Logo of molcellbPermissionsJournals.ASM.orgJournalMCB ArticleJournal InfoAuthorsReviewers
Mol Cell Biol. 2007 Jan; 27(2): 743–757.
Published online 2006 Oct 23. doi:  10.1128/MCB.01557-06
PMCID: PMC1800806

Critical Role for Ebf1 and Ebf2 in the Adipogenic Transcriptional Cascade[down-pointing small open triangle]


The Ebf (O/E) family of helix-loop-helix transcription factors plays a significant role in B lymphocyte and neuronal development. The three primary members of this family, Ebf1, 2, and 3, are all expressed in adipocytes, and Ebf1 promotes adipogenesis when overexpressed in NIH 3T3 fibroblasts. Here we report that these three proteins have adipogenic potential in multiple cellular models and that peroxisome proliferator-activated receptor γ (PPARγ) is required for this effect, at least in part due to direct activation of the PPARγ1 promoter by Ebf1. Ebf1 also directly binds to and activates the C/EBPα promoter, which exerts positive feedback on C/EBPδ expression. Despite this, C/EBPα is dispensable for the adipogenic action of Ebf proteins. Ebf1 itself is induced by C/EBPβ and δ, which bind and activate its promoter. Reduction of Ebf1 and Ebf2 proteins by specific short hairpin RNA blocks differentiation of 3T3-L1 cells, suggesting a critical role for these factors and the absence of functional redundancy between members of this family. Altogether, these data place Ebf1 within the known transcriptional cascade of adipogenesis and suggest critical roles for Ebf1 and Ebf2.

The last decade has seen an enormous surge of interest in the biology of adipocytes, including the developmental processes by which these cells are formed. This has been fuelled by the convergence of a worldwide epidemic of obesity and diabetes (26, 37), with the recent realization that adipose tissue is an active secretory organ regulating a wide array of physiological processes.

Adipogenesis represents a complex series of transcriptional events through which multipotent mesenchymal precursor cells become committed to the adipocyte lineage and ultimately express all of the genes typical of mature fat cells (31). These transcriptional events integrate a variety of extracellular signals that direct fat cell formation in time and space. Most of our knowledge concerning the transcriptional events mediating adipogenesis has come from cultured cell lines such as 3T3-L1 and 3T3-F442A (16, 17) that can be differentiated into fat cells by empirically determined hormonal cocktails. These in vitro systems appear to recapitulate most of the developmental events that take place in vivo and offer the advantage of synchronous differentiation and ease of manipulation. Studies of these cells, as well as cultured mouse embryonic fibroblasts (MEFs) derived from mice with various targeted gene ablations (and more recently, studies from intact animals) have revealed a transcriptional cascade in which the nuclear hormone receptor peroxisome proliferator-activated receptor γ (PPARγ) plays a critical role. In cultured adipocytes and fat pads in vivo, PPARγ is both necessary and sufficient for adipogenesis (4, 33, 41). Other transcription factors have also been shown to play an important role in adipogenesis, including the CCAAT/enhancer binding proteins C/EBPα, C/EBPβ, and C/EBPδ and the basic helix-loop-helix protein SREBP1c (34). In 3T3-L1 cells exposed to a proadipogenic hormonal cocktail, the levels of C/EBPδ and C/EBPβ rise rapidly (6, 50). Within a few days, the levels of these proteins decrease while C/EBPα and PPARγ levels increase. C/EBPα and PPARγ sustain each other's expression through a positive feedback loop (12, 49) and remain elevated throughout the life of the terminally differentiated adipocyte, where they participate in the control of genes involved in lipogenesis, insulin sensitivity, and other pathways.

Although most studies have been devoted to the role of PPARγ and C/EBP proteins, it is known that a large number of other transcription factors and cofactors are regulated during adipogenesis. These factors may play a role in the expression of subsets of genes within the terminally differentiated adipocyte, or they may have a more fundamental impact on the process of differentiation per se. Consistent with this idea, there have been recent reports linking other transcriptional regulators to the adipogenesis cascade, such as the Krüppel-like zinc finger transcription factor family members KLF2 (which represses adipogenesis) (3) and the proadipogenic KLF5 (29) and KLF15 (27). Similarly, the zinc finger-containing factor KROX 20 (7) has been shown to participate early in the adipogenic cascade. A number of other transcription factors have also been shown to negatively regulate adipogenesis, for example, GATA-2 and -3 (39, 40), the forkhead transcription factors FoxO1 (28) and FoxA2 (47), the HMG proteins TCF/Lef (21), and SMAD-3 (8). These findings suggest that fat cell development is a more complex process than previously appreciated, requiring the integration of multiple transcriptional regulators to determine differentiation and function of the mature adipocyte.

In this report, we have focused on the adipogenic potential of a family of transcription factors known as Ebf proteins. The prototypical member of this family, Ebf1, was identified by its role in B-cell lymphopoiesis (18) and was independently identified as playing a role in the development of olfactory neurons (42). Ebf proteins have been implicated in other developmental processes as well, including bone formation (22) and neuronal differentiation (9, 14, 44). Like many other developmental regulators, the Ebf family is highly conserved evolutionarily, with orthologs in Caenorhabditis elegans and Drosophila melanogaster in addition to all vertebrates tested (11, 23). Mammals have four Ebf proteins encoded by distinct genes, designated Ebf1 through 4 (43, 45). Ebf proteins bind their cognate DNA sequences as homo- or heterodimers, the formation of which is mediated by an atypical helix-loop-helix motif. DNA binding by Ebf factors requires a zinc-coordinating motif without homology to classic zinc fingers (19).

We have shown that Ebf1, 2, and 3 are expressed in mouse and human adipose tissue as well as in 3T3-L1 adipocytes (1). More importantly, Ebf1 was shown to promote adipogenesis in 3T3-L1 cells, untransformed MEFs, and NIH 3T3 fibroblasts, while a “dominant-negative” Ebf1, consisting of the Drosophila engrailed repressor fused to the Ebf DNA binding domain, inhibited the differentiation of 3T3-L1 cells. Furthermore, transcriptional profiling in NIH 3T3 fibroblasts suggested that Ebf1 and PPARγ might induce distinct gene sets early in adipogenesis (2).

In the current set of experiments, we examine the role of other Ebf proteins in adipogenesis. Specifically, we demonstrate that multiple Ebf proteins are adipogenic in gain-of-function experiments in NIH 3T3 fibroblasts and in various MEF lines. We show that C/EBPα and PPARγ1 are direct Ebf targets and that Ebf proteins likely amplify the actions of C/EBPβ and C/EBPδ by acting immediately downstream of those factors. Finally, we use short hairpin RNA (shRNA)-mediated knockdown to show that Ebf1 and Ebf2 (but not Ebf3) are required for adipogenesis, indicating that these closely related factors have functionally nonredundant roles in this developmental process.


Cell culture.

All cells were cultured in Dulbecco's modified Eagle's medium (Invitrogen) with 10% bovine calf serum (Invitrogen) at 5% CO2. After retroviral infection (see below), cells were allow to grow to confluence in either 100-mm or 60-mm dishes in Dulbecco's modified Eagle's medium with 10% fetal bovine serum. Two days postconfluence, cells were exposed to differentiation medium containing dexamethasone (1 μM), insulin (5 μg/ml), and isobutylmethylxanthine (0.5 mM) (DMI). After 2 days, cells were maintained in medium containing insulin (5 μg/ml) until ready for harvest at 7 days. PPARγflox/− and PPARγ−/− cells and C/EBPα+/+ and C/EBPα−/− cells were generated as described previously (32, 49). Human preadipocytes were isolated and differentiated as described previously (38).

Oil-red-O staining.

After 7 to 10 days of differentiation, cells were washed once in phosphate-buffered saline and fixed with formaldehyde (Formalde-fresh; Fisher) for 15 min. The staining solution was prepared by dissolving 0.5 g oil-red-O in 100 ml of isopropanol; 60 ml of this solution was mixed with 40 ml of distilled water. After 1 h at room temperature, the staining solution was filtered and added to dishes for 4 h. The staining solution was then removed, and cells were washed twice with distilled water.

Generation of retroviral constructs and retroviral infections.

Retroviruses were constructed in pMSCV vectors (Clontech) with puromycin- or hygromycin-selectable markers. Ebf1, Ebf2, and Ebf3 cDNA were first subcloned into the pSG5-KF1M2 Flag-tag vector (gift from D. Dowhan), and then the Flag-tagged cDNAs were moved to pMSCV. Additionally, all constructs were subcloned into pCDNA3 (Invitrogen) for in vitro transcription and translation. All three Ebf constructs produced a band of the correct size upon immunoblotting with anti-Flag antibody (see Fig. S1 in the supplemental material) (Sigma). PPARγ2 pMSCV retroviral vector was constructed as described previously (32). Viral constructs (10 μg) were transfected using CellPhect transfection kit (Amersham Biosciences) into Phoenix packaging cells along with constructs encoding gag-pol (5 μg) and vesicular stomatitis virus glycoprotein G (5 μg), and supernatants were incubated in the presence of 3 μM trichostatin A (Sigma) and either used immediately or snap frozen and stored at −80°C for later use. Viral supernatants were added to the cells for 4 h in the presence of Polybrene (8 μg/ml). Selection with puromycin (4 μg/ml) or hygromycin (400 μg/ml) was started 24 h after infection and continued until cells infected with a control virus carrying no antibiotic resistance cassette were all killed (usually 4 days for puromycin and 7 days for hygromycin). Experiments were repeated three times.

Isolation of adipocytes, macrophages, and nonmacrophage SVCs from perigonadal adipose tissue.

Five-week-old male C57BL/6J mice (n = 10) were obtained from The Jackson Laboratory (Bar Harbor, MN) and were fed a standard diet (rodent diet 8664; Harlan Teklad, Madison, WI). At 19 weeks of age, mice were euthanized by CO2 inhalation, and epididymal adipose tissue (~0.5 g) was collected and placed in Krebs Ringer HEPES buffer containing 10 mg/ml fatty-acid-poor bovine serum albumin (Sigma-Aldrich, St. Louis, MO). The tissue was minced into fine pieces and centrifuged at 1,000 × g for 10 min to remove erythrocytes and other blood cells. Minced tissue was then digested in 0.12 units/ml of low-endotoxin collagenase (Liberase 3; Roche Applied Science, Indianapolis, IN) at 37°C in a shaking water bath (80 Hz) for 45 min. Samples were then filtered through a sterile 300-μm nylon mesh (Spectrum Laboratories Inc., Rancho Dominguez, CA) to remove undigested fragments. The resulting suspension was centrifuged at 500 × g for 10 min to separate stromal vascular cells (SVCs) from adipocytes. Adipocytes were removed and washed with Krebs Ringer HEPES buffer. They were then suspended in Tri reagent phenol guanidine thiocyanate solution (Molecular Research Center, Inc., Cincinnati, OH), and RNA was isolated according to the manufacturer's instructions. The SVC fraction was incubated in erythrocyte lysis buffer (0.154 mM NH4Cl, 10 mM KHCO3, 0.1 mM EDTA) for 2 min. Cells were then centrifuged at 500 × g for 5 min and resuspended in 100 μl of fluorescence-activated cell sorter buffer (phosphate-buffered saline containing 5 mM EDTA and 0.2% fatty-acid-poor bovine serum albumin). The cells were incubated in the dark on a bidirectional shaker with FcBlock (20 μg/ml; BD Pharmingen, San Jose, CA) for 30 min at 4°C. They were then incubated for 50 min with allophycocyanin-conjugated primary antibody against F4/80 (5 μg/ml; Caltag Laboratories Inc., Burlingame, CA) and phycoerythrin-conjugated antibody against CD11b (Mac-1; 2 μg/ml). F4/80 and CD11b are cell surface markers of mature macrophages (5, 25) that are not found on preadipocytes (10). Control aliquots of SVCs were incubated with allophycocyanin-labeled (2 μg/ml) and phycoerythrin-labeled (5 μg/ml) isotype control antibodies (Caltag Laboratories Inc., Burlingame, CA). After incubation, cells were washed and suspended in fluorescence-activated cell sorter buffer. F4/80+/CD11b+ macrophages and F4/80/CD11b nonmacrophage SVCs were isolated with a MoFlo (DakoCytomation, Fort Collins, CO) fluorescence-activated flow sorter. After sorting, F4/80+/CD11b+ and F4/80/CD11b cells were suspended in Trizol (Invitrogen) and RNA was isolated according to the manufacturer's instructions.

RNA extraction and quantitative PCR (Q-PCR).

Total RNA was isolated from cultured cells using Trizol reagent according to the manufacturer's instructions. First-strand cDNA was synthesized from 2 μg of total RNA and oligo(dT) primers using Moloney murine leukemia virus reverse transcriptase (RETROscript; Ambion). Real-time PCR was performed using SYBR green PCR Master Mix (Stratagene). Specific mouse primers for each gene were as follows: cyclophilin-F, 5′-GCATACAGGTCCTGGCATCT; cyclophilin-R, 5′-TTCACCTTCCCAAAGACCAC; C/EBPα-F, 5′-TGTGCGAGCACGAGACGTC; C/EBPα-R, 5′-AACTCGTCGTTGAAGGCGG; C/EBPβ-F, 5′-CTATTTCTATGAGAAAAGAGGCGTATGTAT; C/EBPβ-R, 5′-ATTCTCCCAAAAAAGTTTATTAAAATGTCT; C/EBPδ-F, 5′-TGCCCACCCTAGAGCTGTG; C/EBPδ-R, 5′-CGCTTTGTGGTTGCTGTTGA; Ebf1-F, 5′-AGGTTGGATTCTGCTACGAAACTT; Ebf1-R, 5′-TGATTCCTCTTAAAAAGGCCTGA; Ebf2-F, 5′-AGCACAAAACTACTTATTCCGATGG; Ebf2-R, 5′-GTCCAACATGGCCGCTTG; Ebf3-F, 5′-TCGTGAATATGCACCGTTTTG; Ebf3-R, 5′-ATGAGTACAGAAAAAATGTCTCGAGG; aP2-F 5′-TTCGATGAAATCACCGCAGA; aP2-R, 5′-AGGGCCCCGCCATCT; PPARγ2-F, 5′-GCATGGTGCCTTCGCTGA; PPARγ2-R, 5′-TGGCATCTCTGTGTCAACCATG; Adipsin-F, 5′-CCTTGCAATACGAGGACAAAGA; Adipsin-R, 5′-CACACCCCAACCAGCCAC; PPARγ1-F, 5′-TGAAAGAAGCGGTGAACCACTG; PPARγ1-R, 5′-TGGCATCTCTGTGTCAACCATG; CHOP-10-F, 5′-CGGAACCTGAGGAGAGAGTG; CHOP-10-R, 5′-GGACGCAGGGTCAAGAGTAG; Pref1-F, 5′-GAAATAGACGTTCGGGCTTG; Pref1-R, 5′-AGGGGTACAGCTGTTGGTTG; GATA2-F, 5′-TGCATGCAAGAGAAGTCACC; GATA2-R, 5′-ACCACCCTTGATGTCCATGT; GATA3-F, 5′-GTCATCCCTGAGCCACATCT; GATA3-R, 5′-GTAGAAGGGGTCGGAGGAAC. Primers for human genes were as follows: Ebf1-F, 5′-TTTTTCGGGTTATCGCTCAG; Ebf1-R, 5′-AATCTCCTCCCCCTTGAAAA; Ebf2-F, 5′-GGGACAAGGACAAAAGAT; Ebf2-R, 5′-TCAATTCAGGTTGCGTTT; Ebf3-F, 5′-GACCCAAATCACTGCTGGTT; Ebf3-R, 5′-TCGATTTCTGCTGCAGTTTG. All reactions were normalized to cyclophilin levels and performed in triplicate. Statistics were performed using the Student t test.

Protein extraction and Western blot analysis.

Whole-cell protein lysates were prepared according to the manufacturer's protocol using a radioimmunoprecipitation assay (RIPA) buffer containing 0.1% sodium dodecyl sulfate (Boston BioProducts) and protease inhibitor cocktail Complete (Roche). Western blot analyses were performed as described previously (20); 30 μg of total protein was loaded on a 10% sodium dodecyl sulfate-polyacrylamide gel electrophoresis gel for each sample. Mouse anti-FLAG antibody (3040; Sigma), 1/2,000 dilution, mouse anti-PPARγ antibody (SC 7273; Santa Cruz Biotechnology), 1/2,000 dilution, and anti-mouse immunoglobulin G (IgG)-peroxidase conjugate (Sigma), 1/5,000 dilution, were used to detect Flag and PPARγ proteins. SuperSignal West Pico stable peroxide solution (Pierce) was used for chemiluminescent detection.

Transfection of adipocytes.

3T3-L1 adipocytes were induced to differentiate as above. Five days after the addition of DMI, cells were transfected using the Amaxa nucleofection system (Amaxa Biosystems). A six-well plate adipocyte was transfected with 3 μg of Ebf1-pCDNA3 or empty vector along with a β-galactosidase expression vector (2 ng) as a control for transfection efficiency. Cells were harvested for RNA extraction 24 h after transfection. Statistical analyses were performed using the Mann-Whitney test.

Chromatin immunoprecipitation (ChIP) assay.

Genomic DNA was extracted, and immunoprecipitation was performed according the manufacturer's protocol (Upstate Biotechnology). Cross-linking was performed as described in the manufacturer's protocol. DNA was sheared using a sonic dismembrator model 100 (Fisher Scientific) at half maximum speed for 10 s. Immunoprecipitation was performed using 2 μg of the following antibodies: C/EBPα, C/EBPβ, and C/EBPδ (SC-61, SC-150, and SC-636; Santa Cruz Biotechnology) or anti-Flag M2 antibody (F1814; Sigma). The following primers were used to screen for C/EBP binding sites on the murine Ebf1 promoter: site A-F, 5′-TTGAATAGCCCTTAGCTGC; site A-R, 5′-GACTACAGTCCACTCCCATTG; site B-F, 5′-CCTCACCTGTACAATGG; site B-R, 5′-GCAGAGACTGCAGTCAACG; site C-F, 5′-TGTGGAACCGCACCGCG; site C-R, 5′-ATCCGGGCTTGCGACCTCG. As a negative control, we used primers to a site approximately 2 kb upstream of the transcriptional start site: F, 5′-AGCCATGAAGGGAGAGGAAT; R, 5′-CCGGTTCTGCTTTGCATACT. The following primers were used to screen for Ebf binding sites in the PPARγ1 promoter: F, 5′-TCCATGACAGACATGGACA; R, 5′-CCGGAGGAGCCGAGCCG. The upstream site was used as a negative control 2 kb from the actual site: F, 5′-CTTAATTCATATAAATAT; R, 5′-CATGTCCATGTCTGTCATG. To check the Ebf site in the C/EBPα promoter, the following primers were used: F, 5′-TCTCTCTCCACTAGCACTATGC; R, 5′-AACTGGCTCGCGCCCGCGCA. For the negative control, a site 2 kb upstream was checked with 5′-GTTTTTAGCCGTCCTCTTG (F) and 5′-GTCTCGTTCACCCTCCACCT (R) primers. A parallel immunoprecipitation with only the secondary antibody (anti-mouse IgG) or the Flag antibody on 3T3-L1 cells infected with the empty retroviruses and differentiated as mentioned above were performed.

Transactivation assays.

The C/EPBα promoter was constructed by the ligation of a 300-bp PCR fragment containing the proximal promoter (−302/+1) as a fragment into the KpnI site of pGL3 basic (Promega). The 0.8-kb Ebf1 promoter in pGL3 basic was described by Smith et al. (36). The C/EBPα, C/EBPβ, and C/EBPδ expression vectors were constructed by cloning into the pCDNA3 vector. NIH 3T3 cells were transfected at 80% confluence using Lipofectamine 2000 (Invitrogen). Transfections (200 ng of each expression construct, 500 ng of each reporter construct) were performed in triplicate and repeated three times. A β-galactosidase expression vector (2 ng) was cotransfected into cells as a control for transfection efficiency. Luciferase and β-galactosidase activity were assessed 24 h after transfection using the Galacto-Star luciferase reporter assay (Roche) according to the manufacturer's instructions. The 0.6-kb PPARγ2 promoter was the gift of Gökhan Hotamisligil (Harvard School of Public Health), while the 0.4-kb PPARγ1 promoter was the gift of Jan Reddy (Northwestern University). Statistics were performed using the Student t test.

shRNA constructs.

Target sequences for Ebf1, Ebf2, and Ebf3 were designed by querying the Whitehead small interfering RNA algorithm (http://jura.wi.mit.edu/bioc/siRNAext) as well as the small interfering RNA designer software from Clontech; at least three sequences represented by both algorithms were subcloned into the pSIREN RetroQ vector (Clontech) using the EcoRI and BamHI restriction sites. An effect had to be seen with at least two hairpins to be reported, although only data from the hairpin delivering the most efficient knockdown is shown. The best target sequences are as follows: Ebf1, GAACCCTGAAATGTGCCGA; Ebf2, CCGTTCTGTGCAGCTGGAAGGA; Ebf3, AATCACAACCAGATCCCCACC. Retroviral production and infection of 3T3-L1 cells was as described above. After selection, cells were grown to confluence and induced to differentiate as described above. Oil-red-O staining was performed at day 7. Each experiment was repeated three times.


DNA probes were labeled with [γ-32P]ATP by incubation with T4 polynucleotide kinase, annealed, and purified on a 6% polyacrylamide Tris-borate-EDTA gel. Recombinant in vitro-translated protein was incubated with labeled probe (20,000 cpm, 3 fmol) for 30 min at room temperature in binding buffer (10 mM HEPES, pH 7.9, 70 mM KCl, dithiothreitol, 1 mM EDTA, 1 mM ZnCl2, 2.3 mM MgCl2, 4% glycerol) with 0.75 μg of poly(dI/dC) per reaction mixture (Amersham). DNA competitors were added 10 min before the addition of the DNA probe. The samples were separated on a 6% polyacrylamide Tris-borate-EDTA gel, which was dried and subjected to autoradiography. Recombinant protein was generated by coupled in vitro transcription-translation using a reticulocyte lysate kit (Promega). A total of 0.5 μl of a 25-μl reaction mix was used for electrophoretic mobility shift assays (EMSAs). Oligonucleotides used for EMSA were as follows: mouse mb-1 promoter Ebf binding site forward, 5′-GAGAGAGACTCAAGGGAATTGTGG-3′; mouse mb-1 promoter Ebf binding site reverse, 5′-CCACAATTCCCTTGAGTCTCTCTC-3′; Ebf binding site mutated mouse mb-1 promoter forward, 5′-AGCCACCTCTCAGCCGTTTTGTGG-3′; Ebf binding site mutated mouse mb-1 promoter reverse, 5′-CCACAAAACGGCTGAGAGGTGGCT-3′; mouse PPARγ1 Ebf binding site forward, 5′-CCTGTAACTCCAGCTCCAGGGGAGCCCACACCT-3′; mouse PPARγ1 Ebf binding site reverse, 5′-AGGTGTGGGCTCCCCTGGAGCTGGAGTTACAGG-3′; mouse C/EBPα Ebf binding site forward, 5′-CGTTTGGACACCAGGGGGCGATGCC-3′; mouse C/EBPα Ebf binding site reverse, 5′-GGCATCGCCCCCTGGTGTCCAAACG-3′; mutated mouse C/EBPα Ebf binding site forward, 5′-CGTTTGGACACCAAGAGGCGATGCC-3′; mutated mouse C/EBPα Ebf binding site reverse, 5′-GGCATCGCCTCTTGGTGTCCAAACG-3′; mutated mouse PPARγ1 binding site forward, 5′-CCTGTAACTCCAGCTCCAAGAGAGCCCACACCT-3; mutated mouse PPARγ1 binding site reverse, 5′-AGGTGTGGGCTCTCTTGGAGCTGGAGTTACAGG-3′.

Diet-induced obesity.

Eight- to 10-week-old male C57BL/6 mice were given a high-fat, high-carbohydrate (n = 11) or normal chow (n = 5) diet for 40 days, after which epididymal fat depots were isolated. Tissues were homogenized in Trizol (Invitrogen), and RNA was isolated according to the manufacturer's recommendations. Ebf levels were measured with Q-PCR and normalized to 36B4; results were analyzed with one-way analysis of variance.


Ebf proteins are differentially regulated during adipogenesis.

To characterize the timing of Ebf induction during adipogenesis, we performed a detailed time course examination in 3T3-L1 cells. Q-PCR-based expression analysis confirmed that Ebf1, 2, and 3 are expressed during adipogenesis, but to differing degrees, such that by day 7 postdifferentiation, one sees relative levels of Ebf1 > Ebf2 > Ebf3 (Fig. (Fig.1A).1A). It is now clear that a number of factors, including CREB (51), Krox20 (7), and lipin 1 (30), are induced very early in the developmental cascade within the first few hours after DMI treatment. We looked at whether any of these Ebf isoforms were seen at these early time points and found that Ebf1 showed a profound increase in expression within 15 min of DMI treatment to levels equivalent to those seen in mature adipocytes. Ebf1 returned to baseline within 3 h but increased again gradually to reach a maximum after 7 days of differentiation. There was a tendency for a small induction in Ebf2 and Ebf3 levels immediately after DMI, but neither of these factors reached the same levels as Ebf1 during early or late time points. In general, Ebf2 increased during adipogenesis and reached a maximum after 7 days, while Ebf3 expression remained relatively low throughout the differentiation process. Levels of Ebf1 and Ebf2 rose before those of C/EBPα, PPARγ1, and PPARγ2 (Fig. (Fig.1B),1B), but the early spike in Ebf1 levels paralleled a similar rise in C/EBPβ and C/EBPδ (Fig. (Fig.1C).1C). Interestingly, Ebf1 shows a similar expression pattern when human preadipocytes are differentiated in vitro, suggesting a role for this factor in human fat development (Fig. (Fig.1D).1D). In human fat, however, Ebf3 rises during differentiation while Ebf2 levels remain low, the inverse of the pattern seen in murine 3T3-L1 cells. We also looked at relative levels of Ebf expression in fat pads from adult mice. In mature adipocytes, Ebf1 levels were highest by far, followed by Ebf2 and then Ebf3 (data not shown), similar to what was observed in mature 3T3-L1 adipocytes. Ebf1 and Ebf3 were more highly expressed in adipocytes than in cells of the stromal-vascular fraction, which includes preadipocytes, while Ebf2 levels were high even in the stromal-vascular cells (Fig. (Fig.1E).1E). This is consistent with elevated levels of Ebf2 seen in stromal cells from other sites such as bone marrow (M.S., unpublished results). Ebf isoforms were expressed at low levels in F4/80+ macrophages.

FIG. 1.
Ebf isoforms are induced early and late during 3T3-L1 adipogenesis. (A) Levels of Ebf isoforms measured by Q-PCR in the first few hours (15 and 30 min and 1, 3, 6, and 12 h) after DMI treatment (left) and over the next several days (1 to 7 days, right). ...

Ectopic expression of Ebf proteins in NIH 3T3 cells promotes adipogenesis.

We next sought to determine which Ebf proteins can promote fat differentiation in nonadipogenic NIH 3T3 fibroblasts. These cells were infected with retroviruses expressing Ebf1, Ebf2, Ebf3, PPARγ2, or empty vector. After selection with puromycin, cells were grown to confluence and induced to differentiate with DMI. Seven days after induction, cells were stained for neutral lipid with oil-red-O (Fig. (Fig.2A).2A). Adipocytes were easily detected in dishes containing cells expressing any Ebf isoform (representing 10 to 15% of all cells), while vector-infected cells exhibited no staining. As previously shown, cells overexpressing PPARγ2 differentiate with very high efficiency (~90%). Consistent with the oil-red-O staining, expression of fat-specific genes was elevated in Ebf-expressing cells (Fig. (Fig.2B2B).

FIG. 2.
Ectopic expression of Ebf isoforms in NIH 3T3 cells promotes adipogenesis. (A) Micrographs of oil-red-O-stained NIH 3T3 cells 7 days after induction of differentiation with DMI. Cells were transduced with retroviruses expressing either Ebf1, Ebf2, Ebf3, ...

PPARγ1 and C/EBPα are direct targets of Ebf1.

PPARγ is the most potent adipogenic transcription factor discovered to date and is required for fat cell differentiation in vitro and in vivo. We sought to determine whether Ebf proteins could induce adipogenesis in the absence of PPARγ by using immortalized PPARγ−/− MEFs (32). Retrovirally mediated expression of Ebf1, Ebf2, or Ebf3 (see Fig. S2A in the supplemental material) was unable to induce differentiation in these cells, as measured by lipid accumulation (Fig. (Fig.3A)3A) or by marker expression (see Fig. S2B in the supplemental material). In contrast, control fibroblasts containing a single intact PPARγ allele (PPARγflox/−) were differentiated by all three Ebf proteins. As expected, retroviral introduction of PPARγ2 allowed robust differentiation of both cell types.

PPARγ1 and C/EBPα are direct targets of Ebf1. (A) Immortalized PPARγ−/− and PPARγflox/− MEFs were infected with retroviruses expressing either Ebf1, Ebf2, Ebf3, PPARγ2, or empty vector. Cells ...

To assess whether Ebf proteins directly induce PPARγ expression, we looked at their effect of Ebf1 on the 0.6-kb PPARγ2 promoter and the 0.4-kb PPARγ1 promoter in transient-transfection assays. No effect was seen on the PPARγ2 promoter construct (data not shown), but Ebf1 induced expression from the PPARγ1 promoter 2.3-fold (Fig. (Fig.3B).3B). We also looked at a possible effect of Ebf1 on the 0.3-kb C/EBPα promoter. Ebf1 increased luciferase expression in this assay by 4.5-fold above baseline (Fig. (Fig.3C),3C), demonstrating a potential direct transcriptional link between these two adipogenic factors. Putative Ebf binding sites within the C/EBPα and PPARγ1 promoters were identified and aligned with the consensus Ebf site as well as a well-characterized Ebf binding site from the mouse immunoglobulin α-associated protein (mb-1) promoter (15, 35) (Fig. (Fig.3D).3D). EMSA confirmed that these putative sites could effectively compete Ebf1 binding from the mb-1 oligonucleotide (Fig. (Fig.3D).3D). Oligonucleotides bearing point mutations in the Ebf binding site of the C/EBPα and PPARγ1 promoters; however, do not compete for binding to the mb-1 promoter (Fig. (Fig.3D).3D). To assess Ebf binding to the native promoters, we also performed ChIP assays in 3T3-L1 cells transduced with Flag-tagged Ebf1 (Fig. (Fig.3E).3E). Ebf1 is bound to the PPARγ1 and C/EBPα promoters even prior to differentiation; this binding increases strongly by 1 h after DMI exposure before gradually decreasing at later time points. This peak at 1 h coincides with the period of maximum endogenous Ebf1 expression in 3T3-L1 differentiation (Fig. (Fig.1A),1A), strongly suggesting that the interaction is functionally relevant. Further support for this notion was obtained by showing that retroviral introduction of Ebf1 into PPARγflox/− fibroblasts causes a large induction in endogenous C/EBPα levels relative to vector-infected control cells (Fig. (Fig.3F).3F). This difference is seen both before and after induction of differentiation with DMI. We did not see a similar effect with PPARγ1, which rose after DMI to an equivalent degree in vector- and Ebf1-infected cells (data not shown). Although the reason for this is unclear, we decided to test whether having a full complement of PPARγ might enhance a direct effect of Ebf1 on PPARγ1. We therefore transfected mature 3T3-L1 adipocytes with an Ebf1 expression plasmid and found enhanced expression of both endogenous C/EBPα and PPARγ1 mRNA (Fig. (Fig.3G3G).

Ebf1 induces C/EBPδ expression via C/EBPα.

Interestingly, levels of C/EBPα and C/EBPδ were elevated by Ebf1 overexpression in both PPARγflox/− and PPARγ−/− fibroblasts (Fig. (Fig.4A)4A) but not in NIH 3T3 cells (Fig. (Fig.4B).4B). NIH 3T3 cells express low levels of C/EBPα during forced differentiation, in contrast to most other models of adipogenesis. This fact, in concert with the coincident rise of C/EBPδ and Ebf1 within the first hour after DMI (Fig. 1A and C), led us to consider whether a feedback loop might exist in which Ebf1 induces C/EBPδ levels in a C/EBPα-dependent manner. C/EBPα−/− MEFs were employed to test whether genetic ablation of C/EBPα could prevent Ebf1 from inducing C/EBPδ. These cells, along with C/EBPα+/+ control MEFs, were transduced with an Ebf1-expressing retrovirus (see Fig. S3A in the supplemental material) and harvested just prior to the induction of differentiation (i.e., on day 0). C/EBPδ was induced by Ebf1 in C/EBPα+/+ control cells but not in C/EBPα−/− cells (Fig. (Fig.4C).4C). The same pattern was seen when Ebf2 or Ebf3 was expressed in these cells (data not shown). C/EBPβ was not induced by Ebf1 in any cellular context. Taken together, these data strongly suggest a cascade in which Ebf1 might participate in the very early stages of adipogenesis by promoting C/EBPδ expression via C/EBPα. Surprisingly, the total absence of C/EBPα did not prevent fat cell formation in the presence of Ebf1, 2, or 3, as measured by lipid accumulation (Fig. (Fig.4D)4D) or by marker expression (see Fig. S3B in the supplemental material). This suggests that Ebf proteins, at least in the setting of ectopic overexpression, can promote adipogenesis without inducing C/EBPδ through C/EBPα.

Ebf1 induces C/EBPδ expression in a C/EBPα-dependent manner, but does not require C/EBPα to promote adipogenesis. (A) C/EBP isoform expression in PPARγflox/− and PPARγ−/− cells transduced ...

C/EBPβ and C/EBPδ promote the induction of Ebf1 during adipogenesis.

C/EBPβ and δ are among the earliest actors in the adipogenic transcriptional cascade. We noted that the sharp early rise in Ebf1 was contemporaneous with a spike in these two factors, and the more gradual elevation in Ebf1 and Ebf2 that occurred over the first few days lies between the peak expression of C/EBPβ and δ on the one hand and PPARγ and C/EBPα on the other (Fig. (Fig.1).1). We thus were interested in the possibility that C/EBPβ and/or δ could be a direct inducer of Ebf expression. To address this, we first tested whether C/EBP proteins could induce expression from an Ebf1 promoter-luciferase construct. We found that C/EBPα, β, and δ were all able to induce expression of this construct (Fig. (Fig.5A).5A). We then identified three potential C/EBP binding elements in this construct using TRANSFAC, designated in Fig. Fig.5A5A as sites A, B, and C. A combination of deletion mapping and transactivation analysis was employed to assess which of these sites might be required for C/EBP action on the Ebf1 promoter. Removal of site A reduced the ability of C/EBPα and δ to induce the reporter but did not affect the ability of C/EBPβ to do so. Subsequent removal of site B did not affect transactivation in response to any of the C/EBPs, but removal of site C (in addition to A and B) prevented all C/EBP action on the basal promoter. These results suggest that all three C/EBP proteins can transactivate the C site but that the A site is transactivated only by C/EBPα or δ.

FIG. 5.
C/EBPβ and δ promote induction of Ebf1. (A) Deletion analysis of Ebf1 promoter was performed to identify potential C/EBP sites. Three putative binding sites for C/EBPs (black boxes A, B, C) and two previously identified transcriptional ...

To confirm that C/EBP proteins occupy the Ebf1 promoter during adipogenesis, we performed ChIP assays at various time points during 3T3-L1 differentiation (Fig. (Fig.5B).5B). C/EBPβ binds site A from day 1 through day 4 and is thereafter replaced by C/EBPα, whereas C/EBPδ does not bind site A. In contrast, site B does not seem to be regulated because all C/EBP proteins occupy this site throughout the differentiation process. Similarly, binding of site C by C/EBPα and δ is not regulated. No binding was seen to a site 2 kb upstream of site A.

Ebf1 and Ebf2, but not Ebf3, are necessary for adipogenesis.

Three Ebf proteins induced adipogenesis in at least four independently derived cell lines (NIH 3T3, PPARγflox/− MEFs, C/EBPα+/+ MEFs, and C/EBPα−/− MEFs). These results could imply that each Ebf isoform exerts a nonredundant action on differentiation. Alternatively, it could be the case that only one or two proteins are required but that any isoform can drive the process forward when overexpressed because of sufficient similarity in their DNA binding activity. To distinguish between these scenarios, we used retroviral delivery of shRNAs to knock down individual Ebf proteins. We first confirmed the efficacy and specificity of each hairpin by cotransfecting NIH 3T3 cells with Flag-tagged Ebf1, Ebf2, or Ebf3 along with the retroviral vector containing the shRNA against each isoform. The absence of adequate antibodies against individual Ebf proteins made this approach necessary. Twenty-four hours after transfection, cell lysates were harvested and analyzed by immunoblotting with an anti-Flag antibody. As shown in Fig. S4 in the supplemental material, we were able to identify hairpins that knocked down individual Ebf proteins at the protein level and that exhibited no cross-reactivity with other Ebfs. We next infected 3T3-L1 preadipocytes with the shRNA-expressing retroviruses. Transduced cells were selected and expanded, and expression of the endogenous Ebf genes was determined by quantitative real-time PCR on the day differentiation was induced. As seen in Fig. Fig.6A,6A, high-efficiency and highly selective knockdown was achieved in these cells as well. At 7 days postdifferentiation, cells were either stained with oil-Red-O or RNA was harvested. As expected, 3T3-L1 cells infected with a control shRNA are able to differentiate fully into adipocytes. In contrast, knockdown of either Ebf1 (Fig. (Fig.6B)6B) or Ebf2 (Fig. (Fig.6C)6C) completely abolished the differentiation process. Cells were cultured for more than 10 days to confirm that differentiation was not simply delayed. Interestingly, knockdown of Ebf3 did not have any effect on the number and appearance of fat cells relative to control cells (Fig. (Fig.6D).6D). These results were confirmed by measuring adipose-selective markers like aP2 and adipsin at day 7. We thus demonstrate a requirement for Ebf1 and Ebf2 in adipogenesis and suggest that these two proteins play nonredundant roles in the differentiation process. Ebf3, in contrast, seems to be less important, consistent with the fact that endogenous levels of this isoform are hardly induced by differentiation at all in these cells (Fig. (Fig.1A1A).

FIG. 6.
Ebf1 and Ebf2 are required for adipogenesis. (A) 3T3-L1 preadipocytes were transduced with constructs containing shRNAs directed against luciferase (control), Ebf1, Ebf2, or Ebf3. Levels of Ebf1, Ebf2, and Ebf3 mRNA were measured by Q-PCR prior to differentiation. ...

Ebf expression is dysregulated in obesity.

The importance of Ebf proteins for adipocyte differentiation in vitro led us to investigate whether levels of these factors are dysregulated in obesity. As shown in Fig. Fig.7,7, Ebf1, 2, and 3 mRNAs are highly induced in the white adipose tissue of mice on a high-fat, high-carbohydrate diet relative to lean controls.

FIG. 7.
Ebf1, 2, and 3 mRNA levels are induced in white adipose tissue in mice on a high-fat, high-carbohydrate (HF/HC) diet. After 40 days of chow (n = 5) or HF/HC (n = 11) diet, mRNA was isolated from epididymal fat pads from male C57BL/6 mice ...


Cell fates are determined by a complex series of transcriptional events that promote terminal differentiation and concomitantly suppress alternate possible lineages. In the case of adipogenesis, extensive work from a number of groups has identified a key role for several transcription factors, most notably PPARγ and the C/EBP proteins α, β, and δ (34). Based on these studies, a model has been proposed wherein C/EBPβ and δ are among the earliest factors to respond to adipogenic stimuli in a committed precursor cell, and these in turn induce expression of PPARγ and C/EBPα (Fig. (Fig.8)8) (12, 34, 48). These latter factors appear to mutually reinforce each other's expression, and both promote the establishment of the ultimate terminally differentiated phenotype. PPARγ and C/EBPα are not equals in this process, however, as ectopic expression of PPARγ in C/EBPα null fibroblasts rescues adipogenesis, while the converse is not true (32, 49).

FIG. 8.
Model of adipogenesis. A model placing Ebf1 into the context of the known adipogenic transcriptional cascade. See the text for details.

In the last few years, several new transcription factors have been discovered to play a role in adipogenesis. Ebf1 was also recently shown to play a positive role in this process (1), although it has not been clear in which part of the cascade Ebf1 acts. Furthermore, other Ebf family members, including Ebf2 and Ebf3, are expressed in adipose tissue but had not been tested for a potential role in the developmental process.

We have addressed these issues using a variety of gain-of-function and loss-of-function approaches in multiple adipogenic and nonadipogenic cell types. Our results indicate that there is a short but intense burst of Ebf1 expression within hours of adding the proadipogenic cocktail, followed by a return to a baseline and a more gradual sustained rise that appears to be maintained for the life of the mature adipocyte. Ebf2 and Ebf3 do not display this sharp rise in the early hours of differentiation, and although they do rise gradually later in development, they never reach peak levels equivalent to that of Ebf1.

These factors promote adipogenesis in NIH 3T3 fibroblasts, placing the Ebf family in a fairly elite class of factors that have this activity. Given the high degree of structural similarity within the Ebf family, however, it was not clear if the three factors tested were acting in a generic “Ebf-like” way when overexpressed or whether each factor had a unique function. To address this, we employed retroviral delivery of shRNA to specifically “knock down” each Ebf isoform in turn. To our surprise, differentiation did not proceed in the presence of reduced levels of either Ebf1 or Ebf2, strongly suggesting that these two factors possess nonredundant activities. Specific Ebf3 “knockdown” did not result in changes in lipid accumulation or expression of marker genes like aP2. These experiments establish the importance of Ebf1 and Ebf2 and inform us that these proteins play a role in the process that cannot be adequately compensated for by other members of the family.

Where can we place Ebf action in the adipogenic cascade? Temporally, they appear at or after the rise in C/EBPβ and δ, suggesting that they might themselves be induced by these factors. This was made clear for Ebf1 in particular by the demonstration of C/EBP binding sites in the proximal promoter. ChIP analysis suggests that C/EBPs may occupy the site A in a differentiation-dependent manner. In contrast, sites B and C are occupied by C/EBPs even prior to the induction of differentiation, suggesting an additional permissive role for C/EBPs in the induction of Ebf1, in which binding to some sites is required but not sufficient for induction of Ebf1 message levels. Alternatively, the C/EBPs occupying site B and C might be regulated by posttranslational modification that alters their capacity to activate the Ebf1 promoter during adipogenesis.

Furthermore, our data suggest that Ebfs promote adipogenesis, at least in part, by inducing expression of C/EBPα and PPARγ1. The evidence that these factors are direct targets of Ebf1 include (i) the presence of Ebf motifs in the PPARγ1 and C/EBPα promoters, both of which bind Ebf1 in vitro, (ii) the ability of these sites to enable reporter gene transactivation by Ebf1 in a transient-transfection assay, (iii) direct binding of Ebf1 to these sites in the native promoter in intact cells in a differentiation-dependent manner, (iv) enhancement of endogenous PPARγ1 and C/EBPα expression in adipocytes transfected with Ebf1, and (v) a requirement for PPARγ in the adipogenic action of Ebf1.

Our experiments place Ebf action between C/EBPβ/δ and C/EBPα/PPARγ, although they do not rule out further downstream actions of Ebf proteins that might also be critical for differentiation. In fact, the ability of Ebf proteins to induce adipogenesis in the absence of C/EBPα (as shown in C/EBPα−/− MEFs as well as in NIH 3T3 cells, which express very low levels of C/EBPα) suggests that other targets are likely to exist. This is reminiscent of the situation in which ectopic addition of PPARγ allows differentiation in the absence of C/EBPα and supports the notion that C/EBPα is more ancillary to the process than PPARγ. Adipocytes differentiated in the absence of C/EBPα accumulate lipid and most markers of the fat phenotype but are not insulin sensitive (13, 34). This is the result of improper Glut4 vesicle trafficking in C/EBPα null adipocytes (46). We do not know yet if this will also be the case in C/EBPα null adipocytes differentiated by ectopic overexpression of Ebf proteins.

Our data also identify a positive feedback loop involving C/EBPδ, Ebf1, and C/EBPα. This was initially suggested by the fact that Ebf1 could induce expression of C/EBPδ in PPARγflox/− and PPARγ−/− cells but not in NIH 3T3 cells, which do not express C/EBPα. The requirement for C/EBPα was confirmed by showing that Ebf1 induces C/EBPδ in C/EBPα+/+ MEFs but not in MEFs lacking C/EBPα. Ebf1 likely exerts this effect by direct transactivation of the C/EBPα promoter. We do note, however, that the Ebf1-C/EBPα-C/EBPδ-positive feedback loop is not absolutely required for adipogenesis, as Ebf1 is perfectly capable of causing fat cell formation in the absence of C/EBPα. We postulate that this feedback loop can act as an accelerant during the early phases of differentiation in normal cells.

These experiments demonstrate a critical role for several different Ebf family members in adipogenesis. Ebf1 and Ebf2 in particular are shown to be both necessary and sufficient for adipogenesis in vitro. The situation with Ebf3 is less clear, in that it promotes differentiation quite well when overexpressed, but its absence has no significant effect on terminal differentiation. Ebf2 and Ebf3 are extraordinarily highly conserved (>85% at the amino acid level), with Ebf2 possessing two short novel sequences not seen in Ebf3, including one in the C-terminal transactivation domain. One possible explanation for our results is that Ebf2 can compensate for the loss of Ebf3, but the opposite is not true because of effects mediated by the Ebf2 unique C-terminal sequence. Further studies involving domain swapping will be required to clarify this point.

Much remains to be learned about Ebf isoform action in adipocyte biology. A key question centers on additional target genes of Ebf proteins during development and in the mature adipocyte. Other developmentally relevant targets are likely, as discussed above, and could include other known adipogenic transcription factors besides PPARγ1 and C/EBPα. PPARγ2 may be such a target. Our studies revealed no Ebf binding sites in the 0.6-kb PPARγ2 promoter, but certainly sites could exist outside of that region. Ebf proteins may also affect antiadipogenic pathways. Fig. S5 in the supplemental material shows the effect of Ebf1 to 3 on the mRNA level of several factors known to repress adipogenesis in transduced NIH 3T3 cells prior to differentiation. The ability of Ebf1 and Ebf2 to repress GATA2 is certainly interesting in this regard.

It is noteworthy that a statistical analysis of overrepresented motifs in the 1-kb promoter regions from 30 adipose-selective genes identified variants of the Ebf consensus sequence as the top-rated hit (data not shown), suggesting that these factors may play a broad role in establishing and maintaining the mature adipose phenotype. Furthermore, the fact that Ebf1, 2, and 3 are elevated in adipose tissue from obese mice suggests a possible role for these proteins in the pathological behavior of adipocytes in the setting of overnutrition. It will be interesting to look at Ebf expression and function in other obese and diabetic models. Ebf1 null mice have been generated and have been reported to be smaller than wild-type littermates, but no specific measurements of body fat or other metabolic parameters were commented upon (14, 24). Ebf3 null animals exhibit perinatal lethality, while Ebf2 null mice have reduced survival and significant neuronal defects (9, 44). We are in the process of characterizing these and other models for metabolic and adipose-related phenotypes. Given the clinical success of PPARγ-based therapies, we will be interested to see if manipulation of Ebf proteins might provide additional avenues in the treatment of metabolic disease.

Supplementary Material

[Supplemental material]


This work was supported by NIH grant DK63906 to E.D.R., Swiss National Foundation award PA00A-105047 to M.A.J., and funds from Astra Zeneca.

We thank James Kirkland at the Adipocyte Core, Boston Obesity Nutrition Research Center, NIH grant DK 46200, for the human adipocyte RNA samples; Karen Inouye for cDNA from isolated murine adipocytes, macrophages, and stromal vascular cells; and Simonetta Westerlund and Magdalen Rhedin for analysis of EBF expression in mice. We also thank members of the Rosen lab for thoughtful discussion.


[down-pointing small open triangle]Published ahead of print on 23 October 2006.

Supplemental material for this article may be found at http://mcb.asm.org/.


1. Akerblad, P., U. Lind, D. Liberg, K. Bamberg, and M. Sigvardsson. 2002. Early B-cell factor (O/E-1) is a promoter of adipogenesis and involved in control of genes important for terminal adipocyte differentiation. Mol. Cell. Biol. 22:8015-8025. [PMC free article] [PubMed]
2. Akerblad, P., R. Mansson, A. Lagergren, S. Westerlund, B. Basta, U. Lind, A. Thelin, R. Gisler, D. Liberg, S. Nelander, K. Bamberg, and M. Sigvardsson. 2005. Gene expression analysis suggests that EBF-1 and PPARgamma2 induce adipogenesis of NIH-3T3 cells with similar efficiency and kinetics. Physiol. Genomics 23:206-216. [PubMed]
3. Banerjee, S. S., M. W. Feinberg, M. Watanabe, S. Gray, R. L. Haspel, D. J. Denkinger, R. Kawahara, H. Hauner, and M. K. Jain. 2003. The Kruppel-like factor KLF2 inhibits peroxisome proliferator-activated receptor-gamma expression and adipogenesis. J. Biol. Chem. 278:2581-2584. [PubMed]
4. Barak, Y., M. C. Nelson, E. S. Ong, Y. Z. Jones, P. Ruiz-Lozano, K. R. Chien, A. Koder, and R. M. Evans. 1999. PPAR gamma is required for placental, cardiac, and adipose tissue development. Mol. Cell 4:585-595. [PubMed]
5. Buckley, P. J., M. R. Smith, M. F. Braverman, and S. A. Dickson. 1987. Human spleen contains phenotypic subsets of macrophages and dendritic cells that occupy discrete microanatomic locations. Am. J. Pathol. 128:505-520. [PMC free article] [PubMed]
6. Cao, Z., R. M. Umek, and S. L. McKnight. 1991. Regulated expression of three C/EBP isoforms during adipose conversion of 3T3-L1 cells. Genes Dev. 5:1538-1552. [PubMed]
7. Chen, Z., J. I. Torrens, A. Anand, B. M. Spiegelman, and J. M. Friedman. 2005. Krox20 stimulates adipogenesis via C/EBPbeta-dependent and -independent mechanisms. Cell Metab. 1:93-106. [PubMed]
8. Choy, L., and R. Derynck. 2003. Transforming growth factor-beta inhibits adipocyte differentiation by Smad3 interacting with CCAAT/enhancer-binding protein (C/EBP) and repressing C/EBP transactivation function. J. Biol. Chem. 278:9609-9619. [PubMed]
9. Corradi, A., L. Croci, V. Broccoli, S. Zecchini, S. Previtali, W. Wurst, S. Amadio, R. Maggi, A. Quattrini, and G. G. Consalez. 2003. Hypogonadotropic hypogonadism and peripheral neuropathy in Ebf2-null mice. Development 130:401-410. [PubMed]
10. Cousin, B., O. Munoz, M. Andre, A. M. Fontanilles, C. Dani, J. L. Cousin, P. Laharrague, L. Casteilla, and L. Penicaud. 1999. A role for preadipocytes as macrophage-like cells. FASEB J. 13:305-312. [PubMed]
11. Dubois, L., and A. Vincent. 2001. The COE-Collier/Olf1/EBF-transcription factors: structural conservation and diversity of developmental functions. Mech. Dev. 108:3-12. [PubMed]
12. Elberg, G., J. M. Gimble, and S. Y. Tsai. 2000. Modulation of the murine peroxisome proliferator-activated receptor gamma 2 promoter activity by CCAAT/enhancer-binding proteins. J. Biol. Chem. 275:27815-27822. [PubMed]
13. El-Jack, A. K., J. K. Hamm, P. F. Pilch, and S. R. Farmer. 1999. Reconstitution of insulin-sensitive glucose transport in fibroblasts requires expression of both PPARgamma and C/EBPalpha. J. Biol. Chem. 274:7946-7951. [PubMed]
14. Garel, S., K. Yun, R. Grosschedl, and J. L. Rubenstein. 2002. The early topography of thalamocortical projections is shifted in Ebf1 and Dlx1/2 mutant mice. Development 129:5621-5634. [PubMed]
15. Gisler, R., S. E. Jacobsen, and M. Sigvardsson. 2000. Cloning of human early B-cell factor and identification of target genes suggest a conserved role in B-cell development in man and mouse. Blood 96:1457-1464. [PubMed]
16. Green, H., and O. Kehinde. 1975. An established preadipose cell line and its differentiation in culture. II. Factors affecting the adipose conversion. Cell 5:19-27. [PubMed]
17. Green, H., and O. Kehinde. 1976. Spontaneous heritable changes leading to increased adipose conversion in 3T3 cells. Cell 7:105-113. [PubMed]
18. Hagman, J., C. Belanger, A. Travis, C. W. Turck, and R. Grosschedl. 1993. Cloning and functional characterization of early B-cell factor, a regulator of lymphocyte-specific gene expression. Genes Dev. 7:760-773. [PubMed]
19. Hagman, J., M. J. Gutch, H. Lin, and R. Grosschedl. 1995. EBF contains a novel zinc coordination motif and multiple dimerization and transcriptional activation domains. EMBO J. 14:2907-2916. [PMC free article] [PubMed]
20. Jimenez, M., C. Yvon, L. Lehr, B. Leger, P. Keller, A. Russell, F. Kuhne, P. Flandin, J. P. Giacobino, and P. Muzzin. 2002. Expression of uncoupling protein-3 in subsarcolemmal and intermyofibrillar mitochondria of various mouse muscle types and its modulation by fasting. Eur. J. Biochem. 269:2878-2884. [PubMed]
21. Kennell, J. A., E. E. O'Leary, B. M. Gummow, G. D. Hammer, and O. A. MacDougald. 2003. T-cell factor 4N (TCF-4N), a novel isoform of mouse TCF-4, synergizes with beta-catenin to coactivate C/EBPalpha and steroidogenic factor 1 transcription factors. Mol. Cell. Biol. 23:5366-5375. [PMC free article] [PubMed]
22. Kieslinger, M., S. Folberth, G. Dobreva, T. Dorn, L. Croci, R. Erben, G. G. Consalez, and R. Grosschedl. 2005. EBF2 regulates osteoblast-dependent differentiation of osteoclasts. Dev. Cell 9:757-767. [PubMed]
23. Liberg, D., M. Sigvardsson, and P. Akerblad. 2002. The EBF/Olf/Collier family of transcription factors: regulators of differentiation in cells originating from all three embryonal germ layers. Mol. Cell. Biol. 22:8389-8397. [PMC free article] [PubMed]
24. Lin, H., and R. Grosschedl. 1995. Failure of B-cell differentiation in mice lacking the transcription factor EBF. Nature 376:263-267. [PubMed]
25. McKnight, A. J., A. J. Macfarlane, P. Dri, L. Turley, A. C. Willis, and S. Gordon. 1996. Molecular cloning of F4/80, a murine macrophage-restricted cell surface glycoprotein with homology to the G-protein-linked transmembrane 7 hormone receptor family. J. Biol. Chem. 271:486-489. [PubMed]
26. Mokdad, A. H., E. S. Ford, B. A. Bowman, W. H. Dietz, F. Vinicor, V. S. Bales, and J. S. Marks. 2003. Prevalence of obesity, diabetes, and obesity-related health risk factors, 2001. JAMA 289:76-79. [PubMed]
27. Mori, T., H. Sakaue, H. Iguchi, H. Gomi, Y. Okada, Y. Takashima, K. Nakamura, T. Nakamura, T. Yamauchi, N. Kubota, T. Kadowaki, Y. Matsuki, W. Ogawa, R. Hiramatsu, and M. Kasuga. 2005. Role of Kruppel-like factor 15 (KLF15) in transcriptional regulation of adipogenesis. J. Biol. Chem. 280:12867-12875. [PubMed]
28. Nakae, J., T. Kitamura, Y. Kitamura, W. H. Biggs III, K. C. Arden, and D. Accili. 2003. The forkhead transcription factor Foxo1 regulates adipocyte differentiation. Dev. Cell 4:119-129. [PubMed]
29. Oishi, Y., I. Manabe, K. Tobe, K. Tsushima, T. Shindo, K. Fujiu, G. Nishimura, K. Maemura, T. Yamauchi, N. Kubota, R. Suzuki, T. Kitamura, S. Akira, T. Kadowaki, and R. Nagai. 2005. Kruppel-like transcription factor KLF5 is a key regulator of adipocyte differentiation. Cell Metab. 1:27-39. [PubMed]
30. Phan, J., M. Peterfy, and K. Reue. 2004. Lipin expression preceding peroxisome proliferator-activated receptor-gamma is critical for adipogenesis in vivo and in vitro. J. Biol. Chem. 279:29558-29564. [PubMed]
31. Pittenger, M. F., A. M. Mackay, S. C. Beck, R. K. Jaiswal, R. Douglas, J. D. Mosca, M. A. Moorman, D. W. Simonetti, S. Craig, and D. R. Marshak. 1999. Multilineage potential of adult human mesenchymal stem cells. Science 284:143-147. [PubMed]
32. Rosen, E. D., C. H. Hsu, X. Wang, S. Sakai, M. W. Freeman, F. J. Gonzalez, and B. M. Spiegelman. 2002. C/EBPalpha induces adipogenesis through PPARgamma: a unified pathway. Genes Dev. 16:22-26. [PMC free article] [PubMed]
33. Rosen, E. D., P. Sarraf, A. E. Troy, G. Bradwin, K. Moore, D. S. Milstone, B. M. Spiegelman, and R. M. Mortensen. 1999. PPAR gamma is required for the differentiation of adipose tissue in vivo and in vitro. Mol. Cell 4:611-617. [PubMed]
34. Rosen, E. D., C. J. Walkey, P. Puigserver, and B. M. Spiegelman. 2000. Transcriptional regulation of adipogenesis. Genes Dev. 14:1293-1307. [PubMed]
35. Sigvardsson, M., D. R. Clark, D. Fitzsimmons, M. Doyle, P. Akerblad, T. Breslin, S. Bilke, R. Li, C. Yeamans, G. Zhang, and J. Hagman. 2002. Early B-cell factor, E2A, and Pax-5 cooperate to activate the early B cell-specific mb-1 promoter. Mol. Cell. Biol. 22:8539-8551. [PMC free article] [PubMed]
36. Smith, E. M., R. Gisler, and M. Sigvardsson. 2002. Cloning and characterization of a promoter flanking the early B cell factor (EBF) gene indicates roles for E-proteins and autoregulation in the control of EBF expression. J. Immunol. 169:261-270. [PubMed]
37. Strauss, R. S., and H. A. Pollack. 2001. Epidemic increase in childhood overweight, 1986-1998. JAMA 286:2845-2848. [PubMed]
38. Tchkonia, T., Y. D. Tchoukalova, N. Giorgadze, T. Pirtskhalava, I. Karagiannides, R. A. Forse, A. Koo, M. Stevenson, D. Chinnappan, A. Cartwright, M. D. Jensen, and J. L. Kirkland. 2005. Abundance of two human preadipocyte subtypes with distinct capacities for replication, adipogenesis, and apoptosis varies among fat depots. Am. J. Physiol. Endocrinol. Metab. 288:E267-E277. [PubMed]
39. Tong, Q., G. Dalgin, H. Xu, C. N. Ting, J. M. Leiden, and G. S. Hotamisligil. 2000. Function of GATA transcription factors in preadipocyte-adipocyte transition. Science 290:134-138. [PubMed]
40. Tong, Q., J. Tsai, G. Tan, G. Dalgin, and G. S. Hotamisligil. 2005. Interaction between GATA and the C/EBP family of transcription factors is critical in GATA-mediated suppression of adipocyte differentiation. Mol. Cell. Biol. 25:706-715. [PMC free article] [PubMed]
41. Tontonoz, P., E. Hu, and B. M. Spiegelman. 1994. Stimulation of adipogenesis in fibroblasts by PPAR gamma 2, a lipid-activated transcription factor. Cell 79:1147-1156. [PubMed]
42. Wang, M. M., and R. R. Reed. 1993. Molecular cloning of the olfactory neuronal transcription factor Olf-1 by genetic selection in yeast. Nature 364:121-126. [PubMed]
43. Wang, S. S., A. G. Betz, and R. R. Reed. 2002. Cloning of a novel Olf-1/EBF-like gene, O/E-4, by degenerate oligo-based direct selection. Mol. Cell. Neurosci. 20:404-414. [PubMed]
44. Wang, S. S., J. W. Lewcock, P. Feinstein, P. Mombaerts, and R. R. Reed. 2004. Genetic disruptions of O/E2 and O/E3 genes reveal involvement in olfactory receptor neuron projection. Development 131:1377-1388. [PubMed]
45. Wang, S. S., R. Y. Tsai, and R. R. Reed. 1997. The characterization of the Olf-1/EBF-like HLH transcription factor family: implications in olfactory gene regulation and neuronal development. J. Neurosci. 17:4149-4158. [PubMed]
46. Wertheim, N., Z. Cai, and T. E. McGraw. 2004. The transcription factor CCAAT/enhancer-binding protein alpha is required for the intracellular retention of GLUT4. J. Biol. Chem. 279:41468-41476. [PubMed]
47. Wolfrum, C., D. Q. Shih, S. Kuwajima, A. W. Norris, C. R. Kahn, and M. Stoffel. 2003. Role of Foxa-2 in adipocyte metabolism and differentiation. J. Clin. Investig. 112:345-356. [PMC free article] [PubMed]
48. Wu, Z., N. L. Bucher, and S. R. Farmer. 1996. Induction of peroxisome proliferator-activated receptor gamma during the conversion of 3T3 fibroblasts into adipocytes is mediated by C/EBPbeta, C/EBPdelta, and glucocorticoids. Mol. Cell. Biol. 16:4128-4136. [PMC free article] [PubMed]
49. Wu, Z., E. D. Rosen, R. Brun, S. Hauser, G. Adelmant, A. E. Troy, C. McKeon, G. J. Darlington, and B. M. Spiegelman. 1999. Cross-regulation of C/EBP alpha and PPAR gamma controls the transcriptional pathway of adipogenesis and insulin sensitivity. Mol. Cell 3:151-158. [PubMed]
50. Yeh, W. C., Z. Cao, M. Classon, and S. L. McKnight. 1995. Cascade regulation of terminal adipocyte differentiation by three members of the C/EBP family of leucine zipper proteins. Genes Dev. 9:168-181. [PubMed]
51. Zhang, J. W., D. J. Klemm, C. Vinson, and M. D. Lane. 2004. Role of CREB in transcriptional regulation of CCAAT/enhancer-binding protein beta gene during adipogenesis. J. Biol. Chem. 279:4471-4478. [PubMed]

Articles from Molecular and Cellular Biology are provided here courtesy of American Society for Microbiology (ASM)
PubReader format: click here to try


Related citations in PubMed

See reviews...See all...

Cited by other articles in PMC

See all...


Recent Activity

Your browsing activity is empty.

Activity recording is turned off.

Turn recording back on

See more...