases induces modulation of pericellular collagen rigidity and affects adipogenesis [152]. Dani and collaborators demonstrated that the inhibition of collagen synthesis with a competitive inhibitor of prolyl hydroxylase (Ethyl 3,4-dihydroxybenzoate) represses the differentiation of preadipocytes into adipocytes and impairs ECM development [153]. In vitro, type I collagen has been reported to favor adipogenesis [154], and denatured, but not native, type IV collagen, promotes adipogenesis in human mesenchymal cells [155].
Microenvironment stiffness and composition are thus under the control of the surrounding ECM that also involves members of the matrix metalloproteinase (MMP) family, and their functions are coordinated by inhibitors of metalloproteinases called TIMPs (for tissue inhibitors of metalloproteinases). Metalloproteases (MMP), where their expression in AT and serum is increased in obesity [156,157,158], exert effects on adipogenesis. For example, MMP14 has a major effect on adipogenesis by altering collagen stiffness in a 3D setting in vitro [152]. Mmp14+/− mice challenged with HFD have defects in AT expansion due to impaired type I collagen cleavage, and develop metabolic alterations [159].
Another ECM protein, fibronectin, inhibits the differentiation of 3T3-F442A preadipocytes into adipocytes [160]. Similarly, the presence of secreted-protein-acidic-and-rich-in-cysteine inhibits adipogenesis by stimulating the deposition of fibronectin [161]. The anti-adipogenic activity of fibronectin can be reversed by keeping cells in a rounded configuration or by exposing cells to cytochalasin D, which disrupts the actin cytoskeleton [160].
Blocking the interaction of integrin alpha 6 with laminin by an anti-integrin alpha 6 antibody, enhances differentiation of adipocytes by repressing RhoA (characterized by anti-adipogenic effects) [162]. Proper integrin signaling is also required for de novo beige fat biogenesis following cold exposure. The integrins β5 and β1 interact with CD81, and are involved in irisin-mediated FAK signaling in beige ASCs [163]. Knockout or antibody-based blockage of either integrins (β5 or β1) or CD81 abolishes the effect of irisin-induced FAK phosphorylation in beige ASCs [163]. Noteworthy, the number of CD81 and ASCs negatively correlates with body weight, insulin resistance, glucose intolerance, and WAT inflammation in experimental models of diet-induced obesity [163].
Another important element of the cellular microenvironment is osmotic pressure which can influence cellular membrane tension, and mechanotransduction. All cells may be exposed to osmotic swelling or shrinkage but in the case of adipocytes, the effect of osmotic pressure might be more pronounced due to the relatively small water compartment [164,165]. Interestingly, the development of the mechanisms controlling osmotic pressure is an important part of the adipocyte phenotype. As shown by Eduardsen et al., osmotic swelling promotes nuclear ERK1/2 activity and decreases IRS phosphorylation in response to insulin [166]. Alternatively, osmotic shrinkage increases phosphorylation of IRS and FAK, lowering the ERK signaling at the same time [166,167,168].

4.4. Effect of Mechanical Cues on Brown Cells

Brown/beige adipocytes differ from white adipocytes in their morphology and their functions. Thermogenic adipocytes need to maintain an extensive and dynamic cytoskeleton to support and organize multilocular lipid droplets and a dense network of mitochondria. This also implies significant differences in mechanical properties. BAT is stiffer than WAT, and BAT stiffness increases when mice are exposed to lower temperature (4 °C). By using atomic force microscopy, Tharp et al. observed that in vitro treatment of brown adipocytes with β-adrenergic agonist (isoproterenol) induces a contractile-like response which stiffens the cell cortex [169]. The contractile-like response in BAT appears to strongly mimic that of cardiomyocytes which can be explained by the common embryonic origin (Myf5+ cells) shared by brown adipocyte and muscle cells [170]. As such, muscle-specific type II myosin heavy chains (MyH) are expressed in brown fat cells conferring a dynamic role of actinomyosin mechanics in thermogenic cells [169]. The repression of actinomyosin-mediated tension by using type II myosin inhibitors (blebbistatin and 2,3-butanedione monoxime) reduces cytoplasmic stiffness and leads to a specific loss of UCP1 expression and oxidative capacity. The authors showed that actinomyosin contractility regulates the thermogenic capacity of brown adipocytes via the mechanosensitive transcriptional co-activators YAP and TAZ (see below). Indeed, YAP overexpression increases UCP1 levels. Accordingly, mice with brown adipocyte-specific (Ucp1-cre) knockout of YAP/TAZ resulted in whitening of BAT, increased fat mass, decreased energy expenditure, and showed hyperinsulinemia and glucose intolerance [169].

5. Actors of the Mechanotransduction: From Tension to Adipose Cell Function

Cell shape is linked to extracellular mechanical properties such as ECM stiffness, stretching or shear stresses exerted by flowing liquids and attachment to other cells (Figure 1 and Appendix D). In the case of fibrotic AT, the changes of ECM stiffness are, therefore, critical in adipocyte mechano-responses. In a model of type II diabetes mellitus, by mixing human decellularized AT ECM and adipocytes, it has been shown that the specific dysfunctional fibrotic cues are part of the ECM and not a property of cells [135]. Another research supported this finding by demonstrating that mechanical changes of the surrounding ECM are transmitted by the cytoskeletal actomyosin stress fibers in adipocytes that were cultured on stiffness-controlled hydrogels [171].

5.1. Cytoskeleton

The cytoskeleton is a dynamic structure essential to maintain cell shape, which provides structural support that functions as a mechano-sensor in combination with focal adhesion proteins [172]. During adipocyte differentiation, ASC undergo morphological transformation to allow the formation and growth of nascent lipid droplets [173] where vimentin transcription increases to proportionally expand the vimentin cage around each droplet [174], while tubulin and actin expression decreases [121]. These cytoskeletal rearrangements are prerequisites for terminal differentiation [121,160,175,176,177]. The inhibition of actin filament polymerization or actomyosin contraction promotes a rounded morphology of adipocytes and subsequently promotes adipogenesis [141,178].
During adipogenesis, changes in cell shape involve disruption of filamentous (F) actin via downregulation of RHO GTPase–RHO-associated kinase (ROCK) signaling and the actin-regulatory proteins (Arp2/3 complex, cofilin, profilin) [120,141,179]. The inactive RHO, RHO GDP, is the predominant RHO species in confluent or rounded human MSCs and promotes adipogenesis, whereas ectopic addition of RHO GTP inhibits it [141]. However, in stiff environments, focal adhesion assembly is promoted, RHO GTP activates ROCK, which, in turn, stimulates F-actin stress fibers formation and translocation of YAP/TAZ to the nucleus, inhibiting adipogenesis [141,180] (Figure 2).
In cultured mature adipocytes, lipid droplet growth is associated with increased Rho-kinase activity [181]. Accordingly, mechanical stretching (e.g., mechanical event induced by lipid droplet enlargement in the cell) on mature adipocytes increases Rho-kinase activity and stress fiber formation, as observed in adipocytes from HFD-fed mice [181]. Thus, adipocyte hypertrophy following 2 weeks of HFD-feeding in mice was associated with a drastic increase in F-actin, increased Rho-kinase activity, and changed expression of actin-regulating proteins, favoring polymerization [182]. Conversely, another study reported that lipid droplet enlargement in adipocytes downregulates cortical F-actin formation [183]. Interestingly, the authors proposed that this downregulation of F-actin organization would lead to decreased insulin-dependent GLUT4 trafficking to the plasma membrane, which governs the insulin-dependent glucose utilization [183].

5.2. Mechano-Induced Cell Metabolism Drives Cell Functions

Recent advances have revealed that the activation of transcriptional programs that determine cell functions can be modulated by the biochemical environment shaped by cell intrinsic metabolic activity exhibited by a cell. Indeed, metabolism impacts intracellular cell signaling pathways by influencing protein post-translational modifications (PTMs) of signaling mediators. Moreover, these PTMs impart a broad influence on transcriptional activation by modulating chromatin state via DNA (hydroxy)methylation, histone or transcription factors PTMs [184,185,186,187,188]. Thus, altered metabolism has been proposed to play an important role in AT dysfunction altering, in turn, whole body energy homeostasis [5,189,190,191].
Actually, cell metabolism is tightly regulated by mechanical cues. Recent work revealed that mechanical forces impact cell metabolism via cytoskeletal reorganization [192,193]. Park et al. found that increasing substrate stiffness correlates with the trapping of E3 ubiquitin ligase tripartite motif (TRIM)-containing protein 21 (TRIM21) within actin stress fibers, thus reducing access to its substrates such as phosphofructokinase (PFK) [194]. On a soft ECM that permits relaxation of the actomyosin cytoskeleton, TRIM21 is released, PFK is degraded thus leading to reduced glycolysis [194] (Figure 3). Accordingly, matrix stiffness increases ATP:ADP ratio and glucose uptake by stimulating glucose transporter 1 (GLUT1) [195,196]. GLUT1 has been reported to be retained at the site of force transmission by ankyrin G which is critical for allowing cells under tension to tune glucose uptake and to fuel the reinforcement of the actin cytoskeleton [196,197]. Moreover, YAP and TAZ that relay stiffness signals, stimulate glucose metabolism by enhancing GLUT1 expression [197]. Bertero et al. also reported that glutamine catabolism is dependent on YAP/TAZ that bind to the promoter of GLS (glutaminase) to promote its transcription [198,199]. Thus mechano-activation of glutaminolysis and glycolysis sustains the metabolic needs of mechano-activated cells and, in turn, alters cell phenotype. In order to elucidate the outmost importance of the mechanics of the environment on adipocyte behavior and the impact of fibrosis on their identity, traditional in vitro models have to be replaced by higher-fidelity microphysiological systems that can mimic physiopathological conditions, as reviewed recently in [200].

5.3. The Nucleus

The nucleus is the largest and stiffest cellular organelle [172]. In adipocytes, the nucleus is compressed against the plasma membrane by large lipid droplets. It can be divided structurally and functionally into the nuclear interior, which houses chromatin, and the surrounding nuclear envelope (Figure 1). The nuclear envelope is composed of two concentric membranes, e.g., the inner and outer nuclear membranes, an underlying nuclear lamina, and nuclear pore complexes that control entry of large molecules into the nuclear interior [172]. The nuclear lamina is a filamentous protein network lining the nucleoplasmic surface of the inner nuclear membrane. It is composed of A-type and B-type lamins, and lamin binding proteins [172,201]. Although B-type lamins are mostly restricted to the nuclear lamina, A-type lamins (encoded by LMNA) are found both at the nuclear periphery and in the nuclear interior interacting with chromatin [202,203,204,205]. When A-type lamin is depleted, cellular elasticity and viscosity of the cytoplasm decrease and the nucleus becomes softer and more deformable [206]. On the other hand, seeding adipocytes on stiff surfaces and subsequent spreading of cells is associated with upregulation of lamin A proteins [207].
During cell differentiation, the nuclear architecture is reorganized [208,209]. For example, in preadipocytes, A-type lamins move from intranuclear structures to the nuclear rim as adipogenesis proceeds [209]. The important role of lamin is illustrated by mutations throughout the LMNA gene that cause various forms of laminopathies, including partial lipodystrophies (Appendix E) [210].
Lamins play an important role in physically connecting the nucleus to the cytoskeleton. Such connections are mediated by the members of the so-called linker of nucleoskeleton and cytoskeleton (LINC) complex [211,212]. These connections mediate intracellular force transmission, cell migration and cell polarization [213]. Through these connections, when mechanical force is applied on the plasma membrane, alterations of the nuclear morphology are induced [214] and translated into chromatin remodeling and epigenetic modifications [215,216]. Interestingly, by using photoconvertible, photo-softening [215] and photo-stiffening [217] hydrogels, Anseth and collaborators demonstrated that human mesenchymal stem cells grown on stiff matrices undergo chromatin remodeling that is characterized by increased histone acetylation and reduced chromatin condensation. Then, an interesting model to consider would be that mechanical cues from the microenvironment are transmitted through integrins, the cytoskeleton, the LINC complex, and lamins to the nucleus, where the mechanical cues are translated into chromatin remodeling and epigenetic modifications [218,219]. Reports have shown how mechanical cues can regulate epigenetic status, through the influence of actin cytoskeleton [216,220,221]. It has been shown that alteration in cytoskeleton tension by compression/stretch in the direction perpendicular to cell alignment induces nuclear shape change, a decrease in histone deacetylase (HDAC) activity, and an increase in histone acetylation [222]. In addition, cells in an elongated shape exhibit elongation of their nucleus and higher levels of nuclear histone H3 acetylation, compared with cells in a circular shape [223]. Although not in adipose cells, this discovery demonstrates the impact of mechanical cues to influence the epigenetic state [223]. The mechanism behind this “biophysical” alteration of histone acetylation relies on altered HDAC activity, possibly because HDAC is sequestered in the cytosol by actin cytoskeleton [224]. Thus, the dynamic reorganization of the nuclear lamina and the cytoskeleton, induced by mechanical cues, may impact on histone acetylation that plays a key role in controlling adipogenesis and adipocyte function [225].

5.4. Mechano-Transduction Mediated by YAP/TAZ

Yes-associated protein (YAP) and PDZ-binding motif (TAZ) are paralogous proteins that act as transcriptional co-regulators [226,227]. The phosphorylation of YAP/TAZ by LATS1/2 (Hippo signaling) leads to cytoplasmic retention and degradation of YAP/TAZ [228]. YAP/TAZ are robust mechano-sensors that mediate cell responses to cell spreading [229,230] and stretching [231], fluid flow-induced shear [232], and to microenvironment stiffness [180,226]. However, the mechanisms by which mechanical cues regulate YAP/TAZ nuclear shuttling are still unclear.
During adipogenesis, activation of LATS2 and concomitant reduction in YAP/TAZ nuclear activity were described [233]. The cytoplasmic retention of TAZ acts on Wnt signaling to prevent β-catenin translocation to the nucleus, which stimulates the pro-proliferator and anti-adipogenic TCF/LEF transcriptional activity [233]. Interestingly, osteoblast-specific deletion of TAZ in mice reduces cell proliferation and osteogenesis, and promotes adipocyte formation, resulting in a trabecular bone loss. Conversely, YAP is selectively expressed in osteoblast-lineage cells where it interacts with β-catenin, increasing β-catenin stability in the nucleus [234]. Interestingly, YAP repression by phosphorylation appears to be mediated by cell spreading and is sufficient to prompt adipogenesis, regardless of a permissive environment [149]. However, ASC-specific overexpression of YAP appears to increase adipogenesis and obesity in mice [235]. Nevertheless, this study reported that Yap overexpression in ASC induced a negative feedback mechanism on the Hippo signaling pathway, leading to suppression of TAZ activity. The latter enhances PPARγ activation and increases adipogenesis [235]. It was also proposed that increased cytoskeleton formation, by overexpression of smooth muscle actin (SMA), induces cell spreading, increases YAP activity and inhibits adipogenesis in human mesenchymal stem cells [236]. Together, these studies suggest that the YAP/TAZ pathway mediates adipogenic regulation induced by microenvironment stiffness and cell spreading.
As adipocyte hypertrophy increases local mechanical stress, it was observed that YAP and TAZ were both activated in mouse and human white adipocytes during obesity [237]. In agreement with the anti-adipogenic effect of YAP/TAZ activity, Olefsky and collaborators have shown that TAZ acts as a PPARγ co-repressor in adipocytes. In obese mice, adipocyte-specific deletion of TAZ increased PPARγ activity and resulted in increased systemic insulin sensitivity and glucose tolerance with an improved AT phenotype [238]. However, compelling and contradictory data demonstrated that the loss of both YAP and TAZ specifically in adipocytes resulted in lipodystrophy, including massive adipocyte death, increased adipogenesis, macrophage infiltration and improved glucose tolerance in HFD fed mice [237]. There are several hypotheses to reconcile these paradoxical results. One possibility is that YAP and TAZ work redundantly to maintain adipocyte survival so that the loss of both simultaneously is fatal for adipocytes. The second possibility is that TAZ has an activity not shared with YAP. TAZ, where its role is better described, represses PPARγ activity in white AT that contributes to insulin resistance, abnormal AT remodeling and local inflammation [238,239,240]. However, the role of YAP in regulating adipocyte differentiation or adipocyte phenotype appears uncertain. It could be involved in the negative feedback mechanism via the Hippo signaling pathway or in the control of cell death [237].

5.5. Membrane

The conversion of forces can also be transduced through electrical signals via the opening of mechanosensitive ion channels present in the cellular membranes [241]. In addition to the above-mentioned regulation of caveolae assembly/disassembly, increased adipocyte volume and subsequent increase in plasma membrane tension may result in the activation of these ion channels. Recently, evidence was provided that two different channels, Piezo1 and SWELL1 (a volume-regulated anion channel (VRAC)), are activated in the context of adipocyte hypertrophy [242,243]. It has been shown that SWELL1 activation in response to mature adipocyte volumetric expansion promotes adipocyte expansion, energy storage, and enhances insulin signaling during increased caloric intake [242]. Piezo1 in mature adipocytes is rather a critical mediator of obesity-induced adipocyte hyperplasia. Indeed, Offermanns and collaborators have shown that the opening of Piezo1 in hypertrophic adipocytes induces the release of the pro-adipogenic fibroblast growth factor 1 (FGF1) which promotes, in turn, ASC differentiation [243]. These data demonstrated that the mechanosensitive ion channels are cell-autonomous sensors of adipocyte volume that regulate AT growth, insulin sensitivity and adipogenesis (Figure 4).

6. Conclusions

The current obesity epidemic needs a great deal of attention on the understanding of adipose biology. Mechanical properties are spatially and temporally regulated to preserve the homoeostasis of AT. Indeed, the adipose microenvironment is a mechanically complex niche, in which a number of inputs regulate AT function. However, the influence of mechanical cues on adipose function, and vice versa, remain underappreciated. The interplay between adipose resident cells and ECM determines the maintenance of a mechanical environment that supports tissue architecture and function. Aberrations in adipose microenvironment play a profound role in the development and progression of obesity. Investigations have partially revealed how adipose cells respond to mechanical stress during AT expansion. There is also evidence that a niche can modulate preadipocyte and adipocyte fate by defining the mechanical properties. We believe that fibrosis and adipocyte hypertrophy ultimately disrupt mechanical homeostasis, altering mechanotransduction signaling through cell–cell, cell–matrix–mediated transcriptional/epigenetic and PTMs mechanisms, leading to progressive ECM deposition and AT dysfunction. Thus, it will be important to dissect the crucial molecular events that connect mechanical cues with defects in AT function. Ultimately, translating these insights into clinical and therapeutic interventions may enable to delay the cycle of obesity and its related complications.

Author Contributions

Conceptualization, S.L., M.L., M.H. and K.C; Literature Review, S.L., M.L., K.D., I.D, S.M., M.H. and K.C.; Figures and Tables, S.L.; Writing—Original Draft Preparation, S.L.; Writing—Review & Editing, S.L., M.L., K.D., I.D., S.M., M.H. and K.C. All authors have read and agreed to the published version of the manuscript.

Funding

This review was founded by the Fondation pour la Recherche Medicale (FRM team 2020–2023) [FDT201904008276, FDT202106012793]. French National Agency of Research (ANR-Captor), Clinical Research program (PHRC), Société Francophone du diabète (SFD), Novo Nordisk foundation, Paris Region Ile de France (DIM).

Institutional Review Board Statement

Not applicable.

Informed Consent Statement

Not applicable.

Data Availability Statement

Not applicable.

Conflicts of Interest

The authors declare no conflict of interest.

Appendix A. Central Role of Adipose Tissue in Regulating Whole-Body Energy Homeostasis

There are at least two classes of fat cells: white and brown. WAT is widely dispersed throughout the body and is organized into anatomically distinct “depots” with 9–18% and 14–28% of the body weight being fat depot, respectively, in lean males and females [119,244]. Major depots reside in the subcutaneous region of the upper (deep and superficial abdominal) and lower body (gluteal–femoral) parts, and in the visceral region (omental, mesenteric, mediastinal and epicardial). Subcutaneous WAT is located under the skin where it acts as a thermal insulator to prevent heat loss, a mechanical cushion for protection against trauma and a barrier against dermal infection [245]. Visceral WAT is wrapped around vital organs such as the heart, pancreas and kidneys. White AT is generally characterized by adipocytes with a single lipid droplet for efficient energy storage, while brown and beige adipocytes (defined as brown-like adipocytes in white fat depots) are characterized by multiple lipid droplets and high mitochondrial content expressing UCP1 for uncoupled oxidative phosphorylation and non-shivering thermogenesis [246]. BAT is present predominantly in rodents, hibernating animals and infant humans where it is mainly located in the interscapular region [246].

Appendix B. In Vivo Development of Fat Depots

Adipogenesis occurs through commitment from multipotent stem cells to preadipocytes, and terminal differentiation from preadipocytes to mature adipocytes with triglyceride-filled lipid droplets. In mammals, adipogenesis starts in late gestation [17,97,247,248] and its rate rapidly rises in response to nutrient availability, leading to a marked postnatal expansion of fat depots [249,250,251]. In mice, adipogenesis in subcutaneous depots starts between embryonic days E13.5 and E18.5 [24,98,248,252] and perigonadal AT depot develops between postnatal day 3 (P3) and the second week of life. The mesenteric AT completes its organogenesis between the second and third weeks of life [17,225,248]. In humans, adipogenesis starts around the 14th–17th week of gestation as a cluster of fat lobules, first in the head and neck, then the trunk, and later in the limbs [253]. By the 28th week of gestation, fat depots are organized [253]. After birth, both the number and the size of adipocytes increase. In a cross-sectional study performed on children, cell size increases to an adult level from the age of 6 months to one year, then reduces between one and two years [250]. The number of adipocytes increases once again at adolescence [18]. Overall adipogenesis progresses in an anterior to posterior, rostral to caudal, and dorsal to ventral direction [254]. BAT appears before WAT and can easily be identified at birth in most mammals [255].
Efforts have begun to indicate the endogenous origin of adipose stem cells (ASC) through the use of flow cytometric studies, single-cell RNA sequencing, and genetic lineage tracing in rodents and humans [256,257,258,259,260,261,262,263,264,265]. Lineage tracing studies in mice revealed that subcutaneous AT depots emerge from Prx1-marked cells that trace the somatic lateral plate mesoderm [266,267,268], while craniofacial AT depots originate from neural crest [269,270]. Visceral AT seems to originate from mesodermal cells marked by Wilms Tumor 1 (WT1) [271,272]. Brown adipocytes are derived from mouse embryo central dermomyotome, which expresses the homeobox transcription factor Engrailed 1 and has also the potential to form dermis and skeletal muscle [273]. BAT and skeletal muscles share the common progenitors expressing myogenic factor 5 (MYF5) [170]. In human, a recent study differentiated functional brown adipocytes from human pluripotent stem cells (MYF5+) through a paraxial mesoderm intermediate PAX3+, suggesting that the developmental origin of brown adipocytes is evolutionarily conserved in mice and humans [274]. By contrast, beige adipocytes develop postnatally and are derived from non-dermomyotome lineage [275]. New data from single-cell RNA sequencing identified a subpopulation of ASCs marked by a cell surface protein CD81, developing to beige adipocytes following cold exposure [163]. Consistent with the unique and remarkable capacity of AT to expand or remodel in various physiological settings, it appears that the molecular characteristics of ASCs are heterogeneous. More data from in vivo lineage tracing are needed to firmly establish the developmental origin and regulation of ASCs. The microenvironment in which ASCs reside strongly controls ASC proliferation, quiescence and differentiation. The role of the neighboring vascular architecture is relevant. The first fat cell clusters are detected in the vicinity of large and small arterioles and are surrounded by extensive stroma [276,277,278], suggesting that adipogenesis appears to be tightly coordinated by angiogenesis in time and space during embryogenesis [279,280,281]. Early studies suggested that innervation of AT follows angiogenesis [278]. Blood vessels secrete growth factors that attract and direct axons to innervate the vasculature. Nerves also release vascular endothelial growth factor A (VEGF-A) to guide and promote angiogenesis sprouting [282]. The setting of the neural network and neural projections has yet to be characterized.

Appendix C. Components of Adipose Tissue ECM

ECM is composed of various macromolecules including collagens, elastins, fibronectins, laminins, proteoglycan and non-proteoglycan polysaccharides [9,65]. Collagens constitute the major structural framework of AT, contributing to tensile strength, cell adhesion, chemotaxis, cell migration and sequestering various growth factors for time- and context-dependent release [9,10,65,107]. Fibronectin and laminin interact with collagen fibers to form networks and provide attachment points for integrin receptors anchored on adipocyte plasma membrane [73].

Appendix D. How Cells Respond to Mechanical Stimuli from Their Environment?

Mechanobiology defines a field at the interface of biology, physics, and bioengineering, which examines how mechanical forces influence cell behavior, cell and tissue morphogenesis, and their modifications in diseases [171,192,283]. Mechanical signaling has the potential to propagate information very quickly, ~40-fold faster than classical chemical signals [284]. Transmitted from the extracellular environment, the mechanical information is integrated into the cell membrane via cell-surface receptors-based adhesions and are routed through the cytoplasmic cytoskeleton directly to the nuclear cortex and cellular signaling pathways [218]. Mechanical deformations of the cell membrane, sensed by caveolae invaginations that define lipid raft-enriched surface domains, can also cause conformational changes of membrane channels and receptors between an open state and a closed state [285]. Cell-to-ECM and cell-to-cell mechanosensing interactions are mediated through integrins (focal adhesions) and cadherins (adherens junctions), respectively. Both junctions are connected on their cytoplasmic side with actin cytoskeleton, allowing transmission of mechanical cues inside the cell [207]. Thus, mechanical signals are transduced actively into the cells through receptor-associated actin filaments, via myosin motors generating different types of forces, such as protrusive and contractile forces [286]. The crosstalk between extracellular forces and intracellular actin structure is fundamental for cells to probe their surroundings, adapt their mechanical properties, and exert the appropriate forces required for their homeostasis in their microenvironment.

Appendix E. Lipodystrophic Laminopathies

The heterozygous LMNA p.R482W mutation, the most frequent mutation causing familial partial Dunnigan lipodystrophy (FPLD2; OMIM ID, 151660), is characterized by the loss of subcutaneous fat in the upper and lower extremities [210,287,288,289]. LMNA p.R482W mutation slows adipogenesis in fibroblastic cells extracted from patients and in human ASC [290,291]. Probably, LMNA mutation disturbs nucleus and cell mechano-sensitivity [292], and nuclear architecture [293], leading to chromatin remodeling [290,291,294]. Lamina has also a key role in chromatin organization and gene silencing by tethering long heterochromatin genomic regions to the nuclear periphery, defining a region called “lamina-associated domains” [292,295]. Collas and collaborators described the effect of a dominant-negative LMNA mutation on chromatin organization regulating the expression of the anti-adipogenic miR-335 in human adipocyte progenitors [291]. It has been also reported that lamins directly regulate epigenetic modifier complexes, such as polycomb repressor complex 2 (PRC2), a predominant H3K27 methyltransferase in mammalian cells [296]. Genome-wide analyses have shown that PRC2 and H3K27me3 are enriched on a large number of developmental regulators such as Wnt gene loci in preadipocytes [297,298]. Indeed, it was established that the H3K27 methyltransferase PRC2 directly represses Wnt genes to facilitate adipogenesis [298]. Moreover, lamin mutations and lamina dysfunction can interfere with cell functions either directly through weakening or strengthening interactions of lamins with signaling factors, or indirectly, by affecting signaling regulated by lamin-binding proteins [292,299,300]. For instance, LMNA mutation in FPLD2 patients impairs the interaction of A-type lamins with the adipogenic factor sterol regulatory element binding protein (SREBP1) [301], a transcription factor that regulates hundreds of genes involved in lipid metabolism and adipogenesis [302].

References

  1. GBD 2015 Obesity Collaborators; Afshin, A.; Forouzanfar, M.H.; Reitsma, M.B.; Sur, P.; Estep, K.; Lee, A.; Marczak, L.; Mokdad, A.H.; Moradi-Lakeh, M.; et al. Health Effects of Overweight and Obesity in 195 Countries over 25 Years. N. Engl. J. Med. 2017, 377, 13–27. [Google Scholar] [CrossRef] [PubMed]
  2. Kusminski, C.M.; Bickel, P.E.; Scherer, P.E. Targeting adipose tissue in the treatment of obesity-associated diabetes. Nat. Rev. Drug Discov. 2016, 15, 639–660. [Google Scholar] [CrossRef] [PubMed]
  3. Rosen, E.D.; Spiegelman, B.M. What we talk about when we talk about fat. Cell 2014, 156, 20–44. [Google Scholar] [CrossRef] [PubMed] [Green Version]
  4. González-Muniesa, P.; Mártinez-González, M.-A.; Hu, F.B.; Després, J.-P.; Matsuzawa, Y.; Loos, R.J.F.; Moreno, L.A.; Bray, G.A.; Martinez, J.A. Obesity. Nat. Rev. Dis. Primers 2017, 3, 17034. [Google Scholar] [CrossRef]
  5. Chouchani, E.T.; Kajimura, S. Metabolic adaptation and maladaptation in adipose tissue. Nat. Metab. 2019, 1, 189–200. [Google Scholar] [CrossRef]
  6. Morigny, P.; Boucher, J.; Arner, P.; Langin, D. Lipid and glucose metabolism in white adipocytes: Pathways, dysfunction and therapeutics. Nat. Rev. Endocrinol. 2021, 17, 276–295. [Google Scholar] [CrossRef]
  7. Ouchi, N.; Parker, J.L.; Lugus, J.J.; Walsh, K. Adipokines in inflammation and metabolic disease. Nat. Rev. Immunol. 2011, 11, 85–97. [Google Scholar] [CrossRef]
  8. Reilly, S.M.; Saltiel, A.R. Adapting to obesity with adipose tissue inflammation. Nat. Rev. Endocrinol. 2017, 13, 633–643. [Google Scholar] [CrossRef]
  9. Marcelin, G.; Gautier, E.L.; Clement, K. Adipose Tissue Fibrosis in Obesity: Etiology and Challenges. Annu. Rev. Physiol. 2022, 84, 135–155. [Google Scholar] [CrossRef]
  10. Marcelin, G.; Silveira, A.L.M.; Martins, L.B.; Ferreira, A.V.M.; Clément, K. Deciphering the cellular interplays underlying obesity-induced adipose tissue fibrosis. J. Clin. Investig. 2019, 129, 4032–4040. [Google Scholar] [CrossRef] [PubMed]
  11. Acosta, J.R.; Douagi, I.; Andersson, D.P.; Bäckdahl, J.; Rydén, M.; Arner, P.; Laurencikiene, J. Increased fat cell size: A major phenotype of subcutaneous white adipose tissue in non-obese individuals with type 2 diabetes. Diabetologia 2016, 59, 560–570. [Google Scholar] [CrossRef] [Green Version]
  12. Mejhert, N.; Rydén, M. Novel aspects on the role of white adipose tissue in type 2 diabetes. Curr. Opin. Pharmacol. 2020, 55, 47–52. [Google Scholar] [CrossRef] [PubMed]
  13. Hoffstedt, J.; Arner, E.; Wahrenberg, H.; Andersson, D.P.; Qvisth, V.; Löfgren, P.; Rydén, M.; Thörne, A.; Wirén, M.; Palmér, M.; et al. Regional impact of adipose tissue morphology on the metabolic profile in morbid obesity. Diabetologia 2010, 53, 2496–2503. [Google Scholar] [CrossRef] [PubMed] [Green Version]
  14. Arner, E.; Westermark, P.O.; Spalding, K.L.; Britton, T.; Rydén, M.; Frisén, J.; Bernard, S.; Arner, P. Adipocyte turnover: Relevance to human adipose tissue morphology. Diabetes 2010, 59, 105–109. [Google Scholar] [CrossRef] [Green Version]
  15. Eriksson-Hogling, D.; Andersson, D.P.; Bäckdahl, J.; Hoffstedt, J.; Rössner, S.; Thorell, A.; Arner, E.; Arner, P.; Rydén, M. Adipose tissue morphology predicts improved insulin sensitivity following moderate or pronounced weight loss. Int. J. Obes. 2015, 39, 893–898. [Google Scholar] [CrossRef]
  16. Pellegrinelli, V.; Carobbio, S.; Vidal-Puig, A. Adipose tissue plasticity: How fat depots respond differently to pathophysiological cues. Diabetologia 2016, 59, 1075–1088. [Google Scholar] [CrossRef] [Green Version]
  17. Berry, D.C.; Jiang, Y.; Graff, J.M. Emerging roles of adipose progenitor cells in tissue development, homeostasis, expansion and thermogenesis. Trends Endocrinol. Metab. 2016, 27, 574–585. [Google Scholar] [CrossRef]
  18. Spalding, K.L.; Arner, E.; Westermark, P.O.; Bernard, S.; Buchholz, B.A.; Bergmann, O.; Blomqvist, L.; Hoffstedt, J.; Näslund, E.; Britton, T.; et al. Dynamics of fat cell turnover in humans. Nature 2008, 453, 783–787. [Google Scholar] [CrossRef] [PubMed]
  19. Rigamonti, A.; Brennand, K.; Lau, F.; Cowan, C.A. Rapid Cellular Turnover in Adipose Tissue. PLoS ONE 2011, 6, e17637. [Google Scholar] [CrossRef] [Green Version]
  20. Kim, S.M.; Lun, M.; Wang, M.; Senyo, S.E.; Guillermier, C.; Patwari, P. Loss of White Adipose Hyperplastic Potential Is Associated with Enhanced Susceptibility to Insulin Resistance. Cell Metab. 2014, 20, 1049–1058. [Google Scholar] [CrossRef] [PubMed] [Green Version]
  21. Berry, R.; Jeffery, E.; Rodeheffer, M.S. Perspective Weighing in on Adipocyte Precursors. Cell Metab. 2013, 19, 8–20. [Google Scholar] [CrossRef] [Green Version]
  22. Guillermier, C.; Fazeli, P.K.; Kim, S.; Lun, M.; Zuflacht, J.P.; Milian, J.; Lee, H.; Francois-Saint-Cyr, H.; Horreard, F.; Larson, D.; et al. Imaging mass spectrometry demonstrates age-related decline in human adipose plasticity. JCI Insight 2017, 2. [Google Scholar] [CrossRef] [Green Version]
  23. Hirsch, J.; Han, P.W. Cellularity of rat adipose tissue: Effects of growth, starvation, and obesity. J. Lipid Res. 1969, 10, 77–82. [Google Scholar] [CrossRef]
  24. Wang, Q.A.; Tao, C.; Gupta, R.K.; Scherer, P.E. Tracking adipogenesis during white adipose tissue development, expansion and regeneration. Nat. Med. 2013, 19, 1338–1344. [Google Scholar] [CrossRef]
  25. Ghaben, A.L.; Scherer, P.E. Adipogenesis and metabolic health. Nat. Rev. Mol. Cell Biol. 2019, 20, 242–258. [Google Scholar] [CrossRef]
  26. Jo, J.; Gavrilova, O.; Pack, S.; Jou, W.; Mullen, S.; Sumner, A.E.; Cushman, S.W.; Periwal, V. Hypertrophy and/or hyperplasia: Dynamics of adipose tissue growth. PLoS Comput. Biol. 2009, 5, e1000324. [Google Scholar] [CrossRef]
  27. Jeffery, E.; Wing, A.; Holtrup, B.; Sebo, Z.; Kaplan, J.L.; Saavedra-Peña, R.; Church, C.D.; Colman, L.; Berry, R.; Rodeheffer, M.S. The Adipose Tissue Microenvironment Regulates Depot-Specific Adipogenesis in Obesity. Cell Metab. 2016, 24, 142–150. [Google Scholar] [CrossRef] [Green Version]
  28. Arner, P.; Andersson, D.P.; Thörne, A.; Wirén, M.; Hoffstedt, J.; Näslund, E.; Thorell, A.; Rydén, M. Variations in the size of the major omentum are primarily determined by fat cell number. J. Clin. Endocrinol. Metab. 2013, 98, E897–E901. [Google Scholar] [CrossRef] [PubMed] [Green Version]
  29. Jeffery, E.; Church, C.D.; Holtrup, B.; Colman, L.; Rodeheffer, M.S. Rapid depot-specific activation of adipocyte precursor cells at the onset of obesity. Nat. Cell Biol. 2015, 17, 376–385. [Google Scholar] [CrossRef]
  30. Vague, J. The degree of masculine differentiation of obesities: A factor determining predisposition to diabetes, atherosclerosis, gout, and uric calculous disease. Am. J. Clin. Nutr. 1956, 4, 20–34. [Google Scholar] [CrossRef]
  31. Rydén, M.; Andersson, D.P.; Bergström, I.B.; Arner, P. Adipose tissue and metabolic alterations: Regional differences in fat cell size and number matter, but differently: A cross-sectional study. J. Clin. Endocrinol. Metab. 2014, 99, E1870–E1876. [Google Scholar] [CrossRef] [PubMed] [Green Version]
  32. Unamuno, X.; Gómez-Ambrosi, J.; Rodríguez, A.; Becerril, S.; Frühbeck, G.; Catalán, V. Adipokine dysregulation and adipose tissue inflammation in human obesity. Eur. J. Clin. Investig. 2018, 48, e12997. [Google Scholar] [CrossRef] [PubMed] [Green Version]
  33. Balistreri, C.R.; Caruso, C.; Candore, G. The Role of Adipose Tissue and Adipokines in Obesity-Related Inflammatory Diseases. Mediat. Inflamm. 2010, 2010, 802078. [Google Scholar] [CrossRef] [PubMed]
  34. Russo, L.; Lumeng, C.N. Properties and functions of adipose tissue macrophages in obesity. Immunology 2018, 155, 407–417. [Google Scholar] [CrossRef]
  35. Weisberg, S.P.; Mccann, D.; Desai, M.; Rosenbaum, M.; Leibel, R.L.; Ferrante, A.W. Obesity Is Associated with Macrophage Accumulation. J. Clin. Investig. 2003, 112, 1796–1808. [Google Scholar] [CrossRef] [PubMed]
  36. Xu, H.; Barnes, G.T.; Yang, Q.; Tan, G.; Yang, D.; Chou, C.J.; Sole, J.; Nichols, A.; Ross, J.S.; Tartaglia, L.A.; et al. Chronic inflammation in fat plays a crucial role in the development of obesity-related insulin resistance. J. Clin. Investig. 2003, 112, 1821–1830. [Google Scholar] [CrossRef] [PubMed]
  37. DiSpirito, J.R.; Zemmour, D.; Ramanan, D.; Cho, J.; Zilionis, R.; Klein, A.M.; Benoist, C.; Mathis, D. Molecular diversification of regulatory T cells in nonlymphoid tissues. Sci. Immunol. 2018, 3, eaat5861. [Google Scholar] [CrossRef]
  38. Li, C.; DiSpirito, J.R.; Zemmour, D.; Spallanzani, R.G.; Kuswanto, W.; Benoist, C.; Mathis, D. TCR Transgenic Mice Reveal Stepwise, Multi-site Acquisition of the Distinctive Fat-Treg Phenotype. Cell 2018, 174, 285–299.e12. [Google Scholar] [CrossRef] [Green Version]
  39. Hildreth, A.D.; Ma, F.; Wong, Y.Y.; Sun, R.; Pellegrini, M.; O’Sullivan, T.E. Single-cell sequencing of human white adipose tissue identifies new cell states in health and obesity. Nat. Immunol. 2021, 22, 639–653. [Google Scholar] [CrossRef]
  40. Jaitin, D.A.; Adlung, L.; Thaiss, C.A.; Weiner, A.; Li, B.; Descamps, H.; Lundgren, P.; Bleriot, C.; Liu, Z.; Deczkowska, A.; et al. Lipid-Associated Macrophages Control Metabolic Homeostasis in a Trem2-Dependent Manner. Cell 2019, 178, 686–698.e14. [Google Scholar] [CrossRef]
  41. Feuerer, M.; Herrero, L.; Cipolletta, D.; Naaz, A.; Wong, J.; Nayer, A.; Lee, J.; Goldfine, A.B.; Benoist, C.; Shoelson, S.; et al. Lean, but not obese, fat is enriched for a unique population of regulatory T cells that affect metabolic parameters. Nat. Med. 2009, 15, 930–939. [Google Scholar] [CrossRef] [PubMed]
  42. Nishimura, S.; Manabe, I.; Nagasaki, M.; Eto, K.; Yamashita, H.; Ohsugi, M.; Otsu, M.; Hara, K.; Ueki, K.; Sugiura, S.; et al. CD8+ effector T cells contribute to macrophage recruitment and adipose tissue inflammation in obesity. Nat. Med. 2009, 15, 914–920. [Google Scholar] [CrossRef] [PubMed]
  43. Nishimura, S.; Manabe, I.; Takaki, S.; Nagasaki, M.; Otsu, M.; Yamashita, H.; Sugita, J.; Yoshimura, K.; Eto, K.; Komuro, I.; et al. Adipose Natural Regulatory B Cells Negatively Control Adipose Tissue Inflammation. Cell Metab. 2013, 18, 759–766. [Google Scholar] [CrossRef] [Green Version]
  44. Kohlgruber, A.C.; Gal-Oz, S.T.; Lamarche, N.M.; Shimazaki, M.; Duquette, D.; Nguyen, H.N.; Mina, A.I.; Paras, T.; Tavakkoli, A.; Von Andrian, U.; et al. γδ T cells producing interleukin-17A regulate adipose regulatory T cell homeostasis and thermogenesis. Nat. Immunol. 2018, 19, 464–474. [Google Scholar] [CrossRef] [PubMed]
  45. Lynch, L.; Hogan, A.E.; Duquette, D.; Lester, C.; Banks, A.; LeClair, K.; Cohen, D.E.; Ghosh, A.; Lu, B.; Corrigan, M.; et al. iNKT Cells Induce FGF21 for Thermogenesis and Are Required for Maximal Weight Loss in GLP1 Therapy. Cell Metab. 2016, 24, 510–519. [Google Scholar] [CrossRef] [PubMed] [Green Version]
  46. Hu, B.; Jin, C.; Zeng, X.; Resch, J.M.; Jedrychowski, M.P.; Yang, Z.; Desai, B.N.; Banks, A.S.; Lowell, B.B.; Mathis, D.; et al. γδ T cells and adipocyte IL-17RC control fat innervation and thermogenesis. Nature 2020, 578, 610–614. [Google Scholar] [CrossRef] [PubMed]
  47. Han, S.J.; Glatman Zaretsky, A.; Andrade-Oliveira, V.; Collins, N.; Dzutsev, A.; Shaik, J.; Morais da Fonseca, D.; Harrison, O.J.; Tamoutounour, S.; Byrd, A.L.; et al. White Adipose Tissue Is a Reservoir for Memory T Cells and Promotes Protective Memory Responses to Infection. Immunity 2017, 47, 1154–1168.e6. [Google Scholar] [CrossRef] [PubMed] [Green Version]
  48. Boulenouar, S.; Michelet, X.; Duquette, D.; Alvarez, D.; Hogan, A.E.; Dold, C.; O’Connor, D.; Stutte, S.; Tavakkoli, A.; Winters, D.; et al. Adipose Type One Innate Lymphoid Cells Regulate Macrophage Homeostasis through Targeted Cytotoxicity. Immunity 2017, 46, 273–286. [Google Scholar] [CrossRef] [Green Version]
  49. Li, C.; Wang, G.; Sivasami, P.; Ramirez, R.N.; Zhang, Y.; Benoist, C.; Mathis, D. Interferon-α-producing plasmacytoid dendritic cells drive the loss of adipose tissue regulatory T cells during obesity. Cell Metab. 2021, 33, 1610–1623.e5. [Google Scholar] [CrossRef]
  50. Talukdar, S.; Oh, D.Y.; Bandyopadhyay, G.; Li, D.; Xu, J.; McNelis, J.; Lu, M.; Li, P.; Yan, Q.; Zhu, Y.; et al. Neutrophils mediate insulin resistance in mice fed a high-fat diet through secreted elastase. Nat. Med. 2012, 18, 1407–1412. [Google Scholar] [CrossRef] [Green Version]
  51. Wang, L.; Luo, Y.; Luo, L.; Wu, D.; Ding, X.; Zheng, H.; Wu, H.; Liu, B.; Yang, X.; Silva, F.; et al. Adiponectin restrains ILC2 activation by AMPK-mediated feedback inhibition of IL-33 signaling. J. Exp. Med. 2021, 218. [Google Scholar] [CrossRef] [PubMed]
  52. Goldberg, E.L.; Shchukina, I.; Youm, Y.H.; Ryu, S.; Tsusaka, T.; Young, K.C.; Camell, C.D.; Dlugos, T.; Artyomov, M.N.; Dixit, V.D. IL-33 causes thermogenic failure in aging by expanding dysfunctional adipose ILC2. Cell Metab. 2021, 33, 2277–2287.e5. [Google Scholar] [CrossRef] [PubMed]
  53. Lee, M.W.; Odegaard, J.I.; Mukundan, L.; Qiu, Y.; Molofsky, A.B.; Nussbaum, J.C.; Yun, K.; Locksley, R.M.; Chawla, A. Activated type 2 innate lymphoid cells regulate beige fat biogenesis. Cell 2015, 160, 74–87. [Google Scholar] [CrossRef] [PubMed] [Green Version]
  54. Molofsky, A.B.; Nussbaum, J.C.; Liang, H.; Van Dyken, S.J.; Cheng, L.E.; Mohapatra, A.; Chawla, A.; Locksley, R.M. Innate lymphoid type 2 cells sustain visceral adipose tissue eosinophils and alternatively activated macrophages. J. Exp. Med. 2013, 210, 535–549. [Google Scholar] [CrossRef]
  55. Brestoff, J.R.; Kim, B.S.; Saenz, S.A.; Stine, R.R.; Monticelli, L.A.; Sonnenberg, G.F.; Thome, J.J.; Farber, D.L.; Lutfy, K.; Seale, P.; et al. Group 2 innate lymphoid cells promote beiging of white adipose tissue and limit obesity. Nature 2015, 519, 242–246. [Google Scholar] [CrossRef] [Green Version]
  56. Wernstedt Asterholm, I.; Tao, C.; Morley, T.S.; Wang, Q.A.; Delgado-Lopez, F.; Wang, Z.V.; Scherer, P.E. Adipocyte inflammation is essential for healthy adipose tissue expansion and remodeling. Cell Metab. 2014, 20, 103–118. [Google Scholar] [CrossRef] [Green Version]
  57. Zhu, Q.; An, Y.A.; Kim, M.; Zhang, Z.; Zhao, S.; Zhu, Y.; Asterholm, I.W.; Kusminski, C.M.; Scherer, P.E. Suppressing adipocyte inflammation promotes insulin resistance in mice. Mol. Metab. 2020, 39, 101010. [Google Scholar] [CrossRef]
  58. Kammoun, H.L.; Kraakman, M.J.; Febbraio, M.A. Adipose tissue inflammation in glucose metabolism. Rev. Endocr. Metab. Disord. 2014, 15, 31–44. [Google Scholar] [CrossRef]
  59. Vila, I.K.; Badin, P.M.; Marques, M.A.; Monbrun, L.; Lefort, C.; Mir, L.; Louche, K.; Bourlier, V.; Roussel, B.; Gui, P.; et al. Immune cell Toll-like receptor 4 mediates the development of obesity- and endotoxemia-associated adipose tissue fibrosis. Cell Rep. 2014, 7, 1116–1129. [Google Scholar] [CrossRef]
  60. Wynn, T.A.; Ramalingam, T.R. Mechanisms of fibrosis: Therapeutic translation for fibrotic disease. Nat. Med. 2012, 18, 1028–1040. [Google Scholar] [CrossRef] [PubMed] [Green Version]
  61. Guzmán-Ruiz, R.; Tercero-Alcázar, C.; Rabanal-Ruiz, Y.; Díaz-Ruiz, A.; El Bekay, R.; Rangel-Zuñiga, O.A.; Navarro-Ruiz, M.C.; Molero, L.; Membrives, A.; Ruiz-Rabelo, J.F.; et al. Adipose tissue depot-specific intracellular and extracellular cues contributing to insulin resistance in obese individuals. FASEB J. 2020, 34, 7520–7539. [Google Scholar] [CrossRef] [PubMed] [Green Version]
  62. Reggio, S.; Rouault, C.; Poitou, C.; Bichet, J.C.; Prifti, E.; Bouillot, J.L.; Rizkalla, S.; Lacasa, D.; Tordjman, J.; Clément, K. Increased Basement Membrane Components in Adipose Tissue During Obesity: Links With TGFβ and Metabolic Phenotypes. J. Clin. Endocrinol. Metab. 2016, 101, 2578–2587. [Google Scholar] [CrossRef] [PubMed] [Green Version]
  63. Pastel, E.; Price, E.; Sjöholm, K.; McCulloch, L.J.; Rittig, N.; Liversedge, N.; Knight, B.; Møller, N.; Svensson, P.A.; Kos, K. Lysyl oxidase and adipose tissue dysfunction. Metabolism 2018, 78, 118–127. [Google Scholar] [CrossRef] [PubMed]
  64. Huang, A.; Lin, Y.; Kao, L.; Chiou, Y.; Lee, G.; Lin, H.; Wu, C.; Chang, C.; Lee, K.; Hsueh, Y.; et al. Inflammation-induced macrophage lysyl oxidase in adipose stiffening and dysfunction in obesity. Clin. Transl. Med. 2021, 11, e543. [Google Scholar] [CrossRef] [PubMed]
  65. Datta, R.; Podolsky, M.J.; Atabai, K. Fat fibrosis: Friend or foe? JCI Insight 2018, 3, e122289. [Google Scholar] [CrossRef] [PubMed]
  66. Khan, T.; Muise, E.S.; Iyengar, P.; Wang, Z.V.; Chandalia, M.; Abate, N.; Zhang, B.B.; Bonaldo, P.; Chua, S.; Scherer, P.E. Metabolic Dysregulation and Adipose Tissue Fibrosis: Role of Collagen VI. Mol. Cell. Biol. 2009, 29, 1575–1591. [Google Scholar] [CrossRef] [Green Version]
  67. Lassen, P.B.; Charlotte, F.; Liu, Y.; Bedossa, P.; Le Naour, G.; Tordjman, J.; Poitou, C.; Bouillot, J.L.; Genser, L.; Zucker, J.D.; et al. The FAT Score, a Fibrosis Score of Adipose Tissue: Predicting Weight-Loss Outcome after Gastric Bypass. J. Clin. Endocrinol. Metab. 2017, 102, 2443–2453. [Google Scholar] [CrossRef] [PubMed]
  68. Pasarica, M.; Sereda, O.R.; Redman, L.M.; Albarado, D.C.; Hymel, D.T.; Roan, L.E.; Rood, J.C.; Burk, D.H.; Smith, S.R. Reduced Adipose Tissue Oxygenation in Human Obesity: Evidence for Rarefaction, Macrophage Chemotaxis, and Inflammation Without an Angiogenic Response. Diabetes 2009, 58, 718–725. [Google Scholar] [CrossRef] [Green Version]
  69. Spencer, M.; Unal, R.; Zhu, B.; Rasouli, N.; McGehee, R.E.; Peterson, C.A.; Kern, P.A. Adipose tissue extracellular matrix and vascular abnormalities in obesity and insulin resistance. J. Clin. Endocrinol. Metab. 2011, 96, E1990–E1998. [Google Scholar] [CrossRef] [Green Version]
  70. Abdennour, M.; Reggio, S.; Le Naour, G.; Liu, Y.; Poitou, C.; Aron-Wisnewsky, J.; Charlotte, F.; Bouillot, J.L.; Torcivia, A.; Sasso, M.; et al. Association of adipose tissue and liver fibrosis with tissue stiffness in morbid obesity: Links with diabetes and BMI loss after gastric bypass. J. Clin. Endocrinol. Metab. 2014, 99, 898–907. [Google Scholar] [CrossRef] [Green Version]
  71. Muir, L.A.; Neeley, C.K.; Meyer, K.A.; Baker, N.A.; Brosius, A.M.; Washabaugh, A.R.; Varban, O.A.; Finks, J.F.; Zamarron, B.F.; Flesher, C.G.; et al. Adipose Tissue Fibrosis, Hypertrophy, and Hyperplasia: Correlations with Diabetes in Human Obesity. Obesity 2016, 24, 597–605. [Google Scholar] [CrossRef] [PubMed]
  72. Díaz-Ruiz, A.; Guzmán-Ruiz, R.; Moreno, N.R.; García-Rios, A.; Delgado-Casado, N.; Membrives, A.; Túnez, I.; El Bekay, R.; Fernández-Real, J.M.; Tovar, S.; et al. Proteasome Dysfunction Associated to Oxidative Stress and Proteotoxicity in Adipocytes Compromises Insulin Sensitivity in Human Obesity. Antioxid. Redox Signal. 2015, 23, 597–612. [Google Scholar] [CrossRef] [PubMed] [Green Version]
  73. Pope, B.D.; Warren, C.R.; Parker, K.K.; Cowan, C.A. Microenvironmental Control of Adipocyte Fate and Function. Trends Cell Biol. 2016, 26, 745–755. [Google Scholar] [CrossRef] [PubMed]
  74. Burke, S.; Nagajyothi, F.; Thi, M.M.; Hanani, M.; Scherer, P.E.; Tanowitz, H.B.; Spray, D.C. Adipocytes in both brown and white adipose tissue of adult mice are functionally connected via gap junctions: Implications for Chagas disease. Microbes Infect. 2014, 16, 893–901. [Google Scholar] [CrossRef] [PubMed] [Green Version]
  75. Zhu, Y.; Li, N.; Huang, M.; Chen, X.; An, Y.A.; Li, J.; Zhao, S.; Funcke, J.B.; Cao, J.; He, Z.; et al. Activating Connexin43 gap junctions primes adipose tissue for therapeutic intervention. Acta Pharm. Sin. B 2022, 12, 3063–3072. [Google Scholar] [CrossRef]
  76. Zhu, Y.; Gao, Y.; Tao, C.; Shao, M.; Zhao, S.; Huang, W.; Yao, T.; Johnson, J.A.; Liu, T.; Cypess, A.M.; et al. Connexin 43 Mediates White Adipose Tissue Beiging by Facilitating the Propagation of Sympathetic Neuronal Signals. Cell Metab. 2016, 24, 420–433. [Google Scholar] [CrossRef] [Green Version]
  77. Revel, J.P.; Karnovsky, M.J. Hexagonal array of subunits in intercellular junctions of the mouse heart and liver. J. Cell Biol. 1967, 33, C7. [Google Scholar] [CrossRef]
  78. Hervé, J.C.; Bourmeyster, N.; Sarrouilhe, D. Diversity in protein-protein interactions of connexins: Emerging roles. Biochim. Biophys. Acta 2004, 1662, 22–41. [Google Scholar] [CrossRef] [Green Version]
  79. Weber, P.A.; Chang, H.C.; Spaeth, K.E.; Nitsche, J.M.; Nicholson, B.J. The permeability of gap junction channels to probes of different size is dependent on connexin composition and permeant-pore affinities. Biophys. J. 2004, 87, 958–973. [Google Scholar] [CrossRef] [Green Version]
  80. Bartness, T.J.; Liu, Y.; Shrestha, Y.B.; Ryu, V. Neural innervation of white adipose tissue and the control of lipolysis. Front. Neuroendocrinol. 2014, 35, 473–493. [Google Scholar] [CrossRef] [Green Version]
  81. Murano, I.; Barbatelli, G.; Giordano, A.; Cinti, S. Noradrenergic parenchymal nerve fiber branching after cold acclimatisation correlates with brown adipocyte density in mouse adipose organ. J. Anat. 2009, 214, 171–178. [Google Scholar] [CrossRef] [PubMed]
  82. Jiang, H.; Ding, X.; Cao, Y.; Wang, H.; Zeng, W. Dense Intra-adipose Sympathetic Arborizations Are Essential for Cold-Induced Beiging of Mouse White Adipose Tissue. Cell Metab. 2017, 26, 686–692.e3. [Google Scholar] [CrossRef] [Green Version]
  83. Zeng, W.; Pirzgalska, R.M.; Pereira, M.M.A.; Kubasova, N.; Barateiro, A.; Seixas, E.; Lu, Y.-H.; Kozlova, A.; Voss, H.; Martins, G.G.; et al. Sympathetic Neuro-adipose Connections Mediate Leptin-Driven Lipolysis. Cell 2015, 163, 84–94. [Google Scholar] [CrossRef] [Green Version]
  84. Néchad, M.; Ruka, E.; Thibault, J. Production of nerve growth factor by brown fat in culture: Relation with the in vivo developmental stage of the tissue. Comp. Biochem. Physiol. Comp. Physiol. 1994, 107, 381–388. [Google Scholar] [CrossRef]
  85. Zeng, X.; Ye, M.; Resch, J.M.; Jedrychowski, M.P.; Hu, B.; Lowell, B.B.; Ginty, D.D.; Spiegelman, B.M. Innervation of thermogenic adipose tissue via a calsyntenin 3β-S100b axis. Nature 2019, 569, 229–235. [Google Scholar] [CrossRef] [PubMed]
  86. Riuzzi, F.; Chiappalupi, S.; Arcuri, C.; Giambanco, I.; Sorci, G.; Donato, R. S100 proteins in obesity: Liaisons dangereuses. Cell. Mol. Life Sci. 2020, 77, 129–147. [Google Scholar] [CrossRef]
  87. Pellegrinelli, V.; Peirce, V.J.; Howard, L.; Virtue, S.; Türei, D.; Senzacqua, M.; Frontini, A.; Dalley, J.W.; Horton, A.R.; Bidault, G.; et al. Adipocyte-secreted BMP8b mediates adrenergic-induced remodeling of the neuro-vascular network in adipose tissue. Nat. Commun. 2018, 9, 4974. [Google Scholar] [CrossRef] [PubMed]
  88. Brito, N.A.; Brito, M.N.; Bartness, T.J. Differential sympathetic drive to adipose tissues after food deprivation, cold exposure or glucoprivation. Am. J. Physiol. Regul. Integr. Comp. Physiol. 2008, 294, R1445–R1452. [Google Scholar] [CrossRef] [Green Version]
  89. Cao, Y.; Wang, H.; Zeng, W. Whole-tissue 3D imaging reveals intra-adipose sympathetic plasticity regulated by NGF-TrkA signal in cold-induced beiging. Protein Cell 2018, 9, 527–539. [Google Scholar] [CrossRef] [Green Version]
  90. Blaszkiewicz, M.; Willows, J.W.; Johnson, C.P.; Townsend, K.L. The Importance of Peripheral Nerves in Adipose Tissue for the Regulation of Energy Balance. Biology 2019, 8, 10. [Google Scholar] [CrossRef] [Green Version]
  91. O’Brien, C.J.O.; Haberman, E.R.; Domingos, A.I. A Tale of Three Systems: Toward a Neuroimmunoendocrine Model of Obesity. Annu. Rev. Cell Dev. Biol. 2021, 37, 549–573. [Google Scholar] [CrossRef] [PubMed]
  92. Giordano, A.; Song, C.K.; Bowers, R.R.; Ehlen, J.C.; Frontini, A.; Cinti, S.; Bartness, T.J. White adipose tissue lacks significant vagal innervation and immunohistochemical evidence of parasympathetic innervation. Am. J. Physiol. Regul. Integr. Comp. Physiol. 2006, 291, R1243–R1255. [Google Scholar] [CrossRef] [PubMed] [Green Version]
  93. Saxena, U.; Klein, M.G.; Goldberg, I.J. Transport of lipoprotein lipase across endothelial cells. Proc. Natl. Acad. Sci. USA 1991, 88, 2254–2258. [Google Scholar] [CrossRef] [PubMed] [Green Version]
  94. Zechner, R.; Kienesberger, P.C.; Haemmerle, G.; Zimmermann, R.; Lass, A. Adipose triglyceride lipase and the lipolytic catabolism of cellular fat stores. J. Lipid Res. 2009, 50, 3–21. [Google Scholar] [CrossRef] [PubMed] [Green Version]
  95. Pepino, M.Y.; Kuda, O.; Samovski, D.; Abumrad, N.A. Structure-function of CD36 and importance of fatty acid signal transduction in fat metabolism. Annu. Rev. Nutr. 2014, 34, 281–303. [Google Scholar] [CrossRef] [Green Version]
  96. Abumradsb, N.A.; Raafat El-Maghrabis, M.; Amrill, E.-Z.; Lopezs, E.; Grimaldill, P.A. Cloning of a Rat Adipocyte Membrane Protein Implicated in Binding or Transport of Long-chain Fatty Acids That Is Induced during Preadipocyte Differentiation. J. Biol. Chem. 1993, 268, 17665–17668. [Google Scholar] [CrossRef]
  97. Tang, W.; Zeve, D.; Suh, J.M.; Bosnakovski, D.; Kyba, M.; Hammer, R.E.; Tallquist, M.D.; Graff, J.M. White Fat Progenitor Cells Reside in the Adipose Vasculature. Science 2008, 322, 583–586. [Google Scholar] [CrossRef] [PubMed] [Green Version]
  98. Jiang, Y.; Berry, D.C.; Tang, W.; Graff, J.M. Independent Stem Cell Lineages Regulate Adipose Organogenesis and Adipose Homeostasis. Cell Rep. 2014, 9, 1007–1022. [Google Scholar] [CrossRef] [Green Version]
  99. Vishvanath, L.; MacPherson, K.A.; Hepler, C.; Wang, Q.A.; Shao, M.; Spurgin, S.B.; Wang, M.Y.; Kusminski, C.M.; Morley, T.S.; Gupta, R.K. Pdgfrβ+ Mural Preadipocytes Contribute to Adipocyte Hyperplasia Induced by High-Fat-Diet Feeding and Prolonged Cold Exposure in Adult Mice. Cell Metab. 2015, 23, 350–359. [Google Scholar] [CrossRef] [PubMed] [Green Version]
  100. Vishvanath, L.; Long, J.Z.; Spiegelman, B.M.; Gupta, R.K. Do Adipocytes Emerge from Mural Progenitors? Cell Stem Cell 2017, 20, 585–586. [Google Scholar] [CrossRef]
  101. Asano, A.; Morimatsu, M.; Nikami, H.; Yoshida, T.; Saito, M. Adrenergic activation of vascular endothelial growth factor mRNA expression in rat brown adipose tissue: Implication in cold-induced angiogenesis. Biochem. J. 1997, 328 Pt 1, 179–183. [Google Scholar] [CrossRef] [PubMed] [Green Version]
  102. Asano, A.; Kimura, K.; Saito, M. Cold-induced mRNA expression of angiogenic factors in rat brown adipose tissue. J. Vet. Med. Sci. 1999, 61, 403–409. [Google Scholar] [CrossRef] [PubMed] [Green Version]
  103. Herold, J.; Kalucka, J. Angiogenesis in Adipose Tissue: The Interplay between Adipose and Endothelial Cells. Front. Physiol. 2021, 11, 624903. [Google Scholar] [CrossRef] [PubMed]
  104. Sung, H.K.; Doh, K.O.; Son, J.E.; Park, J.G.; Bae, Y.; Choi, S.; Nelson, S.M.L.; Cowling, R.; Nagy, K.; Michael, I.P.; et al. Adipose vascular endothelial growth factor regulates metabolic homeostasis through angiogenesis. Cell Metab. 2013, 17, 61–72. [Google Scholar] [CrossRef] [Green Version]
  105. Zhao, Y.; Li, X.; Yang, L.; Eckel-Mahan, K.; Tong, Q.; Gu, X.; Kolonin, M.G.; Sun, K. Transient Overexpression of Vascular Endothelial Growth Factor A in Adipose Tissue Promotes Energy Expenditure via Activation of the Sympathetic Nervous System. Mol. Cell. Biol. 2018, 38, e00242-18. [Google Scholar] [CrossRef] [Green Version]
  106. Trim, W.V.; Lynch, L. Immune and non-immune functions of adipose tissue leukocytes. Nat. Rev. Immunol. 2022, 22, 371–386. [Google Scholar] [CrossRef]
  107. Crewe, C.; An, Y.A.; Scherer, P.E. The ominous triad of adipose tissue dysfunction: Inflammation, fibrosis, and impaired an-giogenesis. J. Clin. Investig. 2017, 127, 74–82. [Google Scholar] [CrossRef] [Green Version]
  108. Pirzgalska, R.M.; Seixas, E.; Seidman, J.S.; Link, V.M.; Sánchez, N.M.; Mahú, I.; Mendes, R.; Gres, V.; Kubasova, N.; Morris, I.; et al. Sympathetic neuron-associated macrophages contribute to obesity by importing and metabolizing norepinephrine. Nat. Med. 2017, 23, 1309–1318. [Google Scholar] [CrossRef]
  109. Hotamisligil, G.S.; Shargill, N.S.; Spiegelman, B.M. Adipose Expression of Tumor Necrosis Factor-α: Direct Role in Obesity-Linked Insulin Resistance. Science 1993, 259, 87–91. [Google Scholar] [CrossRef]
  110. Amano, S.U.; Cohen, J.L.; Vangala, P.; Tencerova, M.; Nicoloro, S.M.; Yawe, J.C.; Shen, Y.; Czech, M.P.; Aouadi, M. Local proliferation of macrophages contributes to obesity-associated adipose tissue inflammation. Cell Metab. 2014, 19, 162–171. [Google Scholar] [CrossRef] [Green Version]
  111. Silva, H.M.; Báfica, A.; Rodrigues-Luiz, G.F.; Chi, J.; D’Emery Alves Santos, P.; Reis, B.S.; Hoytema Van Konijnenburg, D.P.; Crane, A.; Arifa, R.D.N.; Martin, P.; et al. Vasculature-associated fat macrophages readily adapt to inflammatory and metabolic challenges. J. Exp. Med. 2019, 216, 786–806. [Google Scholar] [CrossRef] [Green Version]
  112. Kratz, M.; Coats, B.R.; Hisert, K.B.; Hagman, D.; Mutskov, V.; Peris, E.; Schoenfelt, K.Q.; Kuzma, J.N.; Larson, I.; Billing, P.S.; et al. Metabolic dysfunction drives a mechanistically distinct proinflammatory phenotype in adipose tissue macrophages. Cell Metab. 2014, 20, 614–625. [Google Scholar] [CrossRef] [PubMed] [Green Version]
  113. Gordon, S.; Taylor, P.R. Monocyte and macrophage heterogeneity. Nat. Rev. Immunol. 2005, 5, 953–964. [Google Scholar] [CrossRef] [PubMed]
  114. Biswas, S.K.; Mantovani, A. Orchestration of metabolism by macrophages. Cell Metab. 2012, 15, 432–437. [Google Scholar] [CrossRef] [PubMed] [Green Version]
  115. Brestoff, J.R.; Wilen, C.B.; Moley, J.R.; Li, Y.; Zou, W.; Malvin, N.P.; Rowen, M.N.; Saunders, B.T.; Ma, H.; Mack, M.R.; et al. Intercellular Mitochondria Transfer to Macrophages Regulates White Adipose Tissue Homeostasis and Is Impaired in Obesity. Cell Metab. 2021, 33, 270–282.e8. [Google Scholar] [CrossRef] [PubMed]
  116. Lecoutre, S.; Clément, K.; Dugail, I. Obesity-Related Adipose Tissue Remodeling in the Light of Extracellular Mitochondria Transfer. Int. J. Mol. Sci. 2022, 23, 632. [Google Scholar] [CrossRef] [PubMed]
  117. Torralba, D.; Baixauli, F.; Sánchez-Madrid, F. Mitochondria Know No Boundaries: Mechanisms and Functions of Intercellular Mitochondrial Transfer. Front. Cell Dev. Biol. 2016, 4, 107. [Google Scholar] [CrossRef] [Green Version]
  118. Richard, A.J.; White, U.; Elks, C.M.; Stephens, J.M. Adipose Tissue: Physiology to Metabolic Dysfunction. In Endotext [Internet]; MDText.com, Inc.: South Dartmouth, MA, USA, 2000. [Google Scholar]
  119. Cinti, S. The adipose organ. Prostaglandins. Leukot. Essent. Fatty Acids 2005, 73, 9–15. [Google Scholar] [CrossRef] [PubMed]
  120. Yang, W.; Thein, S.; Lim, C.Y.; Ericksen, R.E.; Sugii, S.; Xu, F.; Robinson, R.C.; Kim, J.B.; Han, W. Arp2/3 complex regulates adipogenesis by controlling cortical actin remodelling. Biochem. J. 2014, 464, 179–192. [Google Scholar] [CrossRef] [PubMed]
  121. Spiegelman, B.M.; Farmer, S.R. Decreases in tubulin and actin gene expression prior to morphological differentiation of 3T3 adipocytes. Cell 1982, 29, 53–60. [Google Scholar] [CrossRef]
  122. Shoham, N.; Gefen, A. The influence of mechanical stretching on mitosis, growth, and adipose conversion in adipocyte cultures. Biomech. Model. Mechanobiol. 2012, 11, 1029–1045. [Google Scholar] [CrossRef] [PubMed]
  123. Shoham, N.; Girshovitz, P.; Katzengold, R.; Shaked, N.T.; Benayahu, D.; Gefen, A. Adipocyte stiffness increases with accumulation of lipid droplets. Biophys. J. 2014, 106, 1421–1431. [Google Scholar] [CrossRef] [PubMed] [Green Version]
  124. Le Lay, S.; Briand, N.; Dugail, I. Adipocyte size fluctuation, mechano-active lipid droplets and caveolae. Adipocyte 2014, 4, 158–160. [Google Scholar] [CrossRef] [PubMed] [Green Version]
  125. Sinha, B.; Köster, D.; Ruez, R.; Gonnord, P.; Bastiani, M.; Abankwa, D.; Stan, R.V.; Butler-Browne, G.; Vedie, B.; Johannes, L.; et al. Cells respond to mechanical stress by rapid disassembly of caveolae. Cell 2011, 144, 402–413. [Google Scholar] [CrossRef] [Green Version]
  126. Scherer, P.E.; Lisanti, M.P.; Baldini, G.; Sargiacomo, M.; Mastick, C.C.; Lodish, H.F. Induction of caveolin during adipogenesis and association of GLUT4 with caveolin-rich vesicles. J. Cell Biol. 1994, 127, 1233–1243. [Google Scholar] [CrossRef] [PubMed]
  127. Blouin, C.M.; Le Lay, S.; Lasnier, F.; Dugail, I.; Hajduch, E. Regulated association of caveolins to lipid droplets during differentiation of 3T3-L1 adipocytes. Biochem. Biophys. Res. Commun. 2008, 376, 331–335. [Google Scholar] [CrossRef] [PubMed] [Green Version]
  128. Briand, N.; Prado, C.; Mabilleau, G.; Lasnier, F.; Le Lièpvre, X.; Covington, J.D.; Ravussin, E.; Le Lay, S.; Dugail, I. Caveolin-1 expression and cavin stability regulate caveolae dynamics in adipocyte lipid store fluctuation. Diabetes 2014, 63, 4032–4044. [Google Scholar] [CrossRef] [Green Version]
  129. Kim, C.A.; Delépine, M.; Boutet, E.; El Mourabit, H.; Le Lay, S.; Meier, M.; Nemani, M.; Bridel, E.; Leite, C.C.; Bertola, D.R.; et al. Association of a homozygous nonsense caveolin-1 mutation with Berardinelli-Seip congenital lipodystrophy. J. Clin. Endocrinol. Metab. 2008, 93, 1129–1134. [Google Scholar] [CrossRef] [Green Version]
  130. Mihai, L.A.; Chin, L.K.; Janmey, P.A.; Goriely, A. A comparison of hyperelastic constitutive models applicable to brain and fat tissues. J. R. Soc. Interface 2015, 12, 20150486. [Google Scholar] [CrossRef]
  131. Sasso, M.; Liu, Y.; Aron-Wisnewsky, J.; Bouillot, J.L.; Abdennour, M.; Clet, M.; Sandrin, L.; le Naour, G.; Bedossa, P.; Tordjman, J.; et al. AdipoScan: A Novel Transient Elastography-Based Tool Used to Non-Invasively Assess Subcutaneous Adipose Tissue Shear Wave Speed in Obesity. Ultrasound Med. Biol. 2016, 42, 2401–2413. [Google Scholar] [CrossRef]
  132. Wenderott, J.K.; Flesher, C.G.; Baker, N.A.; Neeley, C.K.; Varban, O.A.; Lumeng, C.N.; Muhammad, L.N.; Yeh, C.; Green, P.F.; O’Rourke, R.W. Elucidating nanoscale mechanical properties of diabetic human adipose tissue using atomic force microsco-py. Sci. Rep. 2020, 10, 20423. [Google Scholar] [CrossRef]
  133. Divoux, A.; Tordjman, J.; Veyrie, N.; Hugol, D.; Poitou, C.; Aissat, A.; Basdevant, A.; Zucker, J.; Bedossa, P.; Cle, K. Fibrosis in Human Adipose Tissue: Composition, Distribution, and Link with Lipid Metabolism and Fat. Diabetes 2010, 59, 2817–2825. [Google Scholar] [CrossRef] [PubMed] [Green Version]
  134. Unamuno, X.; Gómez-Ambrosi, J.; Becerril, S.; Álvarez-Cienfuegos, F.J.; Ramírez, B.; Rodríguez, A.; Ezquerro, S.; Valentí, V.; Moncada, R.; Mentxaka, A.; et al. Changes in mechanical properties of adipose tissue after bariatric surgery driven by extracellular matrix remodelling and neovascularization are associated with metabolic improvements. Acta Biomater. 2022, 141, 264–279. [Google Scholar] [CrossRef] [PubMed]
  135. Pellegrinelli, V.; Heuvingh, J.; Du Roure, O.; Rouault, C.; Devulder, A.; Klein, C.; Lacasa, M.; Clément, E.; Lacasa, D.; Clément, K. Human adipocyte function is impacted by mechanical cues. J. Pathol. 2014, 233, 183–195. [Google Scholar] [CrossRef] [PubMed]
  136. Cristancho, A.G.; Lazar, M.A. Forming functional fat: A growing understanding of adipocyte differentiation. Nat. Rev. Mol. Cell Biol. 2011, 12, 722–734. [Google Scholar] [CrossRef] [PubMed]
  137. Discher, D.E.; Janmey, P.; Wang, Y.L. Tissue cells feel and respond to the stiffness of their substrate. Science 2005, 310, 1139–1143. [Google Scholar] [CrossRef] [PubMed] [Green Version]
  138. Engler, A.J.; Sen, S.; Sweeney, H.L.; Discher, D.E. Matrix elasticity directs stem cell lineage specification. Cell 2006, 126, 677–689. [Google Scholar] [CrossRef] [PubMed] [Green Version]
  139. Martino, F.; Perestrelo, A.R.; Vinarský, V.; Pagliari, S.; Forte, G. Cellular Mechanotransduction: From Tension to Function. Front. Physiol. 2018, 9, 824. [Google Scholar] [CrossRef]
  140. Pan, H.; Xie, Y.; Zhang, Z.; Li, K.; Hu, D.; Zheng, X.; Fan, Q.; Tang, T. YAP-mediated mechanotransduction regulates osteogenic and adipogenic differentiation of BMSCs on hierarchical structure. Colloids Surf. B. Biointerfaces 2017, 152, 344–353. [Google Scholar] [CrossRef]
  141. McBeath, R.; Pirone, D.M.; Nelson, C.M.; Bhadriraju, K.; Chen, C.S. Cell shape, cytoskeletal tension, and RhoA regulate stem cell lineage commitment. Dev. Cell 2004, 6, 483–495. [Google Scholar] [CrossRef] [Green Version]
  142. Green, H.; Meuth, M. An established pre-adipose cell line and its differentiation in culture. Cell 1974, 3, 127–133. [Google Scholar] [CrossRef]
  143. Kuri-Harcuch, W.; Green, H. Adipose conversion of 3T3 cells depends on a serum factor. Proc. Natl. Acad. Sci. USA 1978, 75, 6107–6109. [Google Scholar] [CrossRef] [Green Version]
  144. Pairault, J.; Green, H. A study of the adipose conversion of suspended 3T3 cells by using glycerophosphate dehydrogenase as differentiation marker. Proc. Natl. Acad. Sci. USA 1979, 76, 5138–5142. [Google Scholar] [CrossRef] [PubMed] [Green Version]
  145. Green, H.; Kehinde, O. An established preadipose cell line and its differentiation in culture. II. Factors affecting the adipose conversion. Cell 1975, 5, 19–27. [Google Scholar] [CrossRef]
  146. Grigoriadis, A.E.; Heersche, J.N.M.; Aubin, J.E. Differentiation of muscle, fat, cartilage, and bone from progenitor cells present in a bone-derived clonal cell population: Effect of dexamethasone. J. Cell Biol. 1988, 106, 2139–2151. [Google Scholar] [CrossRef] [PubMed]
  147. Khetan, S.; Guvendiren, M.; Legant, W.R.; Cohen, D.M.; Chen, C.S.; Burdick, J.A. Degradation-mediated cellular traction directs stem cell fate in covalently crosslinked three-dimensional hydrogels. Nat. Mater. 2013, 12, 458–465. [Google Scholar] [CrossRef] [PubMed] [Green Version]
  148. Kilian, K.A.; Bugarija, B.; Lahn, B.T.; Mrksich, M. Geometric cues for directing the differentiation of mesenchymal stem cells. Proc. Natl. Acad. Sci. USA 2010, 107, 4872–4877. [Google Scholar] [CrossRef] [Green Version]
  149. Oliver-De La Cruz, J.; Nardone, G.; Vrbsky, J.; Pompeiano, A.; Perestrelo, A.R.; Capradossi, F.; Melajová, K.; Filipensky, P.; Forte, G. Substrate mechanics controls adipogenesis through YAP phosphorylation by dictating cell spreading. Biomaterials 2019, 205, 64–80. [Google Scholar] [CrossRef]
  150. Young, D.A.; Choi, Y.S.; Engler, A.J.; Christman, K.L. Stimulation of adipogenesis of adult adipose-derived stem cells using substrates that mimic the stiffness of adipose tissue. Biomaterials 2013, 34, 8581–8588. [Google Scholar] [CrossRef] [Green Version]
  151. Levy, A.; Enzer, S.; Shoham, N.; Zaretsky, U.; Gefen, A. Large, but not small sustained tensile strains stimulate adipogenesis in culture. Ann. Biomed. Eng. 2012, 40, 1052–1060. [Google Scholar] [CrossRef]
  152. Chun, T.H.; Hotary, K.B.; Sabeh, F.; Saltiel, A.R.; Allen, E.D.; Weiss, S.J. A pericellular collagenase directs the 3-dimensional development of white adipose tissue. Cell 2006, 125, 577–591. [Google Scholar] [CrossRef] [PubMed] [Green Version]
  153. Ibrahimi, A.; Bonino, F.; Bardon, S.; Ailhaud, G.; Dani, C. Essential role of collagens for terminal differentiation of preadipocytes. Biochem. Biophys. Res. Commun. 1992, 187, 1314–1322. [Google Scholar] [CrossRef]
  154. Zöller, N.; Schreiner, S.; Petry, L.; Hoffmann, S.; Steinhorst, K.; Kleemann, J.; Jäger, M.; Kaufmann, R.; Meissner, M.; Kippenberger, S. Collagen I Promotes Adipocytogenesis in Adipose-Derived Stem Cells In Vitro. Cells 2019, 8, 302. [Google Scholar] [CrossRef] [Green Version]
  155. Mauney, J.; Volloch, V. Human bone marrow-derived stromal cells show highly efficient stress-resistant adipogenesis on denatured collagen IV matrix but not on its native counterpart: Implications for obesity. Matrix Biol. 2010, 29, 9–14. [Google Scholar] [CrossRef] [PubMed]
  156. Chavey, C.; Mari, B.; Monthouel, M.N.; Bonnafous, S.; Anglard, P.; Van Obberghen, E.; Tartare-Deckert, S. Matrix metalloproteinases are differentially expressed in adipose tissue during obesity and modulate adipocyte differentiation. J. Biol. Chem. 2003, 278, 11888–11896. [Google Scholar] [CrossRef] [PubMed] [Green Version]
  157. Maquoi, E.; Munaut, C.; Colige, A.; Collen, D.; Roger Lijnen, H. Modulation of adipose tissue expression of murine matrix metalloproteinases and their tissue inhibitors with obesity. Diabetes 2002, 51, 1093–1101. [Google Scholar] [CrossRef] [PubMed] [Green Version]
  158. Derosa, G.; Ferrari, I.; D’Angelo, A.; Tinelli, C.; Salvadeo, S.A.T.; Ciccarelli, L.; Piccinni, M.N.; Gravina, A.; Ramondetti, F.; Maffioli, P.; et al. Matrix metalloproteinase-2 and -9 levels in obese patients. Endothelium 2008, 15, 219–224. [Google Scholar] [CrossRef] [PubMed]
  159. Chun, T.H.; Inoue, M.; Morisaki, H.; Yamanaka, I.; Miyamoto, Y.; Okamura, T.; Sato-Kusubata, K.; Weiss, S.J. Genetic link between obesity and MMP14-dependent adipogenic collagen turnover. Diabetes 2010, 59, 2484–2494. [Google Scholar] [CrossRef] [Green Version]
  160. Spiegelman, B.M.; Ginty, C.A. Fibronectin modulation of cell shape and lipogenic gene expression in 3T3-adipocytes. Cell 1983, 35, 657–666. [Google Scholar] [CrossRef]
  161. Nie, J.; Helene Sage, E. SPARC inhibits adipogenesis by its enhancement of beta-catenin signaling. J. Biol. Chem. 2009, 284, 1279–1290. [Google Scholar] [CrossRef] [PubMed] [Green Version]
  162. Liu, J.; DeYoung, S.M.; Zhang, M.; Zhang, M.; Cheng, A.; Saltiel, A.R. Changes in integrin expression during adipocyte differentiation. Cell Metab. 2005, 2, 165–177. [Google Scholar] [CrossRef] [PubMed] [Green Version]
  163. Oguri, Y.; Shinoda, K.; Kim, H.; Alba, D.L.; Bolus, W.R.; Wang, Q.; Brown, Z.; Pradhan, R.N.; Tajima, K.; Yoneshiro, T.; et al. CD81 Controls Beige Fat Progenitor Cell Growth and Energy Balance via FAK Signaling. Cell 2020, 182, 563–577.e20. [Google Scholar] [CrossRef] [PubMed]
  164. Pedersen, S.F.; Kapus, A.; Hoffmann, E.K. Osmosensory mechanisms in cellular and systemic volume regulation. J. Am. Soc. Nephrol. 2011, 22, 1587–1597. [Google Scholar] [CrossRef] [PubMed] [Green Version]
  165. Friis, M.B.; Friborg, C.R.; Schneider, L.; Nielsen, M.B.; Lambert, I.H.; Christensen, S.T.; Hoffman, E.K. Cell shrinkage as a signal to apoptosis in NIH 3T3 fibroblasts. J. Physiol. 2005, 567, 427–443. [Google Scholar] [CrossRef] [PubMed]
  166. Eduardsen, K.; Larsen, S.L.; Novak, I.; Lambert, I.H.; Hoffmann, E.K.; Pedersen, S.F. Cell volume regulation and signaling in 3T3-L1 pre-adipocytes and adipocytes: On the possible roles of caveolae, insulin receptors, FAK and ERK1/2. Cell. Physiol. Biochem. 2011, 28, 1231–1246. [Google Scholar] [CrossRef]
  167. Nielsen, M.B.; Christensen, S.T.; Hoffmann, E.K. Effects of osmotic stress on the activity of MAPKs and PDGFR-beta-mediated signal transduction in NIH-3T3 fibroblasts. Am. J. Physiol. Cell Physiol. 2008, 294. [Google Scholar] [CrossRef]
  168. Baron, V.; Calléja, V.; Ferrari, P.; Alengrin, F.; Van Obberghen, E. p125Fak focal adhesion kinase is a substrate for the insulin and insulin-like growth factor-I tyrosine kinase receptors. J. Biol. Chem. 1998, 273, 7162–7168. [Google Scholar] [CrossRef] [Green Version]
  169. Tharp, K.M.; Kang, M.S.; Timblin, G.A.; Dempersmier, J.; Dempsey, G.E.; Zushin, P.J.H.; Benavides, J.; Choi, C.; Li, C.X.; Jha, A.K.; et al. Actomyosin-Mediated Tension Orchestrates Uncoupled Respiration in Adipose Tissues. Cell Metab. 2018, 27, 602–615.e4. [Google Scholar] [CrossRef] [Green Version]
  170. Seale, P.; Bjork, B.; Yang, W.; Kajimura, S.; Chin, S.; Kuang, S.; Scimè, A.; Devarakonda, S.; Conroe, H.M.; Erdjument-Bromage, H.; et al. PRDM16 controls a brown fat/skeletal muscle switch. Nature 2008, 454, 961–967. [Google Scholar] [CrossRef] [Green Version]
  171. Jansen, K.A.; Donato, D.M.; Balcioglu, H.E.; Schmidt, T.; Danen, E.H.J.; Koenderink, G.H. A guide to mechanobiology: Where biology and physics meet. Biochim. Biophys. Acta 2015, 1853, 3043–3052. [Google Scholar] [CrossRef] [Green Version]
  172. Kirby, T.J.; Lammerding, J. Emerging views of the nucleus as a cellular mechanosensor. Nat. Cell Biol. 2018, 20, 373–381. [Google Scholar] [CrossRef]
  173. Smas, C.M.; Sul, H.S. Control of adipocyte differentiation. Biochem. J. 1995, 309 Pt 3, 697–710. [Google Scholar] [CrossRef] [PubMed] [Green Version]
  174. Franke, W.W.; Hergt, M.; Grund, C. Rearrangement of the vimentin cytoskeleton during adipose conversion: Formation of an intermediate filament cage around lipid globules. Cell 1987, 49, 131–141. [Google Scholar] [CrossRef]
  175. Pairault, J.; Lasnier, F. Dihydrocytochalasin B promotes adipose conversion of 3T3 cells. Biol. Cell 1987, 61, 149–154. [Google Scholar] [CrossRef] [PubMed]
  176. Boone, C.; Gregoire, F.; De Clercq, L.; Remacle, C. The modulation of cell shape influences porcine preadipocyte differentiation. Vitr. Cell. Dev. Biol. Anim. 1999, 35, 61–63. [Google Scholar] [CrossRef] [PubMed]
  177. Rodríguez Fernández, J.L.; Ben-Ze’ev, A. Regulation of fibronectin, integrin and cytoskeleton expression in differentiating adipocytes: Inhibition by extracellular matrix and polylysine. Differentiation. 1989, 42, 65–74. [Google Scholar] [CrossRef] [PubMed]
  178. Schiller, Z.A.; Schiele, N.R.; Sims, J.K.; Lee, K.; Kuo, C.K. Adipogenesis of adipose-derived stem cells may be regulated via the cytoskeleton at physiological oxygen levels in vitro. Stem Cell Res. Ther. 2013, 4, 1–10. [Google Scholar] [CrossRef] [Green Version]
  179. Nobusue, H.; Onishi, N.; Shimizu, T.; Sugihara, E.; Oki, Y.; Sumikawa, Y.; Chiyoda, T.; Akashi, K.; Saya, H.; Kano, K. Regulation of MKL1 via actin cytoskeleton dynamics drives adipocyte differentiation. Nat. Commun. 2014, 5, 3368. [Google Scholar] [CrossRef] [Green Version]
  180. Dupont, S.; Morsut, L.; Aragona, M.; Enzo, E.; Giulitti, S.; Cordenonsi, M.; Zanconato, F.; Le Digabel, J.; Forcato, M.; Bicciato, S.; et al. Role of YAP/TAZ in mechanotransduction. Nature 2011, 474, 179–184. [Google Scholar] [CrossRef]
  181. Hara, Y.; Wakino, S.; Tanabe, Y.; Saito, M.; Tokuyama, H.; Washida, N.; Tatematsu, S.; Yoshioka, K.; Homma, K.; Hasegawa, K.; et al. Rho and Rho-kinase activity in adipocytes contributes to a vicious cycle in obesity that may involve mechanical stretch. Sci. Signal. 2011, 4, ra3. [Google Scholar] [CrossRef]
  182. Hansson, B.; Morén, B.; Fryklund, C.; Vliex, L.; Wasserstrom, S.; Albinsson, S.; Berger, K.; Stenkula, K.G. Adipose cell size changes are associated with a drastic actin remodeling. Sci. Rep. 2019, 9, 1–14. [Google Scholar] [CrossRef] [PubMed]
  183. Kim, J.I.; Park, J.; Ji, Y.; Jo, K.; Han, S.M.; Sohn, J.H.; Shin, K.C.; Han, J.S.; Jeon, Y.G.; Nahmgoong, H.; et al. During Adipocyte Remodeling, Lipid Droplet Configurations Regulate Insulin Sensitivity through F-Actin and G-Actin Reorganization. Mol. Cell. Biol. 2019, 39, e00210-19. [Google Scholar] [CrossRef] [PubMed] [Green Version]
  184. Hoffmeyer, K.; Junghans, D.; Kanzler, B.; Kemler, R. Trimethylation and Acetylation of β-Catenin at Lysine 49 Represent Key Elements in ESC Pluripotency. Cell Rep. 2017, 18, 2815–2824. [Google Scholar] [CrossRef] [PubMed] [Green Version]
  185. Kaelin, W.G.; McKnight, S.L. Influence of metabolism on epigenetics and disease. Cell 2013, 153, 56–69. [Google Scholar] [CrossRef] [PubMed] [Green Version]
  186. Reid, M.A.; Dai, Z.; Locasale, J.W. The impact of cellular metabolism on chromatin dynamics and epigenetics. Nat. Cell Biol. 2017, 19, 1298–1306. [Google Scholar] [CrossRef]
  187. Berger, S.L. The complex language of chromatin regulation during transcription. Nature 2007, 447, 407–412. [Google Scholar] [CrossRef]
  188. Lu, C.; Thompson, C.B. Metabolic regulation of epigenetics. Cell Metab. 2012, 16, 9–17. [Google Scholar] [CrossRef] [Green Version]
  189. Lecoutre, S.; Maqdasy, S.; Petrus, P.; Ludzki, A.; Couchet, M.; Mejhert, N.; Rydén, M. Glutamine metabolism in adipocytes: A bona fide epigenetic modulator of inflammation. Adipocyte 2020, 9, 620–625. [Google Scholar] [CrossRef]
  190. Petrus, P.; Lecoutre, S.; Dollet, L.; Wiel, C.; Sulen, A.; Gao, H.; Tavira, B.; Laurencikiene, J.; Rooyackers, O.; Checa, A.; et al. Glutamine Links Obesity to Inflammation in Human White Adipose Tissue. Cell Metab. 2020, 31, 375–390.e11. [Google Scholar] [CrossRef]
  191. Maqdasy, S.; Lecoutre, S.; Renzi, G.; Frendo-Cumbo, S.; Rizo-Roca, D.; Moritz, T.; Juvany, M.; Hodek, O.; Gao, H.; Couchet, M.; et al. Impaired phosphocreatine metabolism in white adipocytes promotes inflammation. Nat. Metab. 2022, 4, 190–202. [Google Scholar] [CrossRef]
  192. Romani, P.; Valcarcel-Jimenez, L.; Frezza, C.; Dupont, S. Crosstalk between mechanotransduction and metabolism. Nat. Rev. Mol. Cell Biol. 2021, 22, 22–38. [Google Scholar] [CrossRef] [PubMed]
  193. Zanotelli, M.R.; Zhang, J.; Reinhart-King, C.A. Mechanoresponsive metabolism in cancer cell migration and metastasis. Cell Metab. 2021, 33, 1307–1321. [Google Scholar] [CrossRef] [PubMed]
  194. Park, J.S.; Burckhardt, C.J.; Lazcano, R.; Solis, L.M.; Isogai, T.; Li, L.; Chen, C.S.; Gao, B.; Minna, J.D.; Bachoo, R.; et al. Mechanical regulation of glycolysis via cytoskeleton architecture. Nature 2020, 578, 621–626. [Google Scholar] [CrossRef] [PubMed]
  195. Bays, J.L.; DeMali, K.A. It takes energy to resist force. Cell Cycle 2017, 16, 1733–1734. [Google Scholar] [CrossRef] [Green Version]
  196. Salvi, A.M.; Bays, J.L.; Mackin, S.R.; Mege, R.M.; DeMali, K.A. Ankyrin G organizes membrane components to promote coupling of cell mechanics and glucose uptake. Nat. Cell Biol. 2021, 23, 457–466. [Google Scholar] [CrossRef]
  197. Torrino, S.; Bertero, T. Metabo-reciprocity in cell mechanics: Feeling the demands/feeding the demand. Trends Cell Biol. 2022, 32, 624–636. [Google Scholar] [CrossRef]
  198. Bertero, T.; Gaggioli, C. Mechanical forces rewire metabolism in the tumor niche. Mol. Cell. Oncol. 2019, 6, 1592945. [Google Scholar] [CrossRef]
  199. Bertero, T.; Oldham, W.M.; Grasset, E.M.; Bourget, I.; Boulter, E.; Pisano, S.; Hofman, P.; Bellvert, F.; Meneguzzi, G.; Bulavin, D.V.; et al. Tumor-Stroma Mechanics Coordinate Amino Acid Availability to Sustain Tumor Growth and Malignancy. Cell Metab. 2019, 29, 124–140.e10. [Google Scholar] [CrossRef] [Green Version]
  200. Debari, M.K.; Abbott, R.D. Adipose Tissue Fibrosis: Mechanisms, Models, and Importance. Int. J. Mol. Sci. 2020, 21, 6030. [Google Scholar] [CrossRef]
  201. Aebi, U.; Cohn, J.; Buhle, L.; Gerace, L. The nuclear lamina is a meshwork of intermediate-type filaments. Nature 1986, 323, 560–564. [Google Scholar] [CrossRef]
  202. Lund, E.; Collas, P. Nuclear lamins: Making contacts with promoters. Nucleus 2013, 4, 424–430. [Google Scholar] [CrossRef] [PubMed] [Green Version]
  203. Lund, E.; Oldenburg, A.R.; Delbarre, E.; Freberg, C.T.; Duband-Goulet, I.; Eskeland, R.; Buendia, B.; Collas, P. Lamin A/C-promoter interactions specify chromatin state-dependent transcription outcomes. Genome Res. 2013, 23, 1580–1589. [Google Scholar] [CrossRef] [PubMed] [Green Version]
  204. Lund, E.G.; Duband-Goulet, I.; Oldenburg, A.; Buendia, B.; Collas, P. Distinct features of lamin A-interacting chromatin domains mapped by ChIP-sequencing from sonicated or micrococcal nuclease-digested chromatin. Nucleus 2015, 6, 30–39. [Google Scholar] [CrossRef] [Green Version]
  205. Gesson, K.; Rescheneder, P.; Skoruppa, M.P.; Von Haeseler, A.; Dechat, T.; Foisner, R. A-type lamins bind both hetero- and euchromatin, the latter being regulated by lamina-associated polypeptide 2 alpha. Genome Res. 2016, 26, 462–473. [Google Scholar] [CrossRef] [Green Version]
  206. Lee, J.S.H.; Hale, C.M.; Panorchan, P.; Khatau, S.B.; George, J.P.; Tseng, Y.; Stewart, C.L.; Hodzic, D.; Wirtz, D. Nuclear lamin A/C deficiency induces defects in cell mechanics, polarization, and migration. Biophys. J. 2007, 93, 2542–2552. [Google Scholar] [CrossRef] [PubMed] [Green Version]
  207. Hynes, R.O. Integrins: Bidirectional, allosteric signaling machines. Cell 2002, 110, 673–687. [Google Scholar] [CrossRef] [Green Version]
  208. Nmezi, B.; Xu, J.; Fu, R.; Armiger, T.J.; Rodriguez-Bey, G.; Powell, J.S.; Ma, H.; Sullivan, M.; Tu, Y.; Chen, N.Y.; et al. Concentric organization of A- and B-type lamins predicts their distinct roles in the spatial organization and stability of the nuclear lamina. Proc. Natl. Acad. Sci. USA 2019, 116, 4307–4315. [Google Scholar] [CrossRef] [Green Version]
  209. Verstraeten, V.L.R.M.; Renes, J.; Ramaekers, F.C.S.; Kamps, M.; Kuijpers, H.J.; Verheyen, F.; Wabitsch, M.; Steijlen, P.M.; Van Steensel, M.A.M.; Broers, J.L.V. Reorganization of the nuclear lamina and cytoskeleton in adipogenesis. Histochem. Cell Biol. 2011, 135, 251–261. [Google Scholar] [CrossRef] [Green Version]
  210. Vigouroux, C.; Caron-Debarle, M.; Le Dour, C.; Magré, J.; Capeau, J. Molecular mechanisms of human lipodystrophies: From adipocyte lipid droplet to oxidative stress and lipotoxicity. Int. J. Biochem. Cell Biol. 2011, 43, 862–876. [Google Scholar] [CrossRef]
  211. Crisp, M.; Liu, Q.; Roux, K.; Rattner, J.B.; Shanahan, C.; Burke, B.; Stahl, P.D.; Hodzic, D. Coupling of the nucleus and cytoplasm: Role of the LINC complex. J. Cell Biol. 2006, 172, 41–53. [Google Scholar] [CrossRef] [PubMed] [Green Version]
  212. Tapley, E.C.; Starr, D.A. Connecting the nucleus to the cytoskeleton by SUN-KASH bridges across the nuclear envelope. Curr. Opin. Cell Biol. 2013, 25, 57–62. [Google Scholar] [CrossRef] [PubMed] [Green Version]
  213. Méjat, A.; Misteli, T. LINC complexes in health and disease. Nucleus 2010, 1, 40–52. [Google Scholar] [CrossRef] [PubMed] [Green Version]
  214. Maniotis, A.J.; Chen, C.S.; Ingber, D.E. Demonstration of mechanical connections between integrins, cytoskeletal filaments, and nucleoplasm that stabilize nuclear structure. Proc. Natl. Acad. Sci. USA 1997, 94, 849–854. [Google Scholar] [CrossRef] [PubMed] [Green Version]
  215. Killaars, A.R.; Grim, J.C.; Walker, C.J.; Hushka, E.A.; Brown, T.E.; Anseth, K.S. Extended Exposure to Stiff Microenvironments Leads to Persistent Chromatin Remodeling in Human Mesenchymal Stem Cells. Adv. Sci. 2018, 6, 670. [Google Scholar] [CrossRef] [PubMed] [Green Version]
  216. Downing, T.L.; Soto, J.; Morez, C.; Houssin, T.; Fritz, A.; Yuan, F.; Chu, J.; Patel, S.; Schaffer, D.V.; Li, S. Biophysical regulation of epigenetic state and cell reprogramming. Nat. Mater. 2013, 12, 1154–1162. [Google Scholar] [CrossRef] [PubMed]
  217. Killaars, A.R.; Walker, C.J.; Anseth, K.S. Nuclear mechanosensing controls MSC osteogenic potential through HDAC epigenetic remodeling. Proc. Natl. Acad. Sci. USA 2020, 117, 21258–21266. [Google Scholar] [CrossRef] [PubMed]
  218. Swift, J.; Ivanovska, I.L.; Buxboim, A.; Harada, T.; Dingal, P.C.D.P.; Pinter, J.; Pajerowski, J.D.; Spinler, K.R.; Shin, J.W.; Tewari, M.; et al. Nuclear lamin-A scales with tissue stiffness and enhances matrix-directed differentiation. Science 2013, 341. [Google Scholar] [CrossRef] [PubMed] [Green Version]
  219. Alcorta-Sevillano, N.; Macías, I.; Rodríguez, C.I.; Infante, A. Crucial Role of Lamin A/C in the Migration and Differentiation of MSCs in Bone. Cells 2020, 9, 1330. [Google Scholar] [CrossRef] [PubMed]
  220. Crowder, S.W.; Leonardo, V.; Whittaker, T.; Papathanasiou, P.; Stevens, M.M. Material Cues as Potent Regulators of Epigenetics and Stem Cell Function. Cell Stem Cell 2016, 18, 39–52. [Google Scholar] [CrossRef] [PubMed] [Green Version]
  221. Heo, K.S.; Berk, B.C.; Abe, J.I. Disturbed Flow-Induced Endothelial Proatherogenic Signaling Via Regulating Post-Translational Modifications and Epigenetic Events. Antioxid. Redox Signal. 2016, 25, 435–450. [Google Scholar] [CrossRef] [Green Version]
  222. Li, Y.; Chu, J.S.; Kurpinski, K.; Li, X.; Bautista, D.M.; Yang, L.; Paul Sung, K.L.; Li, S. Biophysical regulation of histone acetylation in mesenchymal stem cells. Biophys. J. 2011, 100, 1902–1909. [Google Scholar] [CrossRef] [PubMed] [Green Version]
  223. Jain, N.; Iyer, K.V.; Kumar, A.; Shivashankar, G.V. Cell geometric constraints induce modular gene-expression patterns via redistribution of HDAC3 regulated by actomyosin contractility. Proc. Natl. Acad. Sci. USA 2013, 110, 11349–11354. [Google Scholar] [CrossRef] [Green Version]
  224. Valenzuela-Fernández, A.; Cabrero, J.R.; Serrador, J.M.; Sánchez-Madrid, F. HDAC6: A key regulator of cytoskeleton, cell migration and cell-cell interactions. Trends Cell Biol. 2008, 18, 291–297. [Google Scholar] [CrossRef] [PubMed]
  225. Lecoutre, S.; Petrus, P.; Rydén, M.; Breton, C. Transgenerational Epigenetic Mechanisms in Adipose Tissue Development. Trends Endocrinol. Metab. 2018, 29, 675–685. [Google Scholar] [CrossRef] [PubMed]
  226. Panciera, T.; Azzolin, L.; Cordenonsi, M.; Piccolo, S. Mechanobiology of YAP and TAZ in physiology and disease. Nat. Rev. Mol. Cell Biol. 2017, 18, 758–770. [Google Scholar] [CrossRef]
  227. Heng, B.C.; Zhang, X.; Aubel, D.; Bai, Y.; Li, X.; Wei, Y.; Fussenegger, M.; Deng, X. Role of YAP/TAZ in Cell Lineage Fate Determination and Related Signaling Pathways. Front. Cell Dev. Biol. 2020, 8, 735. [Google Scholar] [CrossRef]
  228. Moon, S.; Kim, W.; Kim, S.; Kim, Y.; Song, Y.; Bilousov, O.; Kim, J.; Lee, T.; Cha, B.; Kim, M.; et al. Phosphorylation by NLK inhibits YAP-14-3-3-interactions and induces its nuclear localization. EMBO Rep. 2017, 18, 61–71. [Google Scholar] [CrossRef] [Green Version]
  229. Nardone, G.; Oliver-De La Cruz, J.; Vrbsky, J.; Martini, C.; Pribyl, J.; Skládal, P.; Pešl, M.; Caluori, G.; Pagliari, S.; Martino, F.; et al. YAP regulates cell mechanics by controlling focal adhesion assembly. Nat. Commun. 2017, 8, 15321. [Google Scholar] [CrossRef]
  230. Mosqueira, D.; Pagliari, S.; Uto, K.; Ebara, M.; Romanazzo, S.; Escobedo-Lucea, C.; Nakanishi, J.; Taniguchi, A.; Franzese, O.; Di Nardo, P.; et al. Hippo pathway effectors control cardiac progenitor cell fate by acting as dynamic sensors of substrate mechanics and nanostructure. ACS Nano 2014, 8, 2033–2047. [Google Scholar] [CrossRef] [Green Version]
  231. Aragona, M.; Panciera, T.; Manfrin, A.; Giulitti, S.; Michielin, F.; Elvassore, N.; Dupont, S.; Piccolo, S. A mechanical checkpoint controls multicellular growth through YAP/TAZ regulation by actin-processing factors. Cell 2013, 154, 1047–1059. [Google Scholar] [CrossRef] [PubMed] [Green Version]
  232. Zhong, W.; Tian, K.; Zheng, X.; Li, L.; Zhang, W.; Wang, S.; Qin, J. Mesenchymal stem cell and chondrocyte fates in a multishear microdevice are regulated by Yes-associated protein. Stem Cells Dev. 2013, 22, 2083–2093. [Google Scholar] [CrossRef] [PubMed] [Green Version]
  233. An, Y.; Kang, Q.; Zhao, Y.; Hu, X.; Li, N. Lats2 modulates adipocyte proliferation and differentiation via hippo signaling. PLoS ONE 2013, 8, e72042. [Google Scholar]
  234. Pan, J.X.; Xiong, L.; Zhao, K.; Zeng, P.; Wang, B.; Tang, F.L.; Sun, D.; Guo, H.H.; Yang, X.; Cui, S.; et al. YAP promotes osteogenesis and suppresses adipogenic differentiation by regulating β-catenin signaling. Bone Res. 2018, 6, 18. [Google Scholar] [CrossRef] [Green Version]
  235. Kamura, K.; Shin, J.; Kiyonari, H.; Abe, T.; Shioi, G.; Fukuhara, A.; Sasaki, H. Obesity in Yap transgenic mice is associated with TAZ downregulation. Biochem. Biophys. Res. Commun. 2018, 505, 951–957. [Google Scholar] [CrossRef] [PubMed]
  236. Talele, N.P.; Fradette, J.; Davies, J.E.; Kapus, A.; Hinz, B. Expression of α-Smooth Muscle Actin Determines the Fate of Mesenchymal Stromal Cells. Stem Cell Rep. 2015, 4, 1016–1030. [Google Scholar] [CrossRef] [PubMed] [Green Version]
  237. Wang, L.; Wang, S.P.; Shi, Y.; Li, R.; Günther, S.; Ong, Y.T.; Potente, M.; Yuan, Z.; Liu, E.; Offermanns, S. YAP and TAZ protect against white adipocyte cell death during obesity. Nat. Commun. 2020, 11, 5455. [Google Scholar] [CrossRef] [PubMed]
  238. El Ouarrat, D.; Isaac, R.; Lee, Y.S.; Oh, D.Y.; Wollam, J.; Lackey, D.; Riopel, M.; Bandyopadhyay, G.; Seo, J.B.; Sampath-Kumar, R.; et al. TAZ Is a Negative Regulator of PPARγ Activity in Adipocytes and TAZ Deletion Improves Insulin Sensitivity and Glucose Tolerance. Cell Metab. 2020, 31, 162–173.e5. [Google Scholar] [CrossRef] [PubMed]
  239. Hong, J.H.; Hwang, E.S.; McManus, M.T.; Amsterdam, A.; Tian, Y.; Kalmukova, R.; Mueller, E.; Benjamin, T.; Spiegelman, B.M.; Sharp, P.A.; et al. TAZ, a transcriptional modulator of mesenchymal stem cell differentiation. Science 2005, 309, 1074–1078. [Google Scholar] [CrossRef] [PubMed] [Green Version]
  240. Kawano, S.; Maruyama, J.; Nagashima, S.; Inami, K.; Qiu, W.; Iwasa, H.; Nakagawa, K.; Ishigami-Yuasa, M.; Kagechika, H.; Nishina, H.; et al. A cell-based screening for TAZ activators identifies ethacridine, a widely used antiseptic and abortifacient, as a compound that promotes dephosphorylation of TAZ and inhibits adipogenesis in C3H10T1/2 cells. J. Biochem. 2015, 158, 413–423. [Google Scholar] [CrossRef] [PubMed]
  241. Douguet, D.; Honoré, E. Mammalian Mechanoelectrical Transduction: Structure and Function of Force-Gated Ion Channels. Cell 2019, 179, 340–354. [Google Scholar] [CrossRef] [PubMed]
  242. Zhang, Y.; Xie, L.; Gunasekar, S.K.; Tong, D.; Mishra, A.; Gibson, W.J.; Wang, C.; Fidler, T.; Marthaler, B.; Klingelhutz, A.; et al. SWELL1 is a regulator of adipocyte size, insulin signalling and glucose homeostasis. Nat. Cell Biol. 2017, 19, 504–517. [Google Scholar] [CrossRef] [PubMed] [Green Version]
  243. Wang, S.; Cao, S.; Arhatte, M.; Li, D.; Shi, Y.; Kurz, S.; Hu, J.; Wang, L.; Shao, J.; Atzberger, A.; et al. Adipocyte Piezo1 mediates obesogenic adipogenesis through the FGF1/FGFR1 signaling pathway in mice. Nat. Commun. 2020, 11, 2303. [Google Scholar] [CrossRef] [PubMed]
  244. Kwok, K.H.M.; Lam, K.S.L.; Xu, A. Heterogeneity of white adipose tissue: Molecular basis and clinical implications. Exp. Mol. Med. 2016, 48, e215. [Google Scholar] [CrossRef] [PubMed] [Green Version]
  245. Zwick, R.K.; Guerrero-Juarez, C.F.; Horsley, V.; Plikus, M.V. Anatomical, Physiological, and Functional Diversity of Adipose Tissue. Cell Metab. 2018, 27, 68–83. [Google Scholar] [CrossRef] [PubMed] [Green Version]
  246. Kajimura, S.; Spiegelman, B.M.; Seale, P. Brown and Beige Fat: Physiological Roles beyond Heat Generation. Cell Metab. 2015, 22, 546–559. [Google Scholar] [CrossRef] [PubMed] [Green Version]
  247. Tang, Q.Q.; Lane, M.D. Adipogenesis: From stem cell to adipocyte. Annu. Rev. Biochem. 2012, 81, 715–736. [Google Scholar] [CrossRef] [Green Version]
  248. Han, J.; Lee, J.-E.; Jin, J.; Lim, J.S.; Oh, N.; Kim, K.; Chang, S.-I.; Shibuya, M.; Kim, H.; Koh, G.Y. The spatiotemporal development of adipose tissue. Development 2011, 138, 5027–5037. [Google Scholar] [CrossRef] [Green Version]
  249. Lemonnier, D. Effect of age, sex, and sites on the cellularity of the adipose tissue in mice and rats rendered obese by a high-fat diet. J. Clin. Investig. 1972, 51, 2907–2915. [Google Scholar] [CrossRef] [Green Version]
  250. Knittle, J.L.; Timmers, K.; Ginsberg-Fellner, F.; Brown, R.E.; Katz, D.P. The growth of adipose tissue in children and adolescents. Cross-sectional and longitudinal studies of adipose cell number and size. J. Clin. Investig. 1979, 63, 239–246. [Google Scholar] [CrossRef] [Green Version]
  251. Kirtland, J.; Harris, P.M. Changes in adipose tissue of the rat due early undernutrition followed by rehabilitation. 3. Changes in cell replication studied with tritiated thymidine. Br. J. Nutr. 1980, 43, 33–43. [Google Scholar] [CrossRef] [Green Version]
  252. Birsoy, K.; Berry, R.; Wang, T.; Ceyhan, O.; Tavazoie, S.; Friedman, J.M.; Rodeheffer, M.S. Analysis of gene networks in white adipose tissue development reveals a role for ETS2 in adipogenesis. Development 2011, 138, 4709–4719. [Google Scholar] [CrossRef] [PubMed] [Green Version]
  253. Poissonnet, C.M.; Burdi, A.R.; Garn, S.M. The chronology of adipose tissue appearance and distribution in the human fetus. Early Hum. Dev. 1984, 10, 1–11. [Google Scholar] [CrossRef]
  254. Crandall, D.L.; Hausman, G.J.; Kral, J.G. A review of the microcirculation of adipose tissue: Anatomic, metabolic, and angiogenic perspectives. Microcirculation 1997, 4, 211–232. [Google Scholar] [CrossRef]
  255. Cannon, B.; Nedergaard, J. Brown adipose tissue: Function and physiological significance. Physiol. Rev. 2004, 84, 277–359. [Google Scholar] [CrossRef] [PubMed]
  256. Hepler, C.; Vishvanath, L.; Gupta, R.K. Sorting out adipocyte precursors and their role in physiology and disease. Genes Dev. 2017, 31, 127–140. [Google Scholar] [CrossRef] [PubMed] [Green Version]
  257. Burl, R.B.; Ramseyer, V.D.; Rondini, E.A.; Pique-Regi, R.; Lee, Y.H.; Granneman, J.G. Deconstructing Adipogenesis Induced by β3-Adrenergic Receptor Activation with Single-Cell Expression Profiling. Cell Metab. 2018, 28, 300–309.e4. [Google Scholar] [CrossRef] [Green Version]
  258. Merrick, D.; Sakers, A.; Irgebay, Z.; Okada, C.; Calvert, C.; Morley, M.P.; Percec, I.; Seale, P. Identification of a mesenchymal progenitor cell hierarchy in adipose tissue. Science 2019, 364, eaav2501. [Google Scholar] [CrossRef]
  259. Nahmgoong, H.; Jeon, Y.G.; Park, E.S.; Choi, Y.H.; Han, S.M.; Park, J.; Ji, Y.; Sohn, J.H.; Han, J.S.; Kim, Y.Y.; et al. Distinct properties of adipose stem cell subpopulations determine fat depot-specific characteristics. Cell Metab. 2022, 34, 458–472.e6. [Google Scholar] [CrossRef]
  260. Schwalie, P.C.; Dong, H.; Zachara, M.; Russeil, J.; Alpern, D.; Akchiche, N.; Caprara, C.; Sun, W.; Schlaudraff, K.U.; Soldati, G.; et al. A stromal cell population that inhibits adipogenesis in mammalian fat depots. Nature 2018, 559, 103–108. [Google Scholar] [CrossRef]
  261. Dong, H.; Sun, W.; Shen, Y.; Baláz, M.; Balázová, L.; Ding, L.; Löffler, M.; Hamilton, B.; Klöting, N.; Blüher, M.; et al. Identification of a regulatory pathway inhibiting adipogenesis via RSPO2. Nat. Metab. 2022, 4, 90–105. [Google Scholar] [CrossRef]
  262. Sárvári, A.K.; Van Hauwaert, E.L.; Markussen, L.K.; Gammelmark, E.; Marcher, A.B.; Ebbesen, M.F.; Nielsen, R.; Brewer, J.R.; Madsen, J.G.S.; Mandrup, S. Plasticity of Epididymal Adipose Tissue in Response to Diet-Induced Obesity at Single-Nucleus Resolution. Cell Metab. 2021, 33, 437–453.e5. [Google Scholar] [CrossRef] [PubMed]
  263. Vijay, J.; Gauthier, M.F.; Biswell, R.L.; Louiselle, D.A.; Johnston, J.J.; Cheung, W.A.; Belden, B.; Pramatarova, A.; Biertho, L.; Gibson, M.; et al. Single-cell analysis of human adipose tissue identifies depot and disease specific cell types. Nat. Metab. 2020, 2, 97–109. [Google Scholar] [CrossRef] [PubMed]
  264. Emont, M.P.; Jacobs, C.; Essene, A.L.; Pant, D.; Tenen, D.; Colleluori, G.; Di Vincenzo, A.; Jørgensen, A.M.; Dashti, H.; Stefek, A.; et al. A single-cell atlas of human and mouse white adipose tissue. Nature 2022, 603, 926–933. [Google Scholar] [CrossRef]
  265. Shan, B.; Barker, C.S.; Shao, M.; Zhang, Q.; Gupta, R.K.; Wu, Y. Multilayered omics reveal sex- and depot-dependent adipose progenitor cell heterogeneity. Cell Metab. 2022, 34, 783–799.e7. [Google Scholar] [CrossRef] [PubMed]
  266. Krueger, K.C.; Costa, M.J.; Du, H.; Feldman, B.J. Characterization of Cre recombinase activity for in vivo targeting of adipocyte precursor cells. Stem Cell Rep. 2014, 3, 1147–1158. [Google Scholar] [CrossRef]
  267. Sanchez-gurmaches, J.; Hung, C.; Guertin, D.A. Emerging Complexities in Adipocyte Origins and Identity. Trends Cell Biol. 2016, 26, 313–326. [Google Scholar] [CrossRef] [Green Version]
  268. Sanchez-Gurmaches, J.; Guertin, D.A. Adipocyte lineages: Tracing back the origins of fat. Biochim. Biophys. Acta Mol. Basis Dis. 2014, 1842, 340–351. [Google Scholar] [CrossRef] [Green Version]
  269. Billon, N.; Iannarelli, P.; Monteiro, M.C.; Glavieux-Pardanaud, C.; Richardson, W.D.; Kessaris, N.; Dani, C.; Dupin, E. The generation of adipocytes by the neural crest. Development 2007, 134, 2283–2292. [Google Scholar] [CrossRef] [PubMed] [Green Version]
  270. Le Lievre, C.S.; Le Douarin, N.M. Mesenchymal derivatives of the neural crest: Analysis of chimaeric quail and chick embryos. Embryol. Exp. Morph 1975, 34, 125–154. [Google Scholar] [CrossRef]
  271. Chau, Y.; Bandiera, R.; Serrels, A.; Martínez-estrada, O.M.; Qing, W.; Lee, M.; Slight, J.; Thornburn, A.; Berry, R.; Mchaffie, S.; et al. Visceral and subcutaneous fat have different origins and evidence supports a mesothelial source. Nat. Cell Biol. 2014, 16, 367–375. [Google Scholar] [CrossRef] [PubMed] [Green Version]
  272. Westcott, G.P.; Emont, M.P.; Li, J.; Jacobs, C.; Tsai, L.; Rosen, E.D. Mesothelial cells are not a source of adipocytes in mice. Cell Rep. 2021, 36, 109388. [Google Scholar] [CrossRef]
  273. Atit, R.; Sgaier, S.K.; Mohamed, O.A.; Taketo, M.M.; Dufort, D.; Joyner, A.L.; Niswander, L.; Conlon, R.A. Beta-catenin activation is necessary and sufficient to specify the dorsal dermal fate in the mouse. Dev. Biol. 2006, 296, 164–176. [Google Scholar] [CrossRef] [PubMed] [Green Version]
  274. Zhang, L.; Avery, J.; Yin, A.; Singh, A.M.; Cliff, T.S.; Yin, H.; Dalton, S. Generation of Functional Brown Adipocytes from Human Pluripotent Stem Cells via Progression through a Paraxial Mesoderm State. Cell Stem Cell 2020, 27, 784–797.e11. [Google Scholar] [CrossRef] [PubMed]
  275. Cohen, P.; Kajimura, S. The cellular and functional complexity of thermogenic fat. Nat. Rev. Mol. Cell Biol. 2021, 22, 393–409. [Google Scholar] [CrossRef] [PubMed]
  276. Hausman, G.J.; Hentges, E.J.; Thomas, G.B. Differentiation of adipose tissue and muscle in hypophysectomized pig fetuses. J. Anim. Sci. 1987, 64, 1255–1261. [Google Scholar] [CrossRef] [PubMed] [Green Version]
  277. Hausman, G.J.; Thomas, G.B. Structural and histochemical aspects of perirenal adipose tissue in fetal pigs: Relationships between stromal-vascular characteristics and fat cell concentration and enzyme activity. J. Morphol. 1986, 190, 271–283. [Google Scholar] [CrossRef]
  278. Hausman, G.J.; Richardson, R.L. Adrenergic innervation of fetal pig adipose tissue. Histochemical and ultrastructural studies. Acta Anat. 1987, 130, 291–297. [Google Scholar] [CrossRef]
  279. Ailhaud, G.; Grimaldi, P.; Négrel, R. Cellular and molecular aspects of adipose tissue development. Annu. Rev. Nutr. 1992, 12, 207–233. [Google Scholar] [CrossRef] [PubMed]
  280. Poissonnet, C.M.; Burdi, A.R.; Bookstein, F.L. Growth and development of human adipose tissue during early gestation. Early Hum. Dev. 1983, 8, 1–11. [Google Scholar] [CrossRef] [Green Version]
  281. Lee, Y.H.; Mottillo, E.P.; Granneman, J.G. Adipose tissue plasticity from WAT to BAT and in between. Biochim. Biophys. Acta 2014, 1842, 358–369. [Google Scholar] [CrossRef] [Green Version]
  282. Carmeliet, P. Angiogenesis in life, disease and medicine. Nature 2005, 438, 932–936. [Google Scholar] [CrossRef] [PubMed]
  283. Tschumperlin, D.J.; Ligresti, G.; Hilscher, M.B.; Shah, V.H. Mechanosensing and fibrosis. J. Clin. Investig. 2018, 128, 74–84. [Google Scholar] [CrossRef] [PubMed] [Green Version]
  284. Na, S.; Collin, O.; Chowdhury, F.; Tay, B.; Ouyang, M.; Wang, Y.; Wang, N. Rapid signal transduction in living cells is a unique feature of mechanotransduction. Proc. Natl. Acad. Sci. USA 2008, 105, 6626–6631. [Google Scholar] [CrossRef] [Green Version]
  285. Sukharev, S.I.; Martinac, B.; Arshavsky, V.Y.; Kung, C. Two types of mechanosensitive channels in the Escherichia coli cell envelope: Solubilization and functional reconstitution. Biophys. J. 1993, 65, 177–183. [Google Scholar] [CrossRef] [Green Version]
  286. Lecuit, T.; Lenne, P.F.; Munro, E. Force generation, transmission, and integration during cell and tissue morphogenesis. Annu. Rev. Cell Dev. Biol. 2011, 27, 157–184. [Google Scholar] [CrossRef] [PubMed] [Green Version]
  287. Decaudain, A.; Vantyghem, M.C.; Guerci, B.; Hécart, A.C.; Auclair, M.; Reznik, Y.; Narbonne, H.; Ducluzeau, P.H.; Donadille, B.; Lebbé, C.; et al. New metabolic phenotypes in laminopathies: LMNA mutations in patients with severe metabolic syndrome. J. Clin. Endocrinol. Metab. 2007, 92, 4835–4844. [Google Scholar] [CrossRef] [Green Version]
  288. Patni, N.; Hatab, S.; Xing, C.; Zhou, Z.; Quittner, C.; Garg, A. A novel autosomal recessive lipodystrophy syndrome due to homozygous LMNA variant. J. Med. Genet. 2020, 57, 422–426. [Google Scholar] [CrossRef] [PubMed]
  289. Patni, N.; Li, X.; Adams-Huet, B.; Vasandani, C.; Gomez-Diaz, R.A.; Garg, A. Regional Body Fat Changes and Metabolic Complications in Children With Dunnigan Lipodystrophy-Causing LMNA Variants. J. Clin. Endocrinol. Metab. 2019, 104, 1099–1108. [Google Scholar] [CrossRef] [Green Version]
  290. Oldenburg, A.R.; Collas, P. Mapping Nuclear Lamin-Genome Interactions by Chromatin Immunoprecipitation of Nuclear Lamins. Methods Mol. Biol. 2016, 1411, 315–324. [Google Scholar] [PubMed]
  291. Oldenburg, A.; Briand, N.; Sørensen, A.L.; Cahyani, I.; Shah, A.; Moskaug, J.Ø.; Collas, P. A lipodystrophy-causing lamin A mutant alters conformation and epigenetic regulation of the anti-adipogenic MIR335 locus. J. Cell Biol. 2017, 216, 2731–2743. [Google Scholar] [CrossRef] [Green Version]
  292. Osmanagic-Myers, S.; Foisner, R. The structural and gene expression hypotheses in laminopathic diseases-not so different after all. Mol. Biol. Cell 2019, 30, 1786–1790. [Google Scholar] [CrossRef] [PubMed]
  293. Vigouroux, C.; Auclair, M.; Dubosclard, E.; Pouchelet, M.; Capeau, J.; Courvalin, J.C.; Buendia, B. Nuclear envelope disorganization in fibroblasts from lipodystrophic patients with heterozygous R482Q/W mutations in the lamin A/C gene. J. Cell Sci. 2001, 114, 4459–4468. [Google Scholar] [CrossRef] [PubMed]
  294. Paulsen, J.; Sekelja, M.; Oldenburg, A.R.; Barateau, A.; Briand, N.; Delbarre, E.; Shah, A.; Sørensen, A.L.; Vigouroux, C.; Buendia, B.; et al. Chrom3D: Three-dimensional genome modeling from Hi-C and nuclear lamin-genome contacts. Genome Biol. 2017, 18, 1–15. [Google Scholar] [CrossRef]
  295. Van Steensel, B.; Belmont, A.S. Lamina-Associated Domains: Links with Chromosome Architecture, Heterochromatin, and Gene Repression. Cell 2017, 169, 780–791. [Google Scholar] [CrossRef] [PubMed] [Green Version]
  296. Schuettengruber, B.; Chourrout, D.; Vervoort, M.; Leblanc, B.; Cavalli, G. Genome regulation by polycomb and trithorax proteins. Cell 2007, 128, 735–745. [Google Scholar] [CrossRef] [PubMed] [Green Version]
  297. Mikkelsen, T.S.; Xu, Z.; Zhang, X.; Wang, L.; Gimble, J.M.; Lander, E.S.; Rosen, E.D. Comparative epigenomic analysis of murine and human adipogenesis. Cell 2010, 143, 156–169. [Google Scholar] [CrossRef] [PubMed] [Green Version]
  298. Wang, L.; Jin, Q.; Lee, J.E.; Su, I.H.; Ge, K. Histone H3K27 methyltransferase Ezh2 represses Wnt genes to facilitate adipogenesis. Proc. Natl. Acad. Sci. USA 2010, 107, 7317–7322. [Google Scholar] [CrossRef] [Green Version]
  299. Brull, A.; Rodriguez, B.M.; Bonne, G.; Muchir, A.; Bertrand, A.T. The Pathogenesis and Therapies of Striated Muscle Laminopathies. Front. Physiol. 2018, 9, 1533. [Google Scholar] [CrossRef] [Green Version]
  300. Serebryannyy, L.; Misteli, T. Protein sequestration at the nuclear periphery as a potential regulatory mechanism in premature aging. J. Cell Biol. 2018, 217, 21–38. [Google Scholar] [CrossRef]
  301. Vadrot, N.; Duband-Goulet, I.; Cabet, E.; Attanda, W.; Barateau, A.; Vicart, P.; Gerbal, F.; Briand, N.; Vigouroux, C.; Oldenburg, A.R.; et al. The p.R482W substitution in A-type lamins deregulates SREBP1 activity in Dunnigan-type familial partial lipodystrophy. Hum. Mol. Genet. 2015, 24, 2096–2109. [Google Scholar] [CrossRef] [Green Version]
  302. Eberlé, D.; Hegarty, B.; Bossard, P.; Ferré, P.; Foufelle, F. SREBP transcription factors: Master regulators of lipid homeostasis. Biochimie 2004, 86, 839–848. [Google Scholar] [CrossRef] [PubMed]
Figure 1. Mechanosensing mechanisms in adipose tissue. Adipocytes receive mechanical forces at the cell plasma membrane arising from the surrounding extracellular matrix (ECM) or from neighboring cells via integrins, cadherins, GAP junctions or other mechanosensitive proteins such as SWELL1 or PIEZO channels. Mechanical cues can emanate from cell–cell adhesion, cell–ECM adhesion, cytoskeleton remodeling (actomyosin, stress fiber, etc.) and from the growth of lipid droplet (plasma membrane (PM)–lipid droplet (LD) forces) that stiffen the cells (purple arrows). Force sensing and transmission at cell–ECM and cell–cell adhesions converge on the cytoskeleton and can directly be transmitted to the nucleus, resulting in chromatin remodeling and transcriptional changes to modulate cell phenotype (inflammation, fibrosis, insulin resistance, and catecholamine resistance).
Figure 1. Mechanosensing mechanisms in adipose tissue. Adipocytes receive mechanical forces at the cell plasma membrane arising from the surrounding extracellular matrix (ECM) or from neighboring cells via integrins, cadherins, GAP junctions or other mechanosensitive proteins such as SWELL1 or PIEZO channels. Mechanical cues can emanate from cell–cell adhesion, cell–ECM adhesion, cytoskeleton remodeling (actomyosin, stress fiber, etc.) and from the growth of lipid droplet (plasma membrane (PM)–lipid droplet (LD) forces) that stiffen the cells (purple arrows). Force sensing and transmission at cell–ECM and cell–cell adhesions converge on the cytoskeleton and can directly be transmitted to the nucleus, resulting in chromatin remodeling and transcriptional changes to modulate cell phenotype (inflammation, fibrosis, insulin resistance, and catecholamine resistance).
Cells 11 02310 g001
Figure 2. Microenvironment stiffness can regulate adipogenesis. During adipogenesis, the cells undergo substantial morphological modifications where spindle-shaped fibroblast-like preadipocytes become rounded mature adipocytes. There is an extensive reorganization of the cytoskeleton that allows lipid droplet growth and expansion. Upon adipogenic stimulation, the inhibition of Ras homolog family member A (RhoA) and Rho-associated protein kinases (ROCKs) disrupts actin cytoskeleton structures allowing adipogenesis. Thus, the inactive form, RHO GDP, promotes adipogenesis. However, in stiff environments, focal adhesion assembly is promoted, RHO GTP activates ROCK, which, in turn, activates F-actin stress fiber formation and translocation of YAP/TAZ to the nucleus, which breaks adipogenesis.
Figure 2. Microenvironment stiffness can regulate adipogenesis. During adipogenesis, the cells undergo substantial morphological modifications where spindle-shaped fibroblast-like preadipocytes become rounded mature adipocytes. There is an extensive reorganization of the cytoskeleton that allows lipid droplet growth and expansion. Upon adipogenic stimulation, the inhibition of Ras homolog family member A (RhoA) and Rho-associated protein kinases (ROCKs) disrupts actin cytoskeleton structures allowing adipogenesis. Thus, the inactive form, RHO GDP, promotes adipogenesis. However, in stiff environments, focal adhesion assembly is promoted, RHO GTP activates ROCK, which, in turn, activates F-actin stress fiber formation and translocation of YAP/TAZ to the nucleus, which breaks adipogenesis.
Cells 11 02310 g002
Figure 3. Hypothetical mechanical regulation of metabolism. Recent studies revealed that mechanical forces altered cell metabolism via cytoskeletal reorganization in cancer cells [194]. In a soft microenvironment, TRIM21 was not trapped by actin stress fiber bundles, which degrade phosphofructokinase (PFK). This led to low glycolysis rates. By contrast, when cells were surrounded by a stiff microenvironment (as in obese adipose tissue), TRIM21 was bound to the F-actin bundles of the cytoskeleton and thus PFK degradation was prevented, leading to high rates of glycolysis. By these mechanisms, we believe that mechanical cues could influence the metabolic phenotype and progression of adipose tissue dysfunction.
Figure 3. Hypothetical mechanical regulation of metabolism. Recent studies revealed that mechanical forces altered cell metabolism via cytoskeletal reorganization in cancer cells [194]. In a soft microenvironment, TRIM21 was not trapped by actin stress fiber bundles, which degrade phosphofructokinase (PFK). This led to low glycolysis rates. By contrast, when cells were surrounded by a stiff microenvironment (as in obese adipose tissue), TRIM21 was bound to the F-actin bundles of the cytoskeleton and thus PFK degradation was prevented, leading to high rates of glycolysis. By these mechanisms, we believe that mechanical cues could influence the metabolic phenotype and progression of adipose tissue dysfunction.
Cells 11 02310 g003
Figure 4. Roles of mechanosensitive channel in adipocyte. SWELL1/VRAC complex or Piezo1 channels are activated in response to increases in adipocyte volume. SWELL1/VRAC activation stimulates insulin–PI3K–AKT2 via GRB2, and thereby supports lipogenesis and continued adipocyte growth in a feed-forward manner [242] However, opening of Piezo1 in mature adipocytes causes the release of the adipogenic fibroblast growth factor 1 (FGF1), which induces adipogenesis through activation of the FGF-receptor-1 [243].
Figure 4. Roles of mechanosensitive channel in adipocyte. SWELL1/VRAC complex or Piezo1 channels are activated in response to increases in adipocyte volume. SWELL1/VRAC activation stimulates insulin–PI3K–AKT2 via GRB2, and thereby supports lipogenesis and continued adipocyte growth in a feed-forward manner [242] However, opening of Piezo1 in mature adipocytes causes the release of the adipogenic fibroblast growth factor 1 (FGF1), which induces adipogenesis through activation of the FGF-receptor-1 [243].
Cells 11 02310 g004
Publisher’s Note: MDPI stays neutral with regard to jurisdictional claims in published maps and institutional affiliations.

Share and Cite

MDPI and ACS Style

Lecoutre, S.; Lambert, M.; Drygalski, K.; Dugail, I.; Maqdasy, S.; Hautefeuille, M.; Clément, K. Importance of the Microenvironment and Mechanosensing in Adipose Tissue Biology. Cells 2022, 11, 2310. https://doi.org/10.3390/cells11152310

AMA Style

Lecoutre S, Lambert M, Drygalski K, Dugail I, Maqdasy S, Hautefeuille M, Clément K. Importance of the Microenvironment and Mechanosensing in Adipose Tissue Biology. Cells. 2022; 11(15):2310. https://doi.org/10.3390/cells11152310

Chicago/Turabian Style

Lecoutre, Simon, Mélanie Lambert, Krzysztof Drygalski, Isabelle Dugail, Salwan Maqdasy, Mathieu Hautefeuille, and Karine Clément. 2022. "Importance of the Microenvironment and Mechanosensing in Adipose Tissue Biology" Cells 11, no. 15: 2310. https://doi.org/10.3390/cells11152310

Note that from the first issue of 2016, this journal uses article numbers instead of page numbers. See further details here.

Article Metrics

Back to TopTop