Sorting out adipocyte precursors and their role in physiology and disease

Maintenance and expansion of WAT in adults

During the first year of life, adipose tissue continues to expand considerably through cellular hypertrophy and the recruitment of new adipocytes (cellular hyperplasia) (Knittle et al. 1979). In adulthood, adipocyte turnover occurs at a rate of ∼10% per year (Spalding et al. 2008). In rodents, the degree of turnover may be higher, varying from depot to depot (Wang et al. 2013; Rivera-Gonzalez et al. 2016). A remarkable and unique property of adipose tissue is the tremendous capacity to expand or contract in different physiological settings. The capacity to expand is most evident in the setting of obesity, where adipose tissue mass can reach upward of 60% of body weight in the morbidly obese (Ortega et al. 2010). The ability of adipose tissue to shrink and then re-expand is perhaps best highlighted by the response of mammary adipose tissue to pregnancy and lactation. During pregnancy and the lactation period, adipocytes are replaced by milk-producing alveolar structures and then reappear as the alveolar structures involute at weaning (Cinti 2009).

The manner by which adipose tissue expands is of great clinical significance (for review, see Hepler and Gupta 2016). Preferential expansion of subcutaneous depots is associated with protection against cardiometabolic disease in obesity (Lee et al. 2013; Karpe and Pinnick 2015). Preferential expansion of visceral depots correlates well with insulin resistance and diabetes. Moreover, expansion of adipose tissue mass through adipocyte hyperplasia is more metabolically favorable than through an increase in cell size (hypertrophy) (Lee et al. 2010). Hyperplasia occurs through the recruitment of adipocyte precursors that undergo adipogenesis and is associated with adipose health and a delayed onset of insulin resistance. As such, de novo adipocyte differentiation in adults appears critical for the maintenance and expansion of adipose depots (Gustafson et al. 2009).

Lineage tracing studies have revealed that the mode of adipose depot expansion in mice fed a high-fat diet occurs in a depot- and sex-dependent manner (Wang et al. 2013; Kim et al. 2014; Jeffery et al. 2015). Epididymal adipose expansion in adult male mice fed a high-fat diet occurs through both adipocyte hypertrophy and hyperplasia, while the inguinal (subcutaneous) depot in these same animals expands almost exclusively through cellular hypertrophy (Wang et al. 2013). In female mice, adipocyte hyperplasia occurs in both the perigonadal WAT and, to some variable extent, the inguinal WAT (Jeffery et al. 2016). A better understanding of how adipose precursors are activated in obesity may lead to therapeutic strategies to alter body fat distribution and/or adipose tissue health.

In recent years, it has become increasingly clear that adipocyte accumulation in areas beyond the major depots plays an important role in physiology and pathology (Shook et al. 2016). Adipocytes accumulate in the skin below the dermis (dermal WAT, referred to here as “dWAT”) and play a critical role in dermal homeostasis (Alexander et al. 2015; Kruglikov and Scherer 2016). This relatively unexplored depot expands in response to cold exposure, wounding, and bacterial infection. Adipose lineage cells in the skin also respond to hair follicle cycling and regulate follicle homeostasis (Festa et al. 2011). Marrow adipose tissue (MAT), located in various skeletal regions, appears to contribute to local skeletal remodeling, hematopoietic regulation, and systemic metabolism (Scheller et al. 2016). Marrow adipocytes are a developmentally distinct white adipocyte population. MAT, but not other WAT depots, is derived from cells once expressing Osx1 (Berry et al. 2015). MAT expansion occurs in response to various pharmacological or physiological triggers, such as glucocorticoids, anti-diabetic thiazolidinediones, estrogen deficiency, obesity, and aging. Likewise, adipocytes also accumulate in the skeletal muscle at sites of injury (Joe et al. 2010; Uezumi et al. 2010). The accumulation of adipocytes in skeletal muscle can impact the regenerative capacity and function of muscle. Similarly, a defining feature of arrhythmogenic cardiomyopathy is the progressive replacement of the myocardium with fibrous tissue and adipocytes (Samanta et al. 2016). The developmental origin and exact properties and function of these lesser-studied adipocytes are still being unraveled; however, it is clear that the ability of new adipocytes to emerge in these tissues is critical for tissue homeostasis and perhaps systemic metabolism.

White adipocyte precursors: in vitro models

Much of our knowledge of the cellular aspects of adipocyte differentiation emanate from the use of cell culture models. Mouse embryonic fibroblasts (MEFs) from mid-stage to late stage embryos can be readily induced to undergo adipogenesis in response to a hormonal/pharmacological cocktail consisting of dexamethasone, iso-butyl-methyl-xanthine, and insulin. This observation lends credence to the views of Flemming that adipocytes can and readily do emerge from ubiquitous fibroblast-like cells. Similarly, adherent fibroblast-like cells within cultures of the adipose stromal vascular fraction (SVF) have been propagated and differentiated into mature adipocytes in vitro (Poznanski et al. 1973; Van et al. 1976; Van Robin and Roncari 1977). Other mesenchymal lineages, such as osteoblasts, myoblasts, and chondrocytes, can also differentiate from cultures of the adherent adipose SVF (Cawthorn et al. 2012). As such, the adherent SVF from adipose tissue is often described as “adipose stem cells” or “mesenchymal stem cells” (Cawthorn et al. 2012). Importantly, cultured MEFs or SVF cells are heterogeneous; whether these cultures contain cells that truly reflect native adipocyte precursors is not certain.

In the 1970s, Green and Kehinde (1974) derived 3T3-L1 cells as a clonal subline of 3T3 immortalized MEFs from Swiss mice. 3T3-L1 cells are viewed as being a “determined” preadipocyte cell line; the cells are morphologically indistinguishable from less adipogenic or nonadipogenic fibroblasts but are highly committed to the adipocyte lineage. The isolation of such committed cells implied that preadipocytes were specialized fibroblasts with a stably unique molecular program. Preadipocyte cell lines have been powerful models for the study of adipocyte determination and differentiation. Pparγ, the “master regulator” of adipocyte differentiation, was originally discovered through the use of 3T3 cells (Chawla and Lazar 1994; Tontonoz et al. 1994). Today, there are dozens of transcriptional regulators implicated in the process of 3T3-L1 adipocyte differentiation. Excellent reviews have been written on this matter (Farmer 2006; Cristancho and Lazar 2011); however, it is now clear that anatomically distinct adipocytes represent, at least to some degree, distinct lineages. As such, whether cultured preadipocyte cell lines represent any particular type of native adipocyte precursor remains unclear. Here, we summarize recent efforts to identify and characterize native adipose precursors, highlighting their regulation and potential contribution to the adipose lineage and physiology (summarized in Fig. 1).

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Figure 1.

Adipocyte precursor heterogeneity in mice. (A) Characterized fetal adipose precursor populations: Three adipose progenitor populations have been characterized in the developing inguinal WAT depot. The degree of overlap in these populations is unclear. Seale and colleagues (Wang et al. 2014) have identified a brown adipocyte progenitor population in BAT that is marked by myogenic factor 5 (Myf5)-Cre and expresses platelet-derived growth factor α (Pdgfrα) and Ebf2. These cells emerge at embryonic day 10.5 (E10.5) and are devoid of MyoD at this stage, suggesting that brown adipocyte lineage commitment has already occurred. (B) Tissue-resident adipose precursors in adult mice: Adipose precursor populations have been identified in multiple tissues and are listed as they were identified and characterized. There is likely a degree of overlap in the populations. Additional populations may exist based on published lineage tracing results.

Native adipocyte precursors: identity, regulation, and function

Adipose tissue mural cells

Another approach commonly used to locate stem/progenitor populations is to follow the expression of lineage-determining factors. Graff and colleagues (Tang et al. 2008) reasoned that the expression of Pparγ, the most critical adipogenic factor, would facilitate the identification of adipose precursors. Tang et al. (2008) generated reporter mice in which the Pparγ locus drives GFP or LacZ expression. The investigators demonstrated that Pparγ⁺ cells reside within the blood vessel walls of adult WAT and resemble mural cells (pericytes and vascular smooth muscle cells). These cells express several mural cell markers, such as Pdgfrβ, NG2 (neural/glial antigen 2), and SMA (α smooth muscle actin). Furthermore, they uniformly express CD34 and Sca1. Importantly, these cells contribute to WAT homeostasis in lean mice (chow-fed), with an estimate of 10%–20% of new adipocytes forming per month depending on the WAT depots examined (Tang et al. 2008; Jiang et al. 2014). This observation was significant, since it provided the first molecular evidence of the hypotheses put forth by Wasserman (1960) and others (Napolitano 1963; Cinti et al. 1984) decades ago that adipose progenitors reside in or near the vasculature of adipose tissue and resemble pericytes. These data provide evidence that the adipocyte precursor niche is created by the microenvironment of the vasculature.

The hypothesis of a perivascular origin of adipogenesis is also supported by mural cell expression of Zfp423 (Gupta et al. 2012; Vishvanath et al. 2016). Zfp423 is a multizinc finger transcription factor that regulates preadipocyte Pparγ expression and adipocyte differentiation in vitro and in vivo (Gupta et al. 2010). Using Zfp423 reporter mice (transgenic Zfp423^GFP reporter mice), we localized the expression of GFP to a distinct subset of endothelial and mural cells in adipose depots (Gupta et al. 2012). Nonendothelial Zfp423⁺ stromal cells express the mural cell marker Pdgfrβ as well as the aforementioned APC markers Pdgfrα, CD34, and Sca1. The expression of Pparγ and other adipose lineage-selective genes is enriched in Zfp423⁺ mural cells (Vishvanath et al. 2016). Furthermore, most Zfp423⁺ mural cells were devoid of CD24 expression, suggesting that they are among the committed preadipocyte population. The enrichment of Pparγ suggests that this population at least overlaps with the mural cell population examined by Graff and colleagues (Tang et al. 2008). Zfp423⁺ mural cells undergo adipocyte differentiation in vitro more readily than Zfp423⁻ mural cells. Importantly, the abundance of Zfp423⁺ mural cells is physiologically regulated in a sex- and depot-dependent manner. In male mice, the frequency of Zfp423⁺ mural cells is higher and more robustly activated by high-fat diet feeding in the gonadal depot than in the inguinal depot; this correlates well with the hyperplastic potential of the depots in obesity (Vishvanath et al. 2016). A number of questions remain unanswered about this precursor population. It is unclear whether these cells proliferate prior to undergoing differentiation or whether the number of Zfp423⁺ mural cells increases by virtue of activating Zfp423 expression in additional Pdgfrβ⁺ cells. Furthermore, it remains unclear what signals activate these cells and/or Zfp423 expression in response to high-fat diet feeding.

The identification of perivascular adipose precursors is consistent with historical observations that primitive adipocytes are associated with the vasculature. However, direct lineage tracing evidence in support of this hypothesis has been lacking. In the modern era of developmental biology, bona fide lineage tracing is best achieved by using an inducible Cre/lox recombination system for “pulse-chase” lineage tracing. In this approach, a defined cell population is irreversibly marked by the expression of a Cre-dependent reporter (usually LacZ or fluorescent protein) at a selected time (“pulse labeling”). This is achieved by transient activation of Cre recombinase. Marked cells are then detected at later time points to determine how the originally labeled precursors contribute to specific cell types (“chase”). Recently, we and others used lineage tracing models to evaluate the contribution of mural preadipose populations to the maintenance and expansion of the adipocyte pool in adults (Jiang et al. 2014; Berry et al. 2016; Vishvanath et al. 2016). Jiang et al. (2014) used SMA-CreERT2 mice to evaluate the contribution of SMA⁺ mural cells to the maintenance of adipose tissue mass in lean animals. Lineage tracing indicated that the adipocyte pool in adult mice is maintained at least in part through differentiation of SMA⁺ mural APCs. In fact, these cells appear critical for maintaining adipose tissue mass; inducible deletion of Pparγ in these cells leads to progressive lipoatrophy and a lipodystrophic-like metabolic phenotype (Jiang et al. 2014).

Our group has recently derived a novel doxycycline-inducible mural cell lineage tracking system in which the Pdgfrβ promoter ultimately drives Cre expression (Fig. 2; Vishvanath et al. 2016). Using this system (termed the “MuralChaser” mouse), we asked whether Pdgfrβ⁺ mural cells contribute to adipocyte hyperplasia associated with diet-induced obesity. Indeed, we identified several mural cell-derived adipocytes in visceral WAT depots formed during the high-fat diet feeding period (Vishvanath et al. 2016). In agreement with prior lineage tracing studies described above, adipocyte hyperplasia was not observed in the inguinal WAT depot of male mice (Wang et al. 2013). These data provide direct genetic evidence that perivascular preadipocytes contribute to adipocyte hyperplasia in obesity. This is in line with fate-mapping studies of perivascular nestin⁺ cells in the adipose tissue (Nestin-Cre; Rosa26R^tdTomato), revealing the presence of labeled gonadal adipocytes following high-fat diet (Iwayama et al. 2015). It is currently unclear whether the adipogenic capacity of perivascular preadipocytes cells is critical for healthy WAT expansion in the setting of obesity. Functional manipulation of adipogenic signaling pathways (e.g., Pparγ deletion) in Pdgfrβ⁺ cells will help clarify the importance of this population.

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Figure 2.

Tracking de novo adipocyte differentiation using the MuralChaser model. (A) Genetic alleles comprising the MuralChaser lineage tracing system. In the presence of doxycycline (DOX), rtTA activates Cre expression. Cre excises the loxP-flanked membrane tdTomato cassette and allows constitutive activation of membrane GFP (mGFP) reporter expression. (B) “Pulse-chase” lineage tracing approach. Doxycycline-containing food was administered to MuralChaser mice for 9 d to label Pdgfrβ⁺ cells (“pulse”). Animals were then switched to a high-fat diet (HFD) or chow diet for 8 wk in the absence of doxycycline (chase). mGFP⁺ adipocytes represent de novo differentiated fat cells formed during the 8-wk period. (C,D) Administration of doxycycline to MuralChaser mice for 9 d resulted in doxycycline-dependent mGFP labeling (arrowhead) of Pdgfrβ⁺ cells lining the endothelium (CD31⁺) of adipose tissue (“pulse”). (E,F) mGFP labeling was not found in mature adipocytes (perilipin⁺). (G,H) Following 8 wk of high-fat diet feeding, numerous mGFP⁺ perilipin⁺ adipocytes were observed (arrows). These cells represent de novo differentiated adipocytes formed during the 8-wk period. Portions of this figure were reproduced with data from Vishvanath et al. (2016), with permission from Elsevier.

Bone marrow LepR⁺ perivascular cells

Adipogenesis occurs within skeletal bone marrow in response to a number of physiological and pharmacological conditions. Adipocyte accumulation is strongly associated with decreased osteogenesis; however, the signals that influence the balance between the two lineages have remained unclear. Morrison and colleagues (Zhou et al. 2014) identified a subpopulation of multipotent perivascular stem cells defined by the expression of the leptin receptor (LepR). LepR⁺ cells emerge postnatally and represent a majority of the colony-forming cells in the bone marrow (Zhou et al. 2014). They actively express Pdgfrα, Pdgfrβ, and Nestin and are targeted by Prx1-Cre mice. These cells are quiescent but proliferate robustly after injury. Lineage analysis indicates that LepR⁺ perivascular cells are multipotent, giving rise to most adipocytes and osteoblasts accumulating after irradiation or fracture (Zhou et al. 2014). LepR⁺ cells also give rise to marrow adipocytes accumulating with age. Importantly, LepR⁺ cells serve as a hematopoietic stem cell niche, secreting cytokines that promote stem cell maintenance (Ding et al. 2012).

Recently, Yue et al. (2016) determined that LepR signaling itself plays an important role in the cell fate decision of LepR⁺ cells. Leptin signaling in Prx1 lineage cells promotes marrow adipogenesis and inhibits osteogenesis in response to high-fat diet feeding. Interestingly, loss of LepR signaling had no impact on adipogenesis induced by irradiation. This indicates that multiple mechanisms control the fate of LepR⁺ cells in skeletal bone marrow.

Recruitment of beige adipocytes in adults

It has long been recognized that WAT depots of cold-exposed rodents can undergo extensive remodeling and adopt a thermogenic phenotype elicited by the emergence of UCP1⁺ multilocular adipocytes (Loncar 1991). However, the aforementioned lineage tracing reveals that most UCP1⁺ cells within WAT depots are not derived from a Myf5⁺ lineage, suggesting a developmental origin largely distinct from the fetal-derived brown adipocytes (Seale et al. 2008; Sanchez-Gurmaches and Guertin 2014). Cold-induced multilocular UCP1⁺ adipocytes appear to be molecularly and likely functionally distinct from brown adipocytes present in brown adipose depots. Beige cells appear to exhibit properties of both white and brown adipocytes; therefore, these cells are now widely designated as “beige adipocytes.” Cells closely resembling beige adipocytes are present in adult humans, particularly in supraclavicular and paravertebral regions (Sharp et al. 2012; Shinoda et al. 2015).

A great number of pharmacological and physiological stimuli can trigger beige adipocyte accumulation in rodents and humans beyond cold exposure, including cancer cachexia, gastric bypass, exercise, and burn trauma (Kir et al. 2014; Neinast et al. 2015; Sidossis et al. 2015; Stanford et al. 2015). Great progress has been made in elucidating the transcriptional machinery and signaling cascades driving brown and beige adipocyte accumulation; however, in most cases, the actual cellular origin of the UCP1⁺ adipocytes accumulating under physiological conditions or in genetic models has remained unclear. Multiple lineage tracing studies now indicate that UCP1⁺ cells naturally appear rapidly and arise predominantly, but not exclusively, by de novo beige adipogenesis in response to β3AR agonism or cold exposure (Wang et al. 2013; Berry et al. 2016; Shao et al. 2016; Vishvanath et al. 2016). It has also been observed that beige adipocytes have the capacity to revert to a white adipocyte phenotype; the cells lose UCP1 expression and become unilocular when animals are returned to thermoneutral conditions following cold exposure (Rosenwald et al. 2013). These same adipocytes can then revert back to UCP1⁺ cells upon re-exposure to cold. This interconversion of phenotypes is often referred to as “transdifferentiation” (Barbatelli et al. 2010). Classically, transdifferentiation most often refers to a direct conversion of one differentiated cell type into another in the absence of dedifferentiation; this type of event is difficult to prove experimentally. Some have postulated that the white-to-beige phenotype switching may simply reflect beige adipocytes in active and inactive (“dormant”) thermogenic states (Kajimura et al. 2015). It should be noted that current adipocyte lineage tracing approaches do not distinguish between these two possibilities. The adiponectin promoter used for adipocyte targeting is expressed in nearly all types of adipocytes. Clear molecular markers discriminating between unilocular white and unilocular dormant beige adipocytes are still needed. As such, the current data support at least two general mechanisms that lead to the natural formation of beige adipocytes during cold exposure: (1) de novo differentiation from precursors and (2) conversion of unilocular adipocytes in multilocular UCP1⁺ adipocytes (activation of “dormant” unilocular beige adipocytes and/or transdifferentiation).

There is now tremendous interest in elucidating the cellular origin of beige adipocytes. Long et al. (2014) first revealed that beige adipocytes express a smooth muscle-like gene program akin to how classical brown adipocytes exhibit a skeletal muscle-like gene signature. Functional studies also suggest a lineage relationship between beige adipocytes and smooth muscle cells. Stromal vascular cells deficient in Myocardin-related transcription factor A (MRTFA) exhibit less of a smooth muscle-like phenotype and more readily undergo beige adipogenesis (McDonald et al. 2015). Multiple lines of lineage tracing evidence further suggest a smooth muscle origin of beige adipocytes (Long et al. 2014; Berry et al. 2016). Inguinal beige adipocytes rapidly emerging (within 7 d) upon cold exposure descend from cells expressing SMA. Furthermore, at least a subpopulation of beige adipocytes formed in cold-exposed mice originates from Myh11⁺ cells (Myh11-CreERT2 lineage tracing) and mural cells expressing Pdgfrβ (MuralChaser lineage tracing). However, beige adipocytes originating from Myh11⁺ and Pdgfrβ⁺ cells appeared only after 2 wk of cold exposure (Berry et al. 2016; Vishvanath et al. 2016). These data suggest that multiple waves of beige adipogenesis may occur, each drawing on somewhat distinct perivascular precursor populations.

We apologize to our colleagues in the field for not being able to discuss all of the outstanding and important studies related to adipose tissue remodeling and adipose precursors. We thank P. Scherer for critical reading of the manuscript, and members of the Touchstone Diabetes Center at University of Texas Southwestern for useful discussion. R.K.G. is supported by National Institute of Diabetes and Digestive and Kidney Diseases R01 DK104789, and C.H. is supported by the National Institutes of Health National Institute of General Medical Sciences training grant T32 GM008203. Graphic illustrations in this review were designed by Visually Medical (http://www.visuallymedical.com).