PMCCPMCCPMCC

Search tips
Search criteria 

Advanced

 
Logo of nihpaAbout Author manuscriptsSubmit a manuscriptNIH Public Access; Author Manuscript; Accepted for publication in peer reviewed journal;
 
Science. Author manuscript; available in PMC Apr 24, 2009.
Published in final edited form as:
PMCID: PMC2597101
NIHMSID: NIHMS68943
White Fat Progenitors Reside in the Adipose Vasculature*
Wei Tang,1 Daniel Zeve,1 Jae Myoung Suh,1 Darko Bosnakovski,1 Michael Kyba,1 Robert E. Hammer,2,4 Michelle D. Tallquist,2 and Jonathan M. Graff1,2,3
1Department of Developmental Biology University of Texas Southwestern Medical Center 6000 Harry Hines Blvd. NB5.118 Dallas, Texas 75390−9133, USA
2Department of Molecular Biology University of Texas Southwestern Medical Center 6000 Harry Hines Blvd. NB5.118 Dallas, Texas 75390−9133, USA
3Department of Internal Medicine University of Texas Southwestern Medical Center 6000 Harry Hines Blvd. NB5.118 Dallas, Texas 75390−9133, USA
4Department of Biochemistry, University of Texas Southwestern Medical Center at Dallas
To whom correspondence should be addressed. Jon.Graff/at/utsouthwestern.edu
White adipose (fat) tissues regulate metabolism, reproduction and lifespan. Adipocytes form throughout life, with the most marked expansion of the lineage occurring during the postnatal period. Adipocytes develop in coordination with the vasculature, but the identity and location of white adipocyte progenitor cells are unknown. We used genetically marked mice to isolate proliferating and renewing adipogenic progenitors. We find that most adipocytes descend from a pool of these proliferating progenitors that are already committed either prenatally or early in postnatal life. These progenitors reside in the mural cell compartment of the adipose vasculature but not in the vasculature of other tissues. Thus, the adipose vasculature appears to function as a progenitor niche and may provide signals for adipocyte development.
How adipocytes (fat cells) develop is a fundamental biological question with important ramifications for human health and disease (1, 2). little is known about the identity, localization, or biological characteristics of endogenous adipocyte progenitors (2). These progenitors likely reside in the adipose stromal-vascular fraction (SVF), a heterogeneous mixture of cells operationally-defined by enzymatic dissociation of adipose depots followed by density separation from adipocytes (1, 3). Peroxisome proliferator-activated receptor gamma (PPARγ), a central regulator of fat formation, is necessary and sufficient for adipogenesis (4, 5). Thus, marking PPARγ-expressing cells in vivo might provide new insights into adipose lineage specification.
To mark and perform lineage analyses on PPARγ-expressing cells, we generated PPARγ-tet transactivator (tTA) (6) “knock-in” mice placing tTA under the control of the PPARγ locus (fig. S1). We introduced into these “PPARγ-tTA” mice two additional alleles: a tTA-responsive Cre allele (tetracycline response element-Cre; “TRE-Cre”) and an element that indelibly expresses lacZ in response to the Cre recombinase (ROSA26-flox-stop-flox-lacZ) (7, 8). With these genetic manipulations, we thereby created a “PPARγ-reporter” strain (“PPARγ-R26R” for PPARγ-Rosa26 reporter) in which the endogenous PPARγ promoter/enhancer induces expression of tTA, leading to Cre expression and an indelible lacZ marking of PPARγ-expressing cells and all descendants (fig. S1). The PPARγ-tTA strain functioned as expected; that is, it was active in adipose depots and repressed by doxycycline (Dox), establishing a tool to examine the adipose lineage (Fig. 1A, figs. S1 and S2).
Figure 1
Figure 1
PPARγ-expressing progenitors proliferate and maintain the precursor pool
To capture the rapid and dramatic expansion of the adipose lineage that occurs during the first postnatal month (1, 9), we Dox-treated the PPARγ-R26R mice starting at different days during this crucial window (fig. S3A). Interestingly, we found homogenous lacZ expression in P30 adipose depots that was not appreciably altered even when Dox administration began in the first postnatal days (Fig. 1A and fig. S3). This surprising result indicated that the vast majority of P30 adipocytes derived from a pre-existing pool of PPARγ-expressing cells, either adipocytes already present prenatally/early postnatally or proliferating precursors. Both interpretations conflict with previous data, however. The possibility that these cells are pre-formed adipocytes is incompatible with the proliferative increase that occurs over this time frame, while the notion that PPARγ-expressing cells are progenitors is inconsistent with cell culture studies (10, 11). To distinguish between the two possible interpretations, we examined the Dox-induced response of another reporter, TRE-H2B-GFP, that is stable in post-mitotic cells but, in contrast to the indelible lacZ marker, becomes diluted in proliferating cells following inhibition of the tet system (12, 13). Strikingly, Dox treatment (P2-P30) markedly reduced adipose depot and adipocyte GFP expression (Fig. 1A), indicating that PPARγ-expressing cells proliferate. Consistent with these data, ~50% of adipocytes were labeled by bromodeoxyuridine (BrdU) when administered between P10 and P30 (fig. S4). The stability of lacZ marking together with the diminishing GFP expression indicate that adipose lineage cells, already instructed to express PPARγ prenatally, proliferate and are the major source of the spurt of adipocyte development observed in the first month of life.
The adipose stromal-vascular fraction (SVF) (fig. S5) is postulated to contain adipocyte progenitors (1, 14). We therefore investigated this location as a possible source from which the proliferating PPARγ-expressing cells characterized above may originate. Indeed we found that a subset of SV cells expressed immunocytochemically-detectable levels of PPARγ as well as the lacZ and GFP reporters (Fig. 1B and fig. S6). These SV resident PPARγ-expressing cells proliferate, as they incorporated BrdU after a brief 2-hour chase, even when the BrdU pulse-chase was initiated after 10 days of Dox pre-treatment to ensure that cells containing both GFP and BrdU expressed GFP prior to initiation of the brief BrdU pulse (fig. S7). In addition, GFP+ SV cells isolated by fluorescence-activated cell sorting (FACS) had considerable proliferative capacity (fig. S8). Further support for the in vivo proliferation of the GFP+ SV cells derives from flow cytometry profiles showing a Dox-induced (P2-P30) decrease in the number and fluorescent intensity of GFP+ SV cells (Fig. 1C and fig. S9). Notably, Dox did not reduce the number or percentage of lacZ+ SV cells, indicating that a pool of PPARγ-expressing cells remains in the SV compartment (Fig. 1D). Together these data indicate that the SV compartment of adipose depots contains PPARγ-expressing cells that divide, are mobilized from and also repopulate the SVF, and behave as an amplifying population that contributes to the adipocyte lineage.
We assessed the adipogenic potential, in vitro and after transplantation, of FACS-isolated GFP+ SV cells (fig. S10). In culture, the sorted GFP+ SV cells underwent spontaneous and insulin-stimulated adipogenesis that was enhanced compared to GFP- SV cells (Figs. 2A, 2B and fig. S11). GFP+ SV adipogenesis mirrored the gene expression patterns described for preadipocyte cell line adipogenesis and the induced adipocytes expressed the perilipin protein with the appropriate subcellular distribution (Fig. 2C) (15). Moreover, freshly isolated GFP+ P30 SV cells transplanted into nude mice led to formation of an ectopic GFP+ depot, containing lipid-laden adipocytes that co-expressed GFP and perilipin (Figs. 2D, 2E and 2F). Thus these GFP+ SV cells have the proliferative and adipogenic properties expected of the endogenous progenitor population.
Figure 2
Figure 2
PPARγ-expressing SV cells are adipogenic and have a unique molecular signature
To characterize the GFP+ SV progenitors and their relationship to other cells present in the adipose depot, we assessed cell surface marker expression using flow cytometry and FACS (fig. S10). The majority of GFP+ SV cells expressed Sca1 and CD34, but not CD105, CD45, TER-119, or Mac-1 (Fig. 2G and fig. S12). When these markers were used to positively or negatively select cells (and independently of the GFP reporter), we again isolated a subset of SV cells that generated a GFP+ ectopic adipose depot after transplantation (fig. S13). Some GFP+ SV cells could potentially be differentiated adipocytes that had yet to accumulate enough lipid to float during the density-based SV fractionation procedure. However, reporters driven by the promoter/enhancer of aP2 (16), a marker of adipocytes and a PPARγ target gene, displayed strong expression in adipose depots and adipocytes but not in SV cells, unlike the PPARγ reporters (Fig. 2G and fig. S14). Immunocytochemical analyses also showed that the GFP+ SV cells did not express perilipin, an adipocyte marker (Fig. 2C). In addition, the FACS-isolated GFP+ SV cells were molecularly distinct from adipocytes, expressing higher levels of the preadipocyte marker Pref-1, the adipogenic inhibitor GATA3, and targets of the anti-adipogenic Wnt (Wisp2) and Hedgehog (Smo, Gli3) pathways and much lower levels of numerous adipocyte markers (e.g., C/EBPα, FAS, leptin, etc.) (Fig. 2H and fig. S15A). Gene expression profiles further defined the GFP+ SV cells as a unique population within adipose tissues (Fig. 2I). Differentially expressed genes include: developmental transcription factors (e.g., goosecoid and Twist2); extracellular matrix genes (e.g., MMP3); anti-angiogenic factors (e.g., Stab1); and signaling cascade components (e.g., EGFR and FGF10) (fig. S15B). Thus, GFP+ SV cells are phenotypically distinct from adipocytes and other SV cells and have a unique molecular signature that allows prospective isolation for transplantation and further lineage analyses.
The local microenvironment (niche) is a crucial determinant of progenitor fate, function, and maintenance (17). In part due to the nature of the SV dissociation and isolation method, the anatomical location and neighboring cells of the SV adipocyte precursors are not known. To investigate the architecture of the SV compartment, we developed an SV particulate (SVP) isolation procedure designed to partially maintain the native SV structure while removing adipocytes that obscure visualization of the precursor location (fig. S16). In the SVPs, the majority of GFP+ cells were arrayed in tube-like structures (Fig. 3A). The GFP+ cells present in freshly isolated tubes did not contain lipid-droplets based upon inspection and lack of lipid staining (Figs. 3A and 3B). Organotypic cultures of SVPs led to formation of lipid-laden GFP+ adipocytes along the tubes, indicating that the tube-associated SVP GFP+ cells were adipogenic (Figs. 3B and 3C). Since the SVP tubes resembled blood vessels, we stained them with antibodies that recognize constituent cells of the vasculature including PECAM (endothelium) and three mural cell markers: SMA, PDGFRβ and NG2 (18). The SVP tubes expressed PECAM and were surrounded by cells that expressed SMA, PDGFRβ and NG2, indicating that they were vessels (Figs. 3D and 3E). Notably, GFP+ SVP cells expressed these mural cell markers (Figs. 3D and 3E). The notion that PPARγ might be expressed in a subset of mural cells is noteworthy as cultured mural cells, similar to mesenchymal stem cells, are multipotent and can be induced to undergo adipogenesis, chondrogenesis, osteogenesis and myogenesis and may provide a progenitor reservoir (18, 19).
Figure 3
Figure 3
SV particulate vessels contain GFP+ precursors that form adipocytes
To investigate the distribution of the GFP+ progenitors within the mural cell compartment, we immunohistochemically-examined sections of freshly frozen PPARγ-GFP P30 adipose depots and other organs. In the adipose vasculature, we again observed co-localization of GFP and mural cell markers (Figs. 4A and 4B). The GFP+ vessels were of various sizes and disseminated throughout the depot (Fig. 4A). However, only a subset of mural cells within a vessel expressed GFP and some adipose vessels did not appear to harbor GFP+ progenitors (fig. S17). Strikingly, mural cells in other examined P30 tissues (including skeletal and cardiac muscles, kidney, retina, pancreas, spleen, lung, etc.) did not express the GFP reporter (Fig. 4C and fig. S18). In older animals (~6 months), we did detect GFP in some small caliber PECAM-positive, SMA-negative adult skeletal muscle vessels (figs. S19 and S20). The majority of these adult skeletal muscle GFP+ cells expressed PECAM, and the cells were not adipogenic (fig. S20). Thus adipose depots appear to contain a unique population of progenitors present in the adipose depot mural cell compartment.
Figure 4
Figure 4
GFP+ cells are present in adipose depot mural cells
PDGFRβ marks mural cells and is required for their development (18). To explore the possibility that PDGFRβ-expressing cells were part of the adipocyte lineage, we X-gal stained adipose depots of P30 mice that contained both a PDGFRβ-Cre transgene (20), which expresses Cre in mural cells and other developing cells, and R26R. As a specificity control, we used SM22-Cre (21), a driver construct expressed in a subset of vascular smooth muscle cells. In these Cre-mediated lineage studies, we found that PDGFRβ-Cre generated strong and relatively homogenous lacZ expression throughout adipose depots in adipocytes and mural cells (Fig. 4D). In contrast, SM22-Cre did not, although lacZ was expressed in a distinct subset of adipose depot vessels (Fig. 4D).
To assess the adipogenic potential of PDGFRβ-expressing mural cells, we isolated PDGFRβ positive and negative cells from white adipose tissues and other organs by FACS, cultured them in insulin or transplanted them into nude mice (fig. S21). In both assays, the adipose depot PDGFRβ+ SV cells had high and substantially more adipogenic potential than PDGFRβ- SV cells (Fig. 4E); this adipogenesis was stimulated by thiazolidinediones (TZDs), diabetes drugs that activate PPARγ (22) (Fig. 4F). In contrast, PDGFRβ+ cells isolated from other organs did not display such potential and were unresponsive to TZDs (fig. S22 and Fig. 4F). Although we could identify sections that contained adipocytes in the non-adipose transplants, these adipocytes were GFP-negative (in contrast to adipocytes present in adipose depot SV PDGFRβ+ transplants) (fig. S22), apparently derived or recruited from host tissues. These data are consistent with the possibility that adipocyte progenitors reside as adipose depot mural cells with distinct properties such as adipogenic potential.
The intertwined epidemics of obesity and diabetes have led to a public health crisis that demands an improved understanding of adipocyte biology (2, 23). Yet the identity of the adipocyte progenitors and their precise location has remained elusive. Exploiting genetic reporters, we show that the pool of murine white adipocyte precursors has largely been committed prenatally or just after birth. These precursors divide, maintain the progenitor pool, and produce adipocytes. Some of these progenitors appear to be mural cells that reside in the vasculature of adipose tissues, results supported by early electron micrographic studies (24, 25). Thus the adipose vasculature appears to function as a progenitor niche and may provide signals for adipocyte development.
Several previous studies have documented an interplay between adipose tissue and the vasculature and shown that this interaction provides possible targets for obesity-diabetes therapies (26-29). The results described here add a new perspective to this interplay. In addition, they provide a foundation for further characterization of the adipose vascular niche and for prospective isolation of the adipocyte progenitors. Such experiments should help establish whether intervention in adipose lineage formation can be an effective therapeutic approach for obesity and diabetes.
Supplementary Material
Supp
Footnotes
*This manuscript has been accepted for publication in Science. This version has not undergone final editing. Please refer to the complete version of record at http://www.sciencemag.org/. Their manuscript may not be reproduced or used in any manner that does not fall within the fair use provisions of the Copyright Act without the prior, written permission of AAAS.
1. Ailhaud G, Grimaldi P, Negrel R. Annu Rev Nutr. 1992;12:207. [PubMed]
2. Gesta S, Tseng YH, Kahn CR. Cell. 2007;131:242. [PubMed]
3. Klaus S, Cassard-Doulcier AM, Ricquier D. J Cell Biol. 1991;115:1783. [PMC free article] [PubMed]
4. Lazar MA. Biochimie. 2005;87:9. [PubMed]
5. Farmer SR. Cell Metab. 2006;4:263. [PMC free article] [PubMed]
6. Kistner A, et al. Proc Natl Acad Sci U S A. 1996;93:10933. [PubMed]
7. Yu TS, Dandekar M, Monteggia LM, Parada LF, Kernie SG. Genesis. 2005;41:147. [PubMed]
8. Soriano P. Nat Genet. 1999;21:70. [PubMed]
9. Cook JR, Kozak LP. Dev Biol. 1982;92:440. [PubMed]
10. Altiok S, Xu M, Spiegelman BM. Genes Dev. 1997;11:1987. [PubMed]
11. Rosen ED, MacDougald OA. Nat Rev Mol Cell Biol. 2006;7:885. [PubMed]
12. Kanda T, Sullivan KF, Wahl GM. Curr Biol. 1998;8:377. [PubMed]
13. Tumbar T, et al. Science. 2004;303:359. [PMC free article] [PubMed]
14. Otto TC, Lane MD. Crit Rev Biochem Mol Biol. 2005;40:229. [PubMed]
15. Ntambi JM, Young-Cheul K. J Nutr. 2000;130:3122S. [PubMed]
16. Graves RA, Tontonoz P, Platt KA, Ross SR, Spiegelman BM. J Cell Biochem. 1992;49:219. [PubMed]
17. Jones DL, Wagers AJ. Nat Rev Mol Cell Biol. 2008;9:11. [PubMed]
18. Armulik A, Abramsson A, Betsholtz C. Circ Res. 2005;97:512. [PubMed]
19. Dellavalle A, et al. Nat Cell Biol. 2007;9:255. [PubMed]
20. Foo SS, et al. Cell. 2006;124:161. [PubMed]
21. Boucher P, Gotthardt M, Li WP, Anderson RG, Herz J. Science. 2003;300:329. [PubMed]
22. Lehmann JM, et al. J Biol Chem. 1995;270:12953. [PubMed]
23. Kopelman PG. Nature. 2000;404:635. [PubMed]
24. Iyama K, Ohzono K, Usuku G. Virchows Arch B Cell Pathol Incl Mol Pathol. 1979;31:143. [PubMed]
25. Cinti S, Cigolini M, Bosello O, Bjorntorp P. J Submicrosc Cytol. 1984;16:243. [PubMed]
26. Rupnick MA, et al. Proc Natl Acad Sci U S A. 2002;99:10730. [PubMed]
27. Kolonin MG, Saha PK, Chan L, Pasqualini R, Arap W. Nat Med. 2004;10:625. [PubMed]
28. Kuo LE, et al. Nat Med. 2007;13:803. [PubMed]
29. Nishimura S, et al. Diabetes. 2007;56:1517. [PubMed]
30. We thank S. Kennedy, T. Wang, R. Adams, R. Evans, S. Kernie, L. Monteggia, M. Osawa, R. Perlingeiro, Q. LaPlant, M. Iacovino as well as members of the Graff lab. JMG is a founder of Reata Pharmaceuticals, a privately-held company designed to address unmet needs in cancer, neurodegenerative, and inflammatory conditions. This work is supported by the NIH and NIDDK (1R01DK064261, 1R01DK066556) and also by the UTSW Excellence in Education Fund. WT, JMG, and UTSW may file a patent application on these studies.