Organoid > Volume 3; 2023 > Article
Yoon and Park: Engineered adipose tissue platforms: recent breakthroughs and future perspectives


As overweight and obesity rates have increased worldwide, the prevalence of metabolic disorders has also grown. Due to the lack of physiologically relevant adipose tissue platforms, research in adipose tissue biology has relied on animal models, leading to false conclusions on pathophysiological mechanisms and therapeutic efficacy. Despite the urgent need for an adipose tissue model, it is still extremely difficult to cultivate mature adipocytes and recapitulate multi-cellular interactions in adipose tissue in vitro. For this reason, adipose tissue modeling requires new technologies that allow better culture conditions for adipocytes and contain a complex network of microenvironments. Herein, we discuss recent technologies, including 3-dimensional (3D) adipocyte spheroids, biomaterial-based 3D culture, 3D bioprinting, and microphysiological systems, which may offer new opportunities to discover drugs targeting adipose tissue.


Adipose tissue is a notably complex organ with intense effects on physiology and pathophysiology [1]. The main function of adipose tissue is maintaining lipid and glucose homeostasis in our body as a regulator of energy balance (Fig. 1A). In addition, it plays important roles in the protection of delicate organs, modulation of cell growth and tissue regeneration, and innate immunity [1]. Adipose tissue has long been considered as a mere physical barrier and a store of energy; however, since the discovery of adipokines, including leptin, estrogen, resistin, and inflammatory cytokines, knowledge of the functional aspects of adipose tissue has expanded markedly [2]. In addition, the tremendous global increase in overweight and obesity rates has prompted intense interest in adipose tissue [3,4]. Many studies have blamed adipose tissue dysfunction for promoting the onset and progression of metabolic diseases, including heart disease [5], stroke [6], type 2 diabetes [7], and even some cancers [3] (Fig. 1B). There is now no doubt that understanding adipose tissue is a key to overcoming those diseases as an important therapeutic target.
The majority of cells found in adipose tissue are adipocytes, which contain droplets of stored fat that can be used for energy [8]. They are derived from precursor cells that develop into one of 3 types of adipose tissue: white adipose tissue, which stores energy; brown adipose tissue, which generates heat; and beige adipose tissue, which possesses properties of both white and brown adipose tissues [1]. Adipocytes are surrounded by the extracellular matrix (ECM), which provides structural support and serves as a reservoir of growth factors and cytokines that modulate adipocyte homeostasis [9]. Even though mature adipocytes occupy ~90% of adipose tissue’s volume, there is considerable cellular heterogeneity [10]. Fibroblasts, vascular endothelial cells, adipose stem cells, and a variety of immune cells in the adipose tissue are now being recognized as vital constituents of the adipose microenvironment, especially in the pathophysiology of obesity [10]. Thus, understanding the complex crosstalk between adipocytes and nearby cells/ECM in different states such as physiological processes or metabolic dysfunction is believed to lead to a greater possibility of developing therapeutics. This goal necessitates the development of in vivo and in vitro models recapitulating the complex microenvironment of human adipose tissue. In general, in vivo models more closely simulate biological events than in vitro models; however, animal studies are time- and resource-intensive, possess ethical concerns, and more importantly, are impacted by the serious physiological differences between humans and animals. To address the shortcomings of in vivo studies, in vitro adipose tissue models also have been developed. A 2-dimensional (2D) adherent culture of adipocytes generated from a preadipocyte cell line (e.g., 3T3-L1) or primary preadipocytes and a floating culture of primary adipocytes are typically used as in vitro culture systems in adipose biology [11]. However, these in vitro models present considerable drawbacks, including technical difficulties in the maturation of adipocytes, long-term cell maintenance, and the absence of multi-cellular interactions [11], failing to capture the inherent complexity of adipose tissue.
Recent advances in technologies including biomaterials, stem cell technology, 3-dimensional (3D) bioprinting, and microfluidics have led to great improvements in modeling adipose tissue in normal and pathological conditions. In this paper, we emphasize the importance of multi-cellular adipose tissue models by reviewing the functional roles and intercellular interactions in adipose tissue. Following that, we introduce some advanced culture platforms, including spheroids, 3D scaffolds, transwell systems, 3D bioprinting systems, and microphysiological systems, which can offer a higher fidelity in recapitulating adipose tissue (Fig. 2). Finally, the last part of this review provides perspectives on the significance of each platform in contributing to the understanding of adipose biology and the development of therapeutics for metabolic diseases.
Ethics statement: This study was a literature review of previously published studies and was therefore exempt from institutional review board approval.

Adipose tissue: roles of cellular and non-cellular components

1. Adipocytes

As cells occupying the largest portion of adipose tissue, adipocytes engage in typical functions related to the metabolic and endocrine roles of adipose tissue. In response to energy flux or external stress signals, adipocytes maintain metabolic homeostasis or participate in other regulatory pathways, such as energy storage and consumption, appetite regulation, insulin sensitivity, and inflammatory responses, through various endocrine factors. In addition, continuous exposure to excessive nutritional and stressful environmental changes leads to metabolic diseases due to adipocyte dysfunction, such as insulin resistance and dyslipidemia. Obesity, which is caused by continuous and excessive energy influx in adipose tissue, is typically accompanied by hypertrophy and hyperplasia of adipocytes, resulting in dysfunction of the adipocytes themselves and further dysfunction of surrounding cells. Moreover, the dysfunction of these adipocytes leads to aggravation of the disease through abnormal interactions with other diseased tissues. A recently identified example is tumorigenesis, which has the potential to increase due to obesity. The International Agency for Research on Cancer has reported that higher amounts of body fat are associated with an increased risk of a number of cancers [12].
In addition to changes in size and functionality of adipocytes, adipocytes are highly plastic cells [13]. Adipocytes can be converted into other types of cells according to the physiological/pathological conditions of the individual [14,15]. The most common event is the dedifferentiation of adipocytes that revert to progenitor- and fibroblast-like cell types. This phenomenon is commonly observed in obesity-induced fibrosis of adipose tissue [14,15], tissue regeneration processes [16,17], and interactions with certain tumors [13]. Furthermore, it has been reported that dedifferentiated adipocytes can induce angiogenesis and increase mature vasculature density [18]. Although high plasticity makes it difficult to utilize in in vitro experiments of primary adipocytes, research on dedifferentiation observed in various pathological processes and its accompanying signaling pathways will greatly contribute to understanding related diseases and developing therapeutics.
Therefore, adipose tissue is an organ that actively interacts with itself and other tissues, and research on its function is important for understanding various fat-related diseases as well as basic physiological phenomena such as fat and glucose metabolism and endocrine signal pathways. In the medical field, in vitro modeling is required to improve the understanding of basic research to elucidate the unknown functions of adipose tissue and various diseases related to adipose tissue. A technology that can enhance the functionality of differentiated adipocytes like in vivo conditions or maintain the phenotype of primary adipocytes for a long time in in vitro culture is required. Adipocytes are mainly cultured in 3 ways. The most used cell source for research on adipose tissue is adipogenic differentiated adipocytes [19,20]. Various types of progenitor cells (cell lines, mesenchymal stem cells [MSCs], or the stromal vascular fraction [SVF]) are used to artificially differentiate cells. However, differentiated adipocytes are not morphologically and functionally like in vivo adipocytes, and it is difficult to maintain a long-term phenotype. Another culture method is ex vivo tissue culture, or the culture of primary adipocytes extracted from adipose tissue [20]. However, it is also difficult to maintain fat cells for a long time without dedifferentiation. Moreover, the lack of co-culture technology for interactions between various cells is still an obstacle to studies using adipocytes.

2. Immune cells

The SVF, which can be obtained from adipose tissue, contains heterogeneous cell populations such as mesenchymal progenitor/stem cells, preadipocytes, endothelial cells, pericytes, T cells, and M2 macrophages [21]. Representative resident cells include immune cells. These immune cells are involved in the systemic immune system through the secretion of cytokines while interacting with external signals or surrounding cells [22]. In particular, resident microphages in adipose tissue play a key role in maintaining metabolic homeostasis and insulin sensitivity. In healthy adipose tissue, anti-inflammatory immune cells (typically M2 macrophages, eosinophils, and regulatory T cells) suppress excessive immune responses and induce healthy cell reactivity through proper adipokine balance. However, when exposed to a stressful environment caused by excessive energy accumulation or abnormal cellular responses, inflammatory immune cells (typically M1 macrophages, CD4 and CD8 T-cells, activated natural killer cells, or neutrophils) are activated and collect in adipose tissue [23]. The composition and functionality of the cells become more inflammatory. Remodeling of these immune cells can lead to insulin resistance, which can lead to more serious secondary diseases [24].
The difficulty in reproducing the complex and systemic reactivity has led to the use of laboratory animals for the study of adipose tissue. In vitro models are being developed to replace in vivo experiments and to conduct more relevant research. To model the pathological state of adipose tissue in vitro, interactions between adipocytes and immune cells are required. In many studies, immune cells activated through inflammation-inducing substances such as lipopolysaccharide or pro-inflammatory cytokines are usually used to induce an inflammatory response of adipocytes in vitro and reproduce insulin resistance [25-27]. However, the inflammatory response caused by a single artificial stimulus has limitations in mimicking the dynamic inflammatory response of in vivo adipose tissue.

3. Endothelial cells

Adipose tissue, a representative endocrine organ, has a well-developed vascular structure that facilitates interactions with other tissues [28]. The development of blood vessels in adipose tissue is not only for the maintenance of adipose tissue, which can grow up to 10 times its original size for energy storage, but also for the acceptance and release of various bioactive molecules such as adipokines, cytokines, nutrients, oxygen and metabolic wastes, and other cells such as immune cells and progenitor cells [29]. In addition, blood vessels contain perivascular/endothelial progenitors, which can act as a niche for adipocytes [30]. Therefore, blood vessels are structurally and functionally transformed according to the systemic metabolic activity and disease state of adipose tissue. Although the mechanisms underlying the changes of blood vessels in fat are still elusive, they can be triggered by several dysfunctional modulations, such as hypoxic signals from an out-of-balance energy state and unusual expansion of adipose tissue, pro-inflammatory signals from immune cells, and fibrosis of the ECM [31,32]. In obesity, the expansion of adipose tissue causes a lack of vascular structure, which induces hypoxic signals. Induction of inflammation due to hypoxia causes the collection and accumulation of pro-inflammatory immune cells and dysfunction of surrounding cells [32]. A variety of dysregulated factors stimulate dysfunction of vascular endothelial cells. In addition, excess free fatty acids released from obese adipocytes can induce inflammatory responses in endothelial cells and lead to dysfunction [33].
For a physiological and pathological understanding of adipose tissue, the role of vascular endothelial cells is important. In addition, proper integration of vascular tissue is required for long-term maintenance of adipose tissue in vitro models. For this purpose, various vascularized adipose tissues have been prepared. As an attempt, vascularized adipose organoids were fabricated through in vitro differentiation of cells using SVF containing various adipose-derived cells [34,35]. As another example, the integration of microfluidics enabled improved co-culture of adipocytes and vascular endothelial cells, which allowed the development of an organ-on-a chip that reproduced highly vascularized adipose tissue [36]. These various attempts reproduced the interactions between adipocytes and vascular endothelial cells in vitro and showed potential as more relevant in vitro models.

4. Fibroblasts

In adipose tissue, fibroblasts play an important role in maintaining adipose tissue homeostasis. Fibroblasts, especially fibroblast-specific protein-1-positive fibroblasts, resist obesity induced by a high-calorie diet and cause adipose tissue loss [37]. This is achieved by fibroblasts regulating the turnover of adipocytes and preadipocytes, preventing maintenance of the pool of preadipocytes and regulating their differentiation into adipocytes. Fibroblasts also induce the expression of transcription factors and related sub-markers for adipogenic differentiation of adipose stromal/progenitor cells (ASCs), thus facilitating differentiation even at low densities [38]. Fibroblast differentiated from ASCs also play a major role in the wound healing process [39]. ASCs in adipose tissue induce differentiation into fibroblasts, which are major effector cells for wound healing, and secrete various ECM and growth factors. In addition, adipocytes are converted into fibroblasts through lipolysis in adipocytes around the wound, and they are involved in skin repair [17]. In an obese environment, it is known that preadipocytes with a myofibroblasts phenotype accumulate ECM and upregulate profibrotic cytokines, along with the upregulation of pro-inflammatory factors of fibroblast-like preadipocytes in adipose tissue [40]. Therefore, fibroblasts and myofibroblasts found in adipose tissue cause the onset of metabolic diseases with the development of obesity.
Collectively, fibroblasts not only maintain adipose tissue homeostasis by regulating the turnover of adipocytes, but also the surrounding microenvironment, particularly pathological factors such as obesity, which promotes the differentiation of precursors such as adipocytes or ASCs into fibroblasts and participates in inflammatory signaling by increasing the collection and proliferation of fibroblasts. The various functionalities of fibroblasts can be effectively utilized in in vitro experiments on adipocytes. In particular, fibroblasts could facilitate the differentiation of preadipocytes into adipocytes in vitro and induce a more in vivo-like phenotype. In addition, since fibroblasts are closely involved in the proliferation, differentiation, and maintenance of adipocytes, they are a factor to consider for the successful reproduction of adipose tissue.

5. Extracellular matrix

The ECM is a non-cellular component found in all tissues. It is an extensive 3D network composed of polymeric proteins such as collagen and elastin, proteoglycans including glycosaminoglycans (GAGs), and glycoproteins such as laminin and fibronectin [41]. Collagen is a major component of the ECM that occupies a significant portion of adipose tissue. It serves to support mechanical stress caused by external force or the expansion of adipocytes. Moreover, it is significantly involved in the adhesion, proliferation, and survival of various cells. In the normal ECM, the types and amounts of collagen are properly balanced, but in disease states such as obesity, fibrosis and dysfunction occur due to the accumulation of various types of collagens, such as collagens type I, III, V, and VI [9]. These changes increase the strength of the adipose tissue, limiting its ability to expand. In particular, an increase in type VI collagen and its cleavage product, endotrophin, can be observed in obese adipose tissue, which is involved in metabolic dysfunction such as insulin resistance and even tumorigenesis [4].
In obesity, along with the accumulation of collagen, the number of proteoglycans composed of a core-protein and multiple GAGs chains, which are characterized by a high degree of sulfation and serve as charge-barriers in the ECM [42], also increases [43]. Proteoglycans have multiple ligands, through which they interact with a variety of cytokines, cell surface receptors, adhesion proteins, and other ECM proteins. Various proteoglycans such as lumican, perlecan, decorin, aggrecan, beta-glycan, and biglycan are abundantly present in adipose tissue, are involved with several soluble factors, and take part in cell adhesion, metabolism, and the immune response. As in the case with collagen, it is known that metabolic disorders such as diabetes are closely related to the composition and content of proteoglycans [43,44].
Multiple tissue receptors, including integrin [45], CD44 [46] and CD36 [47], bind to various ECM components, which are implicated in several physiological pathways. These receptors, which enable cell-cell and cell-ECM interactions, are regulated by the surrounding microenvironment to respond appropriately to the maintenance of tissue homeostasis. For example, integrins are known to be involved in the development of insulin resistance. In the obese state, integrin decreases the expression of Glut4, thereby reducing insulin sensitivity and reducing the phosphorylation of AKT, which is involved in glucose uptake [48].
Taken together, the ECM is dynamically regulated by various cells and signals, showing specificity according to the pathological condition of each tissue. Therefore, an analysis of the ECM in various conditions can represent the inflammatory state of an individual. In obesity, adipose tissue greatly expands in volume by storing excess energy, resulting in a change in the ECM. The phenomenon is due to several factors mentioned above. Comprehensive inflammatory signals caused by obesity consistently lead to low levels of systemic inflammation and further exacerbation of ECM fibrosis, contributing to the development of serious secondary diseases such as diabetes, cardiovascular disease, and tumors.

Advances in engineered adipose tissue platforms

As described above, in vivo adipose tissue is highly complex. Multiple cells forming the tissue communicate directly with one another and change their own internal workings to maintain the functionality of adipose tissue. Adipocyte 2D have served as useful tools to study basic adipose biology; however, they may not account for interactions between multiple cell types and complex biochemical processes occurring under 3D conditions. Therefore, to study the metabolism or various roles of adipose tissue under pathological conditions, beyond single cell-based research, adipocyte co-culture technology with other cells is essential to mimic the precise microenvironment of each patient and to study cell-cell interactions. Over the last decade, various studies have shown intercellular interactions through co-culture with adipocytes (Table 1) [49-62], and there has been increasing interest in developing 3D adipose tissue models, and the complexity of models has been increasing (Table 2) [25,26,36,63-76]. However, there is still a lack of a platform and co-culture technology that can increase the availability of mature adipocytes, enable the maturation of preadipocytes, mimic and appropriately control the complex microenvironment. In this part, we describe the adipocyte culture platforms and co-culture approaches that have been developed.

1. Adipocyte spheroid culture

Spheroids are cellular aggregates and one of the most versatile ways to culture cells in 3D (Fig. 2A). They are better able to recapitulate not only the in vivo morphology, but also cell-cell interactions and tissue architecture, which makes them more physiologically relevant and predictive than 2D models. The formation of scaffold-free spheroids relies on ECM secreted from the cells themselves, which encourages preliminary aggregation through the process of self-assembly. Since the ECM secretion and junctional formation of adipocytes are lower than those of preadipocytes, forming adipocyte aggregates is generally less efficient than forming preadipocyte spheroids. Thus, in several studies, white adipose tissue has been modeled by forming preadipocyte spheroids, followed by subsequent differentiation into adipocytes.
Several technical methods, including the hanging drop, liquid overlay, and magnetic levitation techniques, have been used to generate adipose spheroids. The hanging drop culture comprises a small droplet of cell culture medium suspended by gravity, allowing cellular aggregates growth in 3D. Immortalized human and mouse preadipocytes, as well as primary preadipocytes (SVF and adipose-derived stem cells [ADSCs]) have been incorporated into hanging drop cultures and successfully differentiated into adipocytes [77-80]. In a recent study, Klingelhutz et al. [78] developed adipocyte spheroids by placing preadipocytes in drops on the inside cover of a Petri plate, transferring the spheroids to ultra-low-attachment wells, and providing differentiation cues in the medium (Fig. 3A). The spheroids exhibited unilocular lipid droplets seen in mature adipocytes reaching a volume of ~0.1 cm3 and showed less adiponectin secretion than 2D culture. The adiponectin secretion of adipose spheroids was significantly reduced when exposed to the environmental toxins (indoxyl sulfate and PCB126) for 30 days, which was not achievable in 2D culture, raising the possibility of using 3D adipocyte spheroids as a reliable system to study long-term effects of drugs or toxins [78]. Adipocyte spheroid systems have been similarly developed using a liquid overlay technique, also known as static suspension culture, which is based on a non-adhesive surface that prevents cellular adhesion by coating a culture plate with agarose [81] or elastin-like polypeptide-polyethyleneimine [79]. Magnetic levitation was also utilized to form ADSC-based adipocyte spheroids by mixing the ADSCs with magnetic particles (poly-lysine-based magnetic nanoparticles [80]) and subjecting them to magnetic force during culture to make cells stay levitated against gravity.
Scaffold-free adipose spheroids based on patient-derived ADSCs can be utilized to study adipose tissue diseases because of the dependence of cell adherence on autocrine ECM molecules rather than on artificial substrates, giving rise to realistic patterning of diseased cell differentiation and cell-ECM interactions. Al-Ghadban et al. [81] developed patient-derived ADSC spheroids for modeling lipedema, a connective tissue disorder characterized by fibrosis, hypertrophic adipocyte, inflammation, and leaky blood. Spheroid cultures of ADSCs extracted from lipedema patients and healthy individuals were differentiated into adipocytes. The spheroids derived from lipedema patients showed higher matrix metalloproteinases (MMP11) expression than healthy spheroids, which may explain the increase in fibrosis in lipedema adipose tissue [81]. However, they did not show a higher adipogenic differentiation potential related to hypertrophic adipocytes, as has been documented in lipedema patients, possibly because of low diffusion of nutrients and oxygen throughout the entire spheroid due to the absence of vasculature.
To overcome this drawback, several attempts have been made to generate a vascularized adipose spheroid. 3T3-L1 preadipocytes aggregated with murine endothelial cells (bEND.3) showed the formation of a vascular-like network concomitantly with lipogenesis in perivascular cells upon adipogenesis induction [80]. Spheroid culture of SVF also displayed a vascular-like structure without using exogenous endothelial cells [80]. However, the absence of biomaterial-based scaffolds makes it challenging to create functional vasculature due to the lack of angiogenic signals. Muller et al. [34] solved this problem by developing co-differentiation methods for SVF adipocytes/endothelial cells. The incorporation of SVF spheroids in Matrigel and 4 days of endothelial growth induction in EGM2 medium provided the angiogenic signals to form a highly branched vasculature (Fig. 3B). Subsequent differentiation of perivascular cells using an optimized adipogenic cocktail (without IBMX, rosiglitazone, and indomethacin) successfully preserved the vascular network and induced adipogenesis in a spheroid. The adipose spheroids generated using the optimized method showed much longer and highly branched vascular networks than those generated using the traditional method. These models can serve as useful tools to study interactions between adipocytes and endothelial cells and offer a possibility of long-term maintenance of adipose spheroids by supplying nutrients and oxygen to the sphere center via the vascular network.
Recently, the presence of SVF immune cells in adipose spheroids from SVF was demonstrated by Taylor et al. [82]. As the SVF is composed of a heterogeneous assortment of cells, even including immune cells, it was predicted that adipose organoids generated from SVF might contain immune cell populations. The researchers found that 20%-30% of differentiated adipose spheroid cells were CD45+ immune cells, and of the immune cell populations, macrophages (CD11b+F4/80+) accounted for 60% to 70% [82]. This finding implies that resident immune cells can be maintained in 3D spheroids, even in the absence of M-CSF supplementation in the culture medium, whereas SVF macrophages could not be maintained in 2D culture [82]. This study focused efforts on the evaluation of changes in the lipidome of spheroid in response to dietary interventions, rather than investigating the interaction between adipocytes and immune cells; however, it suggests that scaffold-free adipose spheroids could be utilized to model many pathological conditions associated with immune responses in adipose tissue.

2. Biomaterial scaffold-based 3D adipocyte culture

Three-dimensional porous or permeable biomaterials promote the recapitulation of the structure and functionality of native tissues through cell-to-ECM and cell-to-cell interactions (Fig. 2B). For decades, biomaterial scaffolds such as collagen [83,84], gelatin (Fig. 3D) [85,86], polymer mixtures [87-89] and decellularized ECM [90,91] demonstrated roles in the improvement of adipogenic differentiation efficiency or the maintenance of mature adipocytes. Researchers have recently started to use this strategy for the long-term culture of mature adipocytes in vitro. Louis et al. [83] developed a collagen microfiber-based mature adipocyte culture method, which ensured the maintenance of unilocular mature adipocytes for up to 2 weeks, exhibiting higher cell viability and perilipin expression than 2D-cultured adipocytes. Another study [89] suggested a biofunctionalized hydrogel with components of adipose ECM as a relevant in vitro model for adipose tissue (Fig. 3E). To provide the adipocytes with spatial and chemical cues for effective maturation, they developed a cross-linked hyaluronic acid scaffold containing key adipose ECM components. The 3D cultured preadipocytes in the fabricated scaffolds showed improved in vivo-like morphology and metabolism. However, these strategies are generally employed for adipose tissue regeneration; thus, the development of 3D scaffolds has been focused on the enhancement of in vivo stability and the functional lifetime, supporting adipocytes inside the body.

3. Transwell culture of mature adipocytes

A transwell system is a cell-culture device that provides independent access to both sides of a permeable plastic membrane (Fig. 2C). Harms et al. [92] recently established a new primary adipocyte culture method using transwells, named membrane mature adipocytes aggregate cultures (MAAC). Culturing freshly isolated mature adipocytes underneath the permeable membrane of the transwell system enhanced the maintenance of the integrity and functionality of adipocytes for up to 2 weeks without dedifferentiation (Fig. 3C). Considering that functional maintenance of mature adipocytes is highly required to study long-term drug responses in vitro, the MAAC-based culture system exhibits great potential to serve as a promising drug test platform to treat metabolic diseases. The MAAC method allowed the co-culture of adipocytes with macrophages in a different compartment, which is also advantageous for inducing pathological conditions such as inflammation-associated metabolic disorders. This improved platform can be an experimental tool capable of increasing the in vitro accessibility of primary adipocytes for various applications, such as the pathologic modeling of obesity or drug screening. However, there is still an insufficient understanding of the multi-cellular interactions and dynamic ECM remodeling observed in obese adipose tissue.

4. 3D bioprinting for engineering adipose tissue

Over the last decade, 3D printing emerged as a promising technology to generate the adipose tissue construct for tissue regeneration, due to its efficient control of spatial positioning of multiple cells using various bio-inks (Fig. 2D) [63,64,66-68,93,94]. Cho et al. [66] developed a flexible adipose-vascular tissue assembly composed of a synthetic polymer-based substructure and decellularized ECM bio-inks using planar 3D printing (Fig. 3F). The formation of vascular structures was induced by printing human umbilical vein endothelial cells with vascular tissue-derived ECM bio-ink; meanwhile, the functional maintenance of adipocytes was successfully achieved using adipose tissue-derived ECM bio-ink. To enhance the long-term in vivo retention efficiency, a polycaprolactone module holder was additionally used, resulting in maintenance of its original shape over 7 days. Another study reported that MSCs bioprinted using GelMA bio-ink were robustly differentiated into the adipogenic lineage (Fig. 3G) [68]. The researchers demonstrated that the pore size of the scaffold determined the differentiation and infiltration efficiency, highlighting the importance of the physiochemical properties of bio-ink. This approach can be exploited to develop adipose tissue co-culture platforms to model pathological conditions in vitro [26,65]. Crosstalk between adipocytes and breast cancer cells was recapitulated using 3D bioprinting technology [65]. The spatially controlled deposition of adipocytes and breast cancer cells enabled the generation of direct or indirect contact between the 2 types of cells, showing how adipocytes contribute to cancer aggressiveness. Insulin resistance was also modeled by patterning ADSCs and activated macrophages as a platform to study drug sensitivity [26]. Significant limitations of 3D bioprinting remain in terms of pattern resolution; however, its high flexibility and versatility would offer great advantages in creating complex adipose tissue models. These findings suggest that 3D bioprinting technology may be suitable for the regeneration studies with artificial adipose tissue through improved vascular co-culture and structural simulation. These results also show the possibility of pathological modeling through the mixing of various materials. However, even with this advanced technology, some obstacles remain. Non-cell-friendly printing processes and selection-limiting types of bio-inks and their properties are known as limitations of 3D bioprinting, which are still being improved. In addition, unlike the various evaluations of adipose tissue performed on other platforms, the in vitro maturation of printed artificial tissues and extensive functional evaluation thereof are lacking in 3D bioprinting technology.

5. Microfluidic physiological system for engineering adipose tissue

A microphysiological system (MPS) is an integrated 3D culture platform designed to recapitulate the key organ features (Fig. 2E). This technology presents advantages over other in vitro systems by mimicking the precisely controlled physical, chemical, and biological microenvironment of organs. Because adipose tissue is a dynamic organ composed of a variety of cell populations, which actively interact with other organs by secreting various adipokines, MPS technology has been suggested as a promising tool for emulating its complex physiology [36,76,95-97] and pathology [25,70-72,74,75,98,99].
Rogal et al. [76] developed a two-channel adipose tissue MPS (white adipose tissue-on-a-chip) to solve the issue of low maintenance of mature adipocyte culture. The mature adipocytes cultured within a microfluidic channel were functionally maintained for 36 days while embedded in collagen hydrogel due to the efficient nourishment from vasculature-like perfusion. Isoproterenol-stimulated lipolysis was also observed in the device, showing its potential as a platform for pharmaceutical development and testing (Fig. 3H). Even though media perfusion enabled the long-term maintenance of mature adipocytes in the device, functional regulation of adipocytes through crosstalk with vascular endothelial cells was not achievable in the study. Yang et al. [36] developed a vascularized adipose tissue MPS using adipocytes and endothelial cells. To accomplish adipogenic differentiation and vascularization under the same media conditions, the composition of cell culture media was extensively optimized in the microfluidic device. The adipose tissue generated in the optimized media revealed the formation of an interconnected vascular network (Fig. 3I); however, the cellular function of the vascularized adipose tissue platform was not fully investigated. For disease modeling, MPS has been used to recapitulate the complex pathophysiological features of adipose tissue in disease conditions. An insulin-resistant adipose tissue model was developed using a two-channel microfluidic device, where mature adipocytes were cultured in a microchannel interfaced with a media perfusion chamber [72]. Insulin resistance was efficiently induced by treating mature adipocytes with tumor necrosis factor-alpha, as confirmed through Akt phosphorylation, Glut4 expression, and glucose uptake. Furthermore, the therapeutic efficacy of rosiglitazone, an antidiabetic drug, was evaluated in an MPS device, demonstrating its potential for the study of metabolic diseases and the discovery of new therapeutics. Similarly, Liu et al. [74] reproduced the insulin resistance of adipocytes by culturing adipocytes and immune cells together in adjacent spaces separated by a porous barrier. The insulin sensitivity of the resulting micro-artificial adipose tissue was verified by observing glucose uptake in the presence of insulin and changes in adipokine expression. The currently developed adipose tissue MPS systems have demonstrated the benefits of microfluidic systems, including the long-term cultivation of adipocytes without a loss of viability by supplying medium through flow, and dynamic communication with different types of cells through the different chambers. Although various types of microfluidic devices have been proposed for adipose tissue-related studies, there are still difficulties in inducing cell-ECM interactions or complex in vivo-like inflammatory reactions.


In this review, we discussed the importance of advanced multi-cellular adipose tissue models by reviewing the functional role of cellular components and the ECM in adipose tissue. Next, we focused on discussing the advantages and limitations of current engineering methods for in vivo-like adipose tissue models, including adipocyte spheroids, 3D scaffold-embedded adipocyte culture, transwell-based adipocyte aggregate culture, 3D bioprinting, and MPS (Table 3). Even though recent technologies have enabled better recapitulation of adipose tissue compared to traditional adipocyte culture methods, there is a long way to go to succeed in replicating the complexity of human adipose tissue. Many studies have achieved the generation of multi-cellular adipose tissue in vitro; however, crosstalk between adipocytes and other cellular components including endothelial cells and fibroblasts was not fully validated. In addition, there is a lack of research on the optimization of the ECM and culture medium for co-cultivation, which may lead to inaccuracies in cell response to drug treatments. While adipose tissue naturally interacts with different organs as an important endocrine organ in our body, endocrine-mediated organ-organ interactions have not been recapitulated. The development of multi-organ constructs, such as liver-adipose tissue or pancreas-adipose tissue, will open a new approach to overcome adipose tissue-related metabolic disorders.


Conflict of interest

No potential conflict of interest relevant to this article was reported.



Data availability

Please contact the corresponding author for data availability.

Fig. 1.
Adipocytes communicate with other organs and contribute to pathological states. (A) Adipose tissue directly and indirectly interacts with various organs and maintains energy homeostasis. It interacts with various organs through bioactive molecules or extracellular vesicles. (B) In addition, chronic inflammatory conditions caused by obesity adversely affect the prevalence, development, treatment, and prognosis of metabolic diseases.
Fig. 2.
In vitro adipocyte culture platforms. (A) Adipocyte spheroid culture, (B) three-dimensional (3D) culture with biomaterials, (C) membrane mature adipocyte aggregate culture, (D) 3D bioprinting, (E) microfluidic physiological system.
Fig. 3.
Recent examples of adipocyte culture platforms. This figure presents examples of platforms for culturing (A, B) adipose tissue using spheroid culture, (C) transwell culture, (D, E) 3D cultures with a biomaterial scaffold, (F, G) 3D bioprinting and (H, I) microfluidic technology. Representative images of (A) the hanging drop method for scaffold-free generation of uniform adipose spheroids [78] and (B) scaffold-based vascularized adipose tissue spheroid formation [34]. Reproduced and slightly modified from Klingelhutz et al. Sci Rep 2018;8:523 [78] and Muller et al. Sci Rep 2019;9:7250 [34], according to the Creative Commons license. (C) Membrane mature adipocyte aggregate culture platform. Reproduced from Harms et al. Cell Rep 2019;27:213-25, according to the Creative Commons license [92]. Biomaterial-based 3D adipocyte culture inducing improved adipogenic differentiation with gelatin- [86] (D) and polymer mixture-based scaffolds [89] (E). Reproduced and slightly modified from Contessi et al. J Appl Polym Sci 2019;136:47104 [86] and Louis et al. Biotechnol Bioeng 2017;114:1813-24 [89], with permission from John Wiley and Sons. (F) Representative image of assembled vascularized adipose tissue [66]. (G) Representative images of 3D-printed gelatin-based scaffolds with different pore sizes [68]. Adapted with permission from Cho et al. Adv Healthc Mater 2021;10:e2001693 [66] and Tytgat et al. Macromol Biosci 2020;20: e1900364 [68]. (H) Schematic of the chip platform featuring two independent systems with eight tissue chambers each [76]. (I) A microfluidic device consisting of five cell culture chambers flanked by two fluidic channels and the formation of vascularized adipose tissue [36]. Reproduced from Yang et al. Bioengineering (Basel) 2020;7:114 [36] and Rogal et al. Sci Rep 2020;10:6666 [76], according to the Creative Commons license.
Table 1.
Summary of co-culture studies
Purpose of co-culture Co-culture
Type of in vitro platform used Effect of co-culture Ref.
Adipocytes Other cell types
To induce obese micro-environment 3T3-L1 BMDMs and ATMs Transwell plate (0.4 μm pore) 6-fold increase in miR-223 abundance. [49]
Several fold increase in ATM-derived miR-223 expression.
3T3-L1 RAW 264.7 Transwell plate (0.4 μm pore) Mimicking obesity-related inflammation with lipopolysaccharide stimulation. [50]
Identification of riboflavin pro-inflammatory activity inhibitory effect.
3T3-L1 RAW 264.7 Transwell plate (0.4 μm pore) Improvement of the induction of various inflammatory responses observed in an obese environment. [51]
3T3-L1 RAW 264.7 Transwell plate Increased ANGPTL2 mRNA expression. [52]
MMP9 mRNA upregulation.
Identification of pathways that cause inflammation in adipocytes and immune cells.
3T3-L1 RAW 264.7 Transwell plate Reproduction of the adipose microenvironment. [53]
Verification of anti-inflammatory effects of SO1989.
PHA and SGBS preadipocytes THP-1 Transwell plate (0.4 μm pore) Identification of the role of IL-29 as a regulator. [54]
To recapitulate the interaction between BC cells 3T3-L1 BC cells (MDA-MB-231, MCF7, SUM159, or Hs578t) 3D co-culture system with Matrigel Breast tissue reproduction. [55]
More accurate analysis of EMT and MET
3T3-F442A and PHA BC cells (MDA-MB-436 and E0771) Transwell plate (0.4 μm pore) The increased expression of MVP. [56]
Induction of MVP-related multidrug resistance phenotype in BC cells by adipocytes.
Murine fat tissue or PMA BC cell (MDA-MB-231) Transwell plate (0.4 μm pore) Increased expression of inflammation-related genes in co-culture of adipocytes. [57]
PHA CC cells (PT130 and SW480) Transwell plate Increased CPT1A expression. [58]
Increased survival of CC cells.
Identification of CPT1A, a key regulator of metabolism carried by adipocytes in CC cells.
PHA CC cell (PT93) Transwell plate Transfer of free fatty acids that were released from adipocytes to the CC cells. [59]
Induction of CC cell autophagy by AMPK activation of adipocytes.
PHA GC cells Transwell plate Reduced anoikis [60]
Elevated CD36 and its transcription factor in GC cells.
Higher basal and maximal respiration rates and higher intracellular ATP levels by adipocytes
mADSC, preadipocytes, or primary mesothelial cells OC cell (A224) Transwell plate (0.4 μm pore) Reduction of pyruvic acid and lactic acid production in OC cells by adipocytes [61]
Mediation of the growth inhibitory effect of ITLN1 on OC cells of mature adipocytes.
3T3-L1 LC cell (A549) Ad-CM and transwell plate (0.4 μm pore) A549 cells induced adipocyte lipolysis. [62]
Adipocytes altered energy metabolism, particularly in LC cells through glycolysis.

BMDMs, bone-marrow-derived macrophages; ATMs, adipose tissue macrophages; SGBS, Simpson-Golabi-Behmel syndrome; BC cells, breast cancer cells; CC cells, colon cancer cells; GC cells, gastric cancer cells; OC cell, ovarian cancer cell; PHA, primary human adipocytes; PMA, primary murine adipocytes; mADSC, mature adipocytes differentiated from adipose-derived stem cells; LC cell, lung cancer cell; Ad-CM, conditioned medium from adipocytes; ANGPTL2, angiopoietin-like protein; MMP9, matrix metalloproteinase-9; IL-29, interleukin-29; EMT, epithelial-to-mesenchymal transition; MET, mesenchymal-to-epithelial transition; MVP, major vault protein; CPT1A, carnitine palmitoyltransferase I; AMPK, AMP-activated protein kinase; ATP, adenosine triphosphate; ITLN1, intelectin1.

Table 2.
Adipocyte culture platforms based on 3D printing and microfluidics
Technology used Cell type Cell culture specificity Total culture period Function evaluated Ref.
Extrusion-based 3D printing Human ASCs GelMA and CarMA Day 14 Cell viability [63]
Adipogenic differentiation ability
ASCs/HUVECs co-culture spheroids GelMA Day 7 Adipogenic differentiation and ◦vascularization [64]
MCF-7 and ADSCs Blend of alginate and gelatin Day 10 Optimization of co-culture with MCF-7 and ADSCs [65]
Adipogenic differentiation
Human preadipocyte cells and HUVECs Adipose tissue- and aortas-dECM Week 4 Adipogenic differentiation and vascularization [66]
In vivo volume retention
Integration with host tissue with neovascularization in artificial tissue
3T3-L1 Crosslinked gelatin hydrogels Day 14 Direct cytocompatibility [67]
Adipogenic differentiation
ADMSCs, 3T3-L1 and RAW264.7 Alginate, gelatin, and collagen Day 12 Adipogenic differentiation [26]
Changes of insulin resistance
Drug screening
Human bone marrow MSCs GelMA Day 8 Adipogenic differentiation [68]
Spatial distribution of cells and lipid droplets in scaffolds
Microfluidics 3T3-L1 PDMS chip that can connect with MS Day 13 Monitoring secreted metabolites with MS detection [69]
3T3-L1 and J774A.1 Integrated adipose-tissue-on-chip nanoplasmonic biosensing platform Day 15 Adipocyte differentiation, maturation, and inflammatory stimulation [70]
Cytokine detection
Visceral adipose tissue PDMS on-chip tissue culture platform Day 6 Tissue viability and morphology [71]
Biopsy culture Patient-specific glucose uptake detection
Insulin sensitivity
ASCs, HUVECs and NHLFs Fibrinogen Day 10 Adipogenic differentiation and vascularization [36]
Microfluidic device consisting of 5 cell culture chambers flanked by 2 fluidic channels Vascularized adipose tissue
Primary murine preadipocytes A dual-layer, membrane-based microfluidic devices Day 14 Adipogenesis. [72]
Insulin resistance (lipid droplet, glucose uptake and western blot and Glut-4 localization)
Lipid metabolism
PMA Collagen - Adipocyte culture [73]
PDMS chip with a reservoir for adipocyte capture Adiponectin secretion for sampling
Human preadipocytes and U937 Si-based microfluidic chip with 3 main features Day 20 Adiponectin and IL-6 secretion [74]
- Concentric cell culture compartments Insulin resistance by glucose uptake
- Microchannel arrays
- Media channels
Human preadipocytes Cytoarchitecture on the chip with fiber networks Day 83 Adipogenesis. [75]
Adipocyte hypertrophy
Functional responses to simulated meals and fasting
PHA Collagen Day 47 Validation of adipocyte functionality; glucose and fatty acid metabolism [76]
Tailored microfluidic platform Long-term functionality
- 3D-tissue chambers Drug test with isoproterenol
- Perfusable microchannels
PBMCs and PHA Microfluidic chip separated by porous membrane. Day 14 Adipocyte differentiation [25]
- Lower part: 2 fluidic compartments Immune-metabolic analysis
- Upper part: adipocytes/immune cells culture

3D, 3-dimensional; ASC, adipose tissue-derived stem cells; HUVEC, human umbilical vein endothelial cells; ADSCs, adipose-derived stromal cells; MSCs, mesenchymal stromal cells; ADMSCs, adipose-derived mesenchymal stem cells; PBMCs, peripheral blood mononuclear cells; NHLFs, normal human lung fibroblasts; GelMA, methacrylamide-modified gelatin; CarMA, methacrylate-modified κ-carrageenan; dECM, decellularized extracellular membrane; PDMS, polydimethylsiloxane; MS, mass spectrometry; Glut-4, glucose transporter type 4; IL-6, interleukin-6.

Table 3.
Advantages and disadvantages of each platform
Cell culture Advantages Disadvantages
Spheroid culture Cell-to-cell interaction, simple uniform spheroid production, high-throughput. Time and labor-consuming method, absence of ECM, difficulty in maintenance and utilization of spheroids, uncontrolled levels of differentiation and thus lack of consistency.
3D culture with scaffold Cell-ECM interactions, improved adipogenic differentiation through facile biomolecular integration, functional improvement of the formed adipose tissue, drug resistance, relatively simple process. High dependence of the physical properties of biomaterials on cellular reactivity, scale limitation.
Transwell Long-term culture of mature adipocyte without dedifferentiation while remaining functional, multi-cellular interactions. Expensive commercial tools required (transwell), available only for mature adipocytes, absence of ECM.
3D bioprinting Custom-made detailed architecture, co-culture ability, positioning of cells and biomaterials, high-throughput. Complex printing process, non-cell-friendly process, instrument required for process, limited selection of bio-ink, resolution dependent on the physical properties of bio-ink, lack of tissue maturation in the printout, low availability of mature adipocytes
Microfluidics Mimics blood flow, easy to perform multiplex analysis with a small amount of material, co-culture ability, ease of electrical and optical analysis through sensor integration for monitoring metabolites. Difficulty of high-throughput analysis, complex chip manufacturing process and culture method, size limitation.

3D, 3-dimensional; ECM, extracellular matrix.


1. Zwick RK, Guerrero-Juarez CF, Horsley V, Plikus MV. Anatomical, physiological, and functional diversity of adipose tissue. Cell Metab 2018;27:68-83.
crossref pmid pmc
2. Ahima RS, Flier JS. Adipose tissue as an endocrine organ. Trends Endocrinol Metab 2000;11:327-32.
crossref pmid
3. Quail DF, Dannenberg AJ. The obese adipose tissue microenvironment in cancer development and progression. Nat Rev Endocrinol 2019;15:139-54.
crossref pmid pmc pdf
4. Scully T, Ettela A, LeRoith D, Gallagher EJ. Obesity, type 2 diabetes, and cancer risk. Front Oncol 2021;10:615375.
crossref pmid pmc
5. Oikonomou EK, Antoniades C. The role of adipose tissue in cardiovascular health and disease. Nat Rev Cardiol 2019;16:83-99.
crossref pmid pdf
6. Haley MJ, Mullard G, Hollywood KA, Cooper GJ, Dunn WB, Lawrence CB. Adipose tissue and metabolic and inflammatory responses to stroke are altered in obese mice. Dis Model Mech 2017;10:1229-43.
pmid pmc
7. Lee MW, Lee M, Oh KJ. Adipose tissue-derived signatures for obesity and type 2 diabetes: adipokines, batokines and MicroRNAs. J Clin Med 2019;8:854.
crossref pmid pmc
8. Rosen ED, Spiegelman BM. What we talk about when we talk about fat. Cell 2014;156:20-44.
crossref pmid pmc
9. Ruiz-Ojeda FJ, Méndez-Gutiérrez A, Aguilera CM, Plaza-Díaz J. Extracellular matrix remodeling of adipose tissue in obesity and metabolic diseases. Int J Mol Sci 2019;20:4888.
crossref pmid pmc
10. Corvera S. Cellular heterogeneity in adipose tissues. Annu Rev Physiol 2021;83:257-78.
crossref pmid pmc
11. Sadie-Van Gijsen H. Adipocyte biology: it is time to upgrade to a new model. J Cell Physiol 2019;234:2399-425.
crossref pmid pdf
12. Lauby-Secretan B, Scoccianti C, Loomis D, Grosse Y, Bianchini F, Straif K, et al. Body fatness and cancer: viewpoint of the IARC Working Group. N Engl J Med 2016;375:794-8.
crossref pmid pmc
13. Bielczyk-Maczynska E. White adipocyte plasticity in physiology and disease. Cells 2019;8:1507.
crossref pmid pmc
14. Lessard J, Pelletier M, Biertho L, Biron S, Marceau S, Hould FS, et al. Characterization of dedifferentiating human mature adipocytes from the visceral and subcutaneous fat compartments: fibroblast-activation protein alpha and dipeptidyl peptidase 4 as major components of matrix remodeling. PLoS One 2015;10:e0122065.
crossref pmid pmc
15. Jones JE, Rabhi N, Orofino J, Gamini R, Perissi V, Vernochet C, et al. The adipocyte acquires a fibroblast-like transcriptional signature in response to a high fat diet. Sci Rep 2020;10:2380.
crossref pmid pmc pdf
16. Liao Y, Zeng Z, Lu F, Dong Z, Chang Q, Gao J. In vivo dedifferentiation of adult adipose cells. PLoS One 2015;10:e0125254.
crossref pmid pmc
17. Shook BA, Wasko RR, Mano O, Rutenberg-Schoenberg M, Rudolph MC, Zirak B, et al. Dermal adipocyte lipolysis and myofibroblast conversion are required for efficient skin repair. Cell Stem Cell 2020;26:880-95.
crossref pmid pmc
18. Watanabe H, Goto S, Kato R, Komiyama S, Nagaoka Y, Kazama T, et al. The neovascularization effect of dedifferentiated fat cells. Sci Rep 2020;10:9211.
crossref pmid pmc pdf
19. Ruiz-Ojeda FJ, Rupérez AI, Gomez-Llorente C, Gil A, Aguilera CM. Cell models and their application for studying adipogenic differentiation in relation to obesity: a review. Int J Mol Sci 2016;17:1040.
crossref pmid pmc
20. Dufau J, Shen JX, Couchet M, De Castro Barbosa T, Mejhert N, Massier L, et al. In vitro and ex vivo models of adipocytes. Am J Physiol Cell Physiol 2021;320:C822-41.
crossref pmid
21. Ramakrishnan VM, Boyd NL. The adipose stromal vascular fraction as a complex cellular source for tissue engineering applications. Tissue Eng Part B Rev 2018;24:289-99.
crossref pmid pmc
22. Chung KJ, Nati M, Chavakis T, Chatzigeorgiou A. Innate immune cells in the adipose tissue. Rev Endocr Metab Disord 2018;19:283-92.
crossref pmid pdf
23. Wensveen FM, Valentić S, Šestan M, Wensveen TT, Polić B. Interactions between adipose tissue and the immune system in health and malnutrition. Semin Immunol 2015;27:322-33.
crossref pmid
24. Wu H, Ballantyne CM. Metabolic inflammation and insulin resistance in obesity. Circ Res 2020;126:1549-64.
crossref pmid pmc
25. Kongsuphol P, Gupta S, Liu Y, Bhuvanendran Nair Gourikutty S, Biswas SK, Ramadan Q. In vitro micro-physiological model of the inflamed human adipose tissue for immune-metabolic analysis in type II diabetes. Sci Rep 2019;9:4887.
crossref pmid pmc pdf
26. Park SB, Koh B, Jung WH, Choi KJ, Na YJ, Yoo HM, et al. Development of a three-dimensional in vitro co-culture model to increase drug selectivity for humans. Diabetes Obes Metab 2020;22:1302-15.
crossref pmid pdf
27. Mohallem R, Aryal UK. Regulators of TNFα mediated insulin resistance elucidated by quantitative proteomics. Sci Rep 2020;10:20878.
crossref pmid pmc pdf
28. Gu P, Xu A. Interplay between adipose tissue and blood vessels in obesity and vascular dysfunction. Rev Endocr Metab Disord 2013;14:49-58.
crossref pmid pdf
29. Herold J, Kalucka J. Angiogenesis in adipose tissue: the interplay between adipose and endothelial cells. Front Physiol 2021;11:624903.
crossref pmid pmc
30. Tang W, Zeve D, Suh JM, Bosnakovski D, Kyba M, Hammer RE, et al. White fat progenitor cells reside in the adipose vasculature. Science 2008;322:583-6.
crossref pmid pmc
31. Li M, Qian M, Kyler K, Xu J. Adipose tissue-endothelial cell interactions in obesity-induced endothelial dysfunction. Front Cardiovasc Med 2021;8:681581.
crossref pmid pmc
32. Spencer M, Unal R, Zhu B, Rasouli N, McGehee RE Jr, Peterson CA, et al. Adipose tissue extracellular matrix and vascular abnormalities in obesity and insulin resistance. J Clin Endocrinol Metab 2011;96:E1990-8.
crossref pmid pmc
33. Ghosh A, Gao L, Thakur A, Siu PM, Lai CW. Role of free fatty acids in endothelial dysfunction. J Biomed Sci 2017;24:50.
crossref pmid pmc pdf
34. Muller S, Ader I, Creff J, Leménager H, Achard P, Casteilla L, et al. Human adipose stromal-vascular fraction self-organizes to form vascularized adipose tissue in 3D cultures. Sci Rep 2019;9:7250.
crossref pmid pmc pdf
35. Ioannidou A, Alatar S, Schipper R, Baganha F, Åhlander M, Hornell A, et al. Hypertrophied human adipocyte spheroids as in vitro model of weight gain and adipose tissue dysfunction. J Physiol 2022;600:869-83.
crossref pmid pdf
36. Yang F, Cohen RN, Brey EM. Optimization of co-culture conditions for a human vascularized adipose tissue model. Bioengineering (Basel) 2020;7:114.
crossref pmid pmc
37. Zhang R, Gao Y, Zhao X, Gao M, Wu Y, Han Y, et al. FSP1-positive fibroblasts are adipogenic niche and regulate adipose homeostasis. PLoS Biol 2018;16:e2001493.
crossref pmid pmc
38. Ejaz A, Hatzmann FM, Hammerle S, Ritthammer H, Mattesich M, Zwierzina M, et al. Fibroblast feeder layer supports adipogenic differentiation of human adipose stromal/progenitor cells. Adipocyte 2019;8:178-89.
crossref pmid pmc pdf
39. Zhou ZQ, Chen Y, Chai M, Tao R, Lei YH, Jia YQ, et al. Adipose extracellular matrix promotes skin wound healing by inducing the differentiation of adipose‑derived stem cells into fibroblasts. Int J Mol Med 2019;43:890-900.
crossref pmid
40. Keophiphath M, Achard V, Henegar C, Rouault C, Clément K, Lacasa D. Macrophage-secreted factors promote a profibrotic phenotype in human preadipocytes. Mol Endocrinol 2009;23:11-24.
crossref pmid pmc
41. Davies JA. Extracellular matrix. In: eLS, John Wiley & Sons Ltd.; 2001.
42. Zhang F, Zhang Z, Linhardt RJ. Chapter 3: glycosaminoglycans. Cummings RD, Pierce JM. In: Handbook of glycomics San Diego: Academic Press; 2009 59-80.
43. Pessentheiner AR, Ducasa GM, Gordts PL. Proteoglycans in obesity-associated metabolic dysfunction and meta-inflammation. Front Immunol 2020;11:769.
crossref pmid pmc
44. Pessentheiner AR, Quach A, Al-Azzam N, Liu S, Downes M, Evans RM, et al. Adipose tissue heparan sulfate proteoglycans: critical regulators of adipocyte metabolism and glucose homeostasis. FASEB J 2020;34(S1):1.
45. Ruiz-Ojeda FJ, Wang J, Bäcker T, Krueger M, Zamani S, Rosowski S, et al. Active integrins regulate white adipose tissue insulin sensitivity and brown fat thermogenesis. Mol Metab 2021;45:101147.
crossref pmid pmc
46. Kang HS, Liao G, DeGraff LM, Gerrish K, Bortner CD, Garantziotis S, et al. CD44 plays a critical role in regulating diet-induced adipose inflammation, hepatic steatosis, and insulin resistance. PLoS One 2013;8:e58417.
crossref pmid pmc
47. Cai L, Wang Z, Ji A, Meyer JM, van der Westhuyzen DR. Scavenger receptor CD36 expression contributes to adipose tissue inflammation and cell death in diet-induced obesity. PLoS One 2012;7:e36785.
crossref pmid pmc
48. Hatem-Vaquero M, Griera M, García-Jerez A, Luengo A, Álvarez J, Rubio JA, et al. Peripheral insulin resistance in ILK-depleted mice by reduction of GLUT4 expression. J Endocrinol 2017;234:115-28.
crossref pmid
49. Ying W, Riopel M, Bandyopadhyay G, Dong Y, Birmingham A, Seo JB, et al. Adipose tissue macrophage-derived exosomal mirnas can modulate in vivo and in vitro insulin sensitivity. Cell 2017;171:372-84.
crossref pmid
50. Mazur-Bialy AI, Pocheć E. Riboflavin reduces pro-inflammatory activation of adipocyte-macrophage co-culture. potential application of vitamin B2 enrichment for attenuation of insulin resistance and metabolic syndrome development. Molecules 2016;21:1724.
crossref pmid pmc
51. Kim M, Song K, Kim YS. Alantolactone improves palmitate-induced glucose intolerance and inflammation in both lean and obese states in vitro: adipocyte and adipocyte-macrophage co-culture system. Int Immunopharmacol 2017;49:187-94.
crossref pmid
52. Kim J, Lee SK, Jang YJ, Park HS, Kim JH, Hong JP, et al. Enhanced ANGPTL2 expression in adipose tissues and its association with insulin resistance in obese women. Sci Rep 2018;8:13976.
crossref pmid pmc pdf
53. Yang N, Tang Q, Qin W, Li Z, Wang D, Zhang W, et al. Treatment of obesity-related inflammation with a novel synthetic pentacyclic oleanane triterpenoids via modulation of macrophage polarization. EBioMedicine 2019;45:473-86.
crossref pmid pmc
54. Lin TY, Chiu CJ, Kuan CH, Chen FH, Shen YC, Wu CH, et al. IL-29 promoted obesity-induced inflammation and insulin resistance. Cell Mol Immunol 2020;17:369-79.
crossref pmid pdf
55. Pallegar NK, Garland CJ, Mahendralingam M, Viloria-Petit AM, Christian SL. A novel 3-dimensional co-culture method reveals a partial mesenchymal to epithelial transition in breast cancer cells induced by adipocytes. J Mammary Gland Biol Neoplasia 2019;24:85-97.
crossref pmid pdf
56. Lehuédé C, Li X, Dauvillier S, Vaysse C, Franchet C, Clement E, et al. Adipocytes promote breast cancer resistance to chemotherapy, a process amplified by obesity: role of the major vault protein (MVP). Breast Cancer Res 2019;21:7.
crossref pmid pmc pdf
57. Blücher C, Iberl S, Schwagarus N, Müller S, Liebisch G, Höring M, et al. Secreted factors from adipose tissue reprogram tumor lipid metabolism and induce motility by modulating PPARα/ANGPTL4 and FAK. Mol Cancer Res 2020;18:1849-62.
crossref pmid pdf
58. Xiong X, Wen YA, Fairchild R, Zaytseva YY, Weiss HL, Evers BM, et al. Upregulation of CPT1A is essential for the tumor-promoting effect of adipocytes in colon cancer. Cell Death Dis 2020;11:736.
crossref pmid pmc pdf
59. Wen YA, Xing X, Harris JW, Zaytseva YY, Mitov MI, Napier DL, et al. Adipocytes activate mitochondrial fatty acid oxidation and autophagy to promote tumor growth in colon cancer. Cell Death Dis 2017;8:e2593.
crossref pmid pmc pdf
60. Tan Y, Lin K, Zhao Y, Wu Q, Chen D, Wang J, et al. Adipocytes fuel gastric cancer omental metastasis via PITPNC1-mediated fatty acid metabolic reprogramming. Theranostics 2018;8:5452-68.
crossref pmid pmc
61. Au-Yeung CL, Yeung TL, Achreja A, Zhao H, Yip KP, Kwan SY, et al. ITLN1 modulates invasive potential and metabolic reprogramming of ovarian cancer cells in omental microenvironment. Nat Commun 2020;11:3546.
crossref pmid pmc pdf
62. Li FF, Zhang H, Li JJ, Cao YN, Dong X, Gao C. Interaction with adipocytes induces lung adenocarcinoma A549 cell migration and tumor growth. Mol Med Rep 2018;18:1973-80.
crossref pmid pmc
63. Tytgat L, Van Damme L, Ortega Arevalo MD, Declercq H, Thienpont H, Otteveare H, et al. Extrusion-based 3D printing of photo-crosslinkable gelatin and κ-carrageenan hydrogel blends for adipose tissue regeneration. Int J Biol Macromol 2019;140:929-38.
crossref pmid
64. Benmeridja L, De Moor L, De Maere E, Vanlauwe F, Ryx M, Tytgat L, et al. High-throughput fabrication of vascularized adipose microtissues for 3D bioprinting. J Tissue Eng Regen Med 2020;14:840-54.
crossref pmid pdf
65. Chaji S, Al-Saleh J, Gomillion CT. Bioprinted three-dimensional cell-laden hydrogels to evaluate adipocyte-breast cancer cell interactions. Gels 2020;6:10.
crossref pmid pmc
66. Cho WW, Kim BS, Ahn M, Ryu YH, Ha DH, Kong JS, et al. Flexible adipose-vascular tissue assembly using combinational 3D printing for volume-stable soft tissue reconstruction. Adv Healthc Mater 2021;10:e2001693.
crossref pmid pdf
67. Contessi Negrini N, Celikkin N, Tarsini P, Farè S, Święszkowski W. Three-dimensional printing of chemically crosslinked gelatin hydrogels for adipose tissue engineering. Biofabrication 2020;12:025001.
crossref pmid pdf
68. Tytgat L, Kollert MR, Van Damme L, Thienpont H, Ottevaere H, Duda GN, et al. Evaluation of 3D printed gelatin-based scaffolds with varying pore size for MSC-based adipose tissue engineering. Macromol Biosci 2020;20:e1900364.
crossref pmid pdf
69. Dugan CE, Grinias JP, Parlee SD, El-Azzouny M, Evans CR, Kennedy RT. Monitoring cell secretions on microfluidic chips using solid-phase extraction with mass spectrometry. Anal Bioanal Chem 2017;409:169-78.
crossref pmid pdf
70. Zhu J, He J, Verano M, Brimmo AT, Glia A, Qasaimeh MA, et al. An integrated adipose-tissue-on-chip nanoplasmonic biosensing platform for investigating obesity-associated inflammation. Lab Chip 2018;18:3550-60.
crossref pmid pmc pdf
71. Zambon A, Zoso A, Gagliano O, Magrofuoco E, Fadini GP, Avogaro A, et al. High temporal resolution detection of patient-specific glucose uptake from human ex vivo adipose tissue on-chip. Anal Chem 2015;87:6535-43.
crossref pmid
72. Tanataweethum N, Zhong F, Trang A, Lee C, Cohen RN, Bhushan A. Towards an insulin resistant adipose model on a chip. Cell Mol Bioeng 2020;14:89-99.
crossref pmid pmc pdf
73. Godwin LA, Brooks JC, Hoepfner LD, Wanders D, Judd RL, Easley CJ. A microfluidic interface for the culture and sampling of adiponectin from primary adipocytes. Analyst 2015;140:1019-25.
crossref pmid pmc
74. Liu Y, Kongsuphol P, Chiam SY, Zhang QX, Gourikutty SB, Saha S, et al. Adipose-on-a-chip: a dynamic microphysiological in vitro model of the human adipose for immune-metabolic analysis in type II diabetes. Lab Chip 2019;19:241-53.
crossref pmid
75. Pope BD, Warren CR, Dahl MO, Pizza CV, Henze DE, Sinatra NR, et al. Fattening chips: hypertrophy, feeding, and fasting of human white adipocytes in vitro. Lab Chip 2020;20:4152-65.
crossref pmid pmc
76. Rogal J, Binder C, Kromidas E, Roosz J, Probst C, Schneider S, et al. WAT-on-a-chip integrating human mature white adipocytes for mechanistic research and pharmaceutical applications. Sci Rep 2020;10:6666.
crossref pmid pmc pdf
77. Naderi N, Wilde C, Haque T, Francis W, Seifalian AM, Thornton CA, et al. Adipogenic differentiation of adipose-derived stem cells in 3-dimensional spheroid cultures (microtissue): implications for the reconstructive surgeon. J Plast Reconstr Aesthet Surg 2014;67:1726-34.
crossref pmid
78. Klingelhutz AJ, Gourronc FA, Chaly A, Wadkins DA, Burand AJ, Markan KR, et al. Scaffold-free generation of uniform adipose spheroids for metabolism research and drug discovery. Sci Rep 2018;8:523.
crossref pmid pmc pdf
79. Turner PA, Gurumurthy B, Bailey JL, Elks CM, Janorkar AV. Adipogenic differentiation of human adipose-derived stem cells grown as spheroids. Process Biochem 2017;59:312-20.
crossref pmid pmc
80. Daquinag AC, Souza GR, Kolonin MG. Adipose tissue engineering in three-dimensional levitation tissue culture system based on magnetic nanoparticles. Tissue Eng Part C Methods 2013;19:336-44.
crossref pmid
81. Al-Ghadban S, Pursell IA, Diaz ZT, Herbst KL, Bunnell BA. 3D spheroids derived from human lipedema ASCs demonstrated similar adipogenic differentiation potential and ECM remodeling to non-lipedema ASCs in vitro. Int J Mol Sci 2020;21:8350.
crossref pmid pmc
82. Taylor J, Sellin J, Kuerschner L, Krähl L, Majlesain Y, Förster I, et al. Generation of immune cell containing adipose organoids for in vitro analysis of immune metabolism. Sci Rep 2020;10:21104.
crossref pmid pmc pdf
83. Louis F, Kitano S, Mano JF, Matsusaki M. 3D collagen microfibers stimulate the functionality of preadipocytes and maintain the phenotype of mature adipocytes for long term cultures. Acta Biomater 2019;84:194-207.
crossref pmid
84. Emont MP, Yu H, Jun H, Hong X, Maganti N, Stegemann JP, et al. Using a 3D culture system to differentiate visceral adipocytes in vitro. Endocrinology 2015;156:4761-8.
crossref pmid pmc
85. Huber B, Borchers K, Tovar GE, Kluger PJ. Methacrylated gelatin and mature adipocytes are promising components for adipose tissue engineering. J Biomater Appl 2016;30:699-710.
crossref pmid pdf
86. Contessi Negrini N, Tarsini P, Tanzi MC, Farè S. Chemically crosslinked gelatin hydrogels as scaffolding materials for adipose tissue engineering. J Appl Polym Sci 2019;136:47104.
crossref pdf
87. Qi D, Wu S, Kuss MA, Shi W, Chung S, Deegan PT, et al. Mechanically robust cryogels with injectability and bioprinting supportability for adipose tissue engineering. Acta Biomater 2018;74:131-42.
crossref pmid
88. Kessler L, Gehrke S, Winnefeld M, Huber B, Hoch E, Walter T, et al. Methacrylated gelatin/hyaluronan-based hydrogels for soft tissue engineering. J Tissue Eng 2017;8:2041731417744157.
crossref pmid pmc pdf
89. Louis F, Pannetier P, Souguir Z, Le Cerf D, Valet P, Vannier JP, et al. A biomimetic hydrogel functionalized with adipose ECM components as a microenvironment for the 3D culture of human and murine adipocytes. Biotechnol Bioeng 2017;114:1813-24.
crossref pmid pdf
90. Zhao Y, Fan J, Bai S. Biocompatibility of injectable hydrogel from decellularized human adipose tissue in vitro and in vivo. J Biomed Mater Res B Appl Biomater 2019;107:1684-94.
crossref pmid pdf
91. Tan QW, Zhang Y, Luo JC, Zhang D, Xiong BJ, Yang JQ, et al. Hydrogel derived from decellularized porcine adipose tissue as a promising biomaterial for soft tissue augmentation. J Biomed Mater Res A 2017;105:1756-64.
crossref pmid pdf
92. Harms MJ, Li Q, Lee S, Zhang C, Kull B, Hallen S, et al. Mature human white adipocytes cultured under membranes maintain identity, function, and can transdifferentiate into brown-like adipocytes. Cell Rep 2019;27:213-25.
crossref pmid
93. Kambe Y, Ogino S, Yamanaka H, Morimoto N, Yamaoka T. Adipose tissue regeneration in a 3D-printed poly(lactic acid) frame-supported space in the inguinal region of rats. Biomed Mater Eng 2020;31:203-10.
crossref pmid
94. Van Damme L, Briant E, Blondeel P, Van Vlierberghe S. Indirect versus direct 3D printing of hydrogel scaffolds for adipose tissue regeneration. MRS Adv 2020;5:855-64.
95. Yokomizo A, Morimoto Y, Nishimura K, Takeuchi S. Temporal observation of adipocyte microfiber using anchoring device. Micromachines (Basel) 2019;10:358.
crossref pmid pmc
96. Loskill P, Sezhian T, Tharp KM, Lee-Montiel FT, Jeeawoody S, Reese WM, et al. WAT-on-a-chip: a physiologically relevant microfluidic system incorporating white adipose tissue. Lab Chip 2017;17:1645-54.
crossref pmid pmc
97. Li X, Brooks JC, Hu J, Ford KI, Easley CJ. 3D-templated, fully automated microfluidic input/output multiplexer for endocrine tissue culture and secretion sampling. Lab Chip 2017;17:341-9.
crossref pmid pmc
98. Ramadan Q, Gourikutty SB, Zhang QX. OOCHIP: compartmentalized microfluidic perfusion system with porous barriers for enhanced cell-cell crosstalk in organ-on-a-chip. Micromachines (Basel) 2020;11:565.
crossref pmid pmc
99. Ahluwalia A, Misto A, Vozzi F, Magliaro C, Mattei G, Marescotti MC, et al. Systemic and vascular inflammation in an in-vitro model of central obesity. PLoS One 2018;13:e0192824.
crossref pmid pmc

Editorial Office
Room 319, Hall 1 of Chonbuk National University Dental College, 20, Geonji-ro, Deokjin-gu, Jeonju 54907, Korea
Tel: +82-63-270-4024    E-mail:                

Copyright © 2024 by The Organoid Society.

Developed in M2PI

Close layer
prev next