LS14: A Novel Human Adipocyte Cell Line that Produces Prolactin
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《内分泌学杂志》
Departments of Cell Biology (E.R.H., T.D.B., C.E.S.C., N.B.-J.) and Surgery (K.S.G., J.J.S.), University of Cincinnati College of Medicine, Cincinnati, Ohio 45257
Abstract
Adipose tissue is an integral component within the endocrine system. Adipocytes produce numerous bioactive substances, and their dysregulation has serious pathophysiological consequences. We previously reported that human adipose tissue from several depots produces significant amounts of prolactin (PRL). To study locally produced PRL, we sought an acceptable in vitro model. Consequently, we developed an adipocyte cell line derived from a metastatic liposarcoma. The cell line, designated LS14, has been in continuous culture for 2 yr. These cells exhibit many properties of primary preadipocytes, including the ability to undergo terminal differentiation, as judged by morphological alterations, lipid accumulation, and increase in glycerol-3-phosphate dehydrogenase. LS14 cells express many adipose-associated genes, such as adipocyte fatty acid-binding protein (aP2), hormone-sensitive lipase, lipoprotein lipase, preadipocyte factor 1, adiponectin, leptin, and IL-6. Similar to primary adipocytes, LS14 cells also produce and respond to PRL, thus making them an attractive model to study adipose PRL production and function. The expression of PRL was confirmed at the transcriptional level by RT-PCR, and PRL secretion was determined by the Nb2 bioassay. Addition of exogenous PRL to LS14 cells resulted in a dose-dependent inhibition of IL-6 release. In summary, we have established a novel human adipocyte cell line with many characteristics of primary adipocytes. The LS14 cells open up new avenues for research on human adipocyte biology and add to the repertoire of nonpituitary, PRL-producing cell lines.
Introduction
A DIPOSE TISSUE serves as an energy reservoir as well as a highly active endocrine tissue. It responds to and secretes numerous hormones, cytokines, and fat-specific molecules, collectively referred to as adipokines. Adipose-derived secretory molecules have extensive regulatory roles in metabolic homeostasis, with their major effects exerted on the brain, liver, gastrointestinal tract, and muscle (1, 2). Aberrant production of adipokines is associated with the pathophysiology of the metabolic syndrome, diabetes, and cardiovascular diseases (3). The incidence of excess adiposity or obesity has been steadily increasing in the past several decades, promoting active research on the endocrine functions of adipose tissue.
In vitro models for adipose tissue function in humans consist of primary adipocytes and cell lines. The utility of primary cells is limited by their inherent senescence as well as by patient to patient variations. Given the paucity of suitable human adipocyte cell lines (4, 5, 6, 7), most investigators have been using murine adipogenic cell lines such as 3T3-L1 and 3T3–442A. Although capable of high levels of differentiation, murine adipocytes do not accurately represent the full spectrum of hormonal and metabolic characteristics of human adipocytes (8).
Our laboratory, which has long been studying prolactin (PRL) production and function (9), became interested in adipose tissue because of a recent serendipitous observation. Upon investigating PRL production in human breast tissue, we discovered that adipose, rather than glandular, tissue was the major source of locally produced PRL (10). This novel observation led us to examine and then confirm PRL production by sc and visceral adipose depots (11). We recently identified mature adipocytes as the primary source of the hormone (12).
In humans, PRL is produced in multiple nonpituitary sites. Extrapituitary PRL production was first reported in 1978, with the detection of PRL expression in the human decidua (13), followed by the identification of a human lymphoid B cell line that produces PRL (14). Since then, multiple sites of PRL production, including breast, prostate, brain, myometrium, and, most recently, adipose, have been identified (15). Because the prolactin receptor (PRLR) is ubiquitously expressed, most tissues have the capacity to respond to PRL signaling. Given the multiple production sites of PRL and the widespread expression of its receptor, extrapituitary PRL most likely acts as an autocrine or paracrine signaling protein.
Given the difficulty in obtaining sufficient human adipose tissue and the unavoidable variability of clinical specimens, we sought to develop a suitable in vitro model. Because induced cell immortality, e.g. by viral transformation, often results in loss of typical characteristics, we looked for an alternative approach and elected to use a liposarcoma. Liposarcomas are the second most common soft tissue malignancies in humans (16). Although most liposarcomas have defective adipogenesis, this can often be overcome by exposure to exogenous ligands (17). In this study we report the successful generation of a liposarcoma-derived, novel, human adipocyte cell line, LS14. These cells have been in continuous culture since early 2004, exhibit a gene expression pattern similar to that in primary human preadipocytes, and are capable of terminal differentiation. LS14 cells express many adipocyte-specific genes as well as PRL and the PRLR.
Materials and Methods
Cloning of LS14 cells
Tumor tissue was obtained during abdominal surgery in November 2003 from a 52-yr-old man diagnosed with a recurrent liposarcoma. The patient gave informed consent in accordance with the rules and regulations of the institutional review board of University of Cincinnati. Liposarcoma cell cultures were established from a small tumor sample, processed within 30 min of excision. After mincing, fragments were suspended in Hanks’ balanced salt solution containing 1 mg/ml collagenase type I (Worthington Biochemical Corp., Lakewood, NJ) and digested for 12 h at 37 C with gentle agitation. After filtration through a 70-μm pore size nylon filter, the cells were sedimented by centrifugation at 500 x g for 10 min. Cells were seeded into tissue culture flasks (Nunc, Rochester, NY) containing various combinations of tissue culture medium and were incubated at 37 C with 95% relative humidity in 5% CO2. Optimal growth was observed in DMEM/Ham’s F-12 medium (1:1; Mediatech, Herndon, VA) without phenol red, supplemented with 10% fetal bovine serum (FBS; Invitrogen Life Technologies, Inc., Carlsbad, CA), 1% ITS+ additive (BD Clontech, Mountain View, CA), and 100 μg/ml of the antibiotic normocin (Invivogen, San Diego, CA). Cells were subcultured by a brief exposure to trypsin/EDTA (Invitrogen Life Technologies, Inc.). At passage 8, clonal cultures were generated by limiting dilution in 96-well plates. Several single-cell-derived cultures were obtained, of which clone 14, designated LS14 cells, showed the best growth and differentiation properties.
Harvesting of primary visceral human adipocytes
Fresh visceral (omentum) adipose tissue was obtained from morbidly obese patients undergoing gastric bypass surgery. The study was approved by the University of Cincinnati institutional review board, and informed consent was obtained from each patient. Tissue was minced and digested with collagenase I (Worthington Biochemical Corp.) at 200 U/g tissue in Hanks’ balanced salt solution containing 3% (wt/vol) fatty acid-free BSA (U.S. Biological Corp., Swampscott, MA) and 200 nM adenosine. Digestion was performed at 37 C with agitation for 30–40 min. After filtration through 70-μm pore size stainless steel mesh, mature adipocytes were collected from the floating cell layer after centrifugation at 200 x g for 1 min. Preadipocytes were sedimented by centrifugation at 800 x g for 10 min. Cell fractions were kept frozen at –70 C. For RNA extraction, either mature adipocytes or preadipocytes were pooled from several patients.
Cell growth
LS14 cells were plated into 96-well plates (Nunc) at a density of 500 cells/well. After 12–18 h, the plating medium was replaced with growth medium at eight wells per treatment. Cell growth was evaluated on various days by incubation with [3-(4,5-dimethylthiazol-2-yl)2,5-diphenyl tetrazolium bromide (MTT) at a final concentration of 0.5 mg/ml for 2 h. This was followed by aspiration of the medium and solubilization of the formazan product with dimethylsulfoxide. Absorbance was read at 570 nm using a 96-well plate reader (Bio-Tek, Winooski, VT).
Adipogenesis
LS14 cells were suspended in serum-free basal adipogenesis medium (BAM) consisting of phenol red-free DMEM/Ham’s F-12 (1:1) containing 33 μM biotin, 17 μM pantothenic acid, 1 μM human insulin (Sigma-Aldrich Corp., St. Louis, MO), 10 μg/ml apotransferrin (Sigma-Aldrich Corp.), 1 nM T3 (Sigma-Aldrich Corp.), 2 μM rosiglitazone (Kemprotec, Middlesbrough, UK), 200 μM ascorbate phosphate, 4 μM oleic acid/BSA, 4 μM linoleic acid/BSA, and 1 μM tripalmitate/BSA (U.S. Biochemical Corp., Cleveland, OH). Cells were plated at 35,000 cells/cm2 onto 6 well plates coated with type I rat tail collagen (50 μg/ml). After 24 h, adipogenesis was induced by incubation in BAM containing 250 μM isobutylmethylxanthine (IBMX; BIOMOL, Plymouth Meeting, PA). After 72 h of IBMX treatment, cells were incubated in BAM, which was replaced every fourth day throughout differentiation. The progress of differentiation was followed microscopically using Oil Red O staining. Briefly, cells were plated onto collagen-coated glass slides and differentiated as described above. Cells were fixed with 4% paraformaldehyde and stained with 0.1% Oil Red O in 60% triethyl phosphate for 30 min (18). After washing, cells were mounted in glycerol jelly and photographed using a SPOT-2 digital CCD camera mounted on Nikon Microphot microscope (Nikon Corp., Melville, NY).
Glycerol-3-phosphate dehydrogenase (GPDH) activity
GPDH (sn-glycerol-3-phosphate dehydrogenase; EC 1.1.1.8) activity was determined spectrophotometrically by measuring the disappearance of reduced nicotinamide adenine dinucleotide during GPDH-catalyzed reduction of dihydroxyacetone phosphate under zero-order conditions by the method of Wise and Green (19). Enzyme activity was expressed as units per milligram of protein, where 1 U of activity is defined as the oxidation of 1 nmol reduced nicotinamide adenine dinucleotide/min. Proteins was measured by a bicinchoninic acid method (20).
Conventional and real-time RT-PCR
Total RNA was isolated with Tri-Reagent (MRC, Cincinnati, OH), and its concentration and purity were determined spectrophotometrically. First-strand cDNA synthesis was performed using SuperScript II reverse transcriptase (Invitrogen Life Technologies, Inc.), oligo(deoxythymidine)12–18 (Invitrogen Life Technologies, Inc.), 5x first-strand synthesis buffer, 20 mM deoxy-NTPs (Roche, Indianapolis, IN), and RNasin placental ribonuclease inhibitor (Promega Corp., Madison, WI). Conventional PCR was performed using Immomix PCR premix (Bioline, Randolph, MA) and intron-spanning primers (Table 1). Standard cycle conditions were 96 C for 6 min for polymerase activation, followed by 28–34 cycles of 94, 57, and 72 C, each for 45 sec. Products were resolved on a 1.5% agarose gel containing ethidium bromide and photographed.
Quantitative real-time PCR was performed using Immolase heat-activated Taq DNA polymerase (Bioline) and appropriate primers (Table 1). SYBR Green I (Invitrogen Life Technologies, Inc.) was used for fluorometric product detection using a SmartCycler I (Cepheid, Sunnyvale, CA). Cycle parameters were 96 C for 6 min for polymerase activation, followed by 45 cycles of 94 C for 15 sec, 57 C for 15 sec, and 72 C for 30 sec, and finally an optical read stage at 83.5 C for 6 sec. Product purity was confirmed by DNA melting curve analysis and agarose gel electrophoresis. PCR efficiency was determined for each reaction using the LinRegPCR program (21) or, for a given primer set, by cDNA dose-response curve analysis. 2-Microglobulin (B2M) was used as a reference gene. Efficiencies for PRL and B2M were 79% and 97%, respectively. Fold changes in gene expression were calculated from the cycle threshold and efficiency measurements, using the method of Pfaffl et al. (22).
Nb2 bioassay for PRL
PRL release into conditioned medium (CM) was determined using the rat Nb2 lymphoma bioassay as previously described (10). Briefly, starved Nb2 cells were plated in 96-well plates (20,000 cells/well) and incubated with human PRL (National Institute of Diabetes and Digestive and Kidney Diseases, Bethesda, MD) in triplicate, and CM aliquots were incubated in duplicate. After 3 d, cell number was determined by the MTT method. The amount of PRL in the CM was calculated from the standard curve, with a lowest detectable level of 2 pg/well. To verify assay specificity, Nb2 cells were coincubated with CM aliquots and protein A purified anti-human PRL (anti-hPRL) antibodies (Upstate Biotechnology, Inc., Charlottesville, VA) or normal rabbit IgG.
IL-6 measurement
After treatment with hPRL, IL-6 levels in CM were determined by solid-phase sandwich ELISA, using optimized and validated monoclonal antibody pairs to hIL-6 (Cytosets, BioSource International, Camarillo, CA). Bound biotinylated antibodies were detected using avidin/horseradish peroxidase and a fluorometric horseradish peroxidase substrate (QuantaBlue, Pierce Chemical Co., Rockford, IL) and were measured at 325 nm excitation and 420 nm emission on a Gemini fluorescence plate reader (Molecular Devices, Sunnyvale, CA). The detection limit was 20 pg/ml.
Data analysis
When appropriate, values are expressed as the mean ± SEM. The effect of PRL on IL-6 release was analyzed by Student’s t test using the program Kaleidagraph 4.0 (Synergy Software, Reading, PA). Significance was set at P < 0.05.
Results
Optimization of growth and differentiation of LS14 cells
After cloning, we obtained eight cultures. Clone 14, designated LS14, exhibited the best growth and differentiation characteristics and was used to optimize the culture conditions for both growth and differentiation. Although LS14 cells grew faster in medium containing the Clonetics (Cambrex, Walkersville, MD) Smooth Muscle Growth Medium-3 supplements (Fig. 1), they had a lower rate of differentiation (data not shown). Cells grown in 10% FBS alone showed suppressed growth parameters. A medium containing 5% FBS, 5% FetalClone III (HyClone, Logan, UT), 1% ITS+ additive, and bovine pituitary extract (15 μg/ml protein; Invitrogen Life Technologies, Inc.) was selected for the remainder of the studies because it provided the best compromise between cell growth (doubling time of 96 h) and differentiation. Notably, when cultured in BAM, the cells showed lack of growth (Fig. 1).
Induction of differentiation was accomplished using serum-free medium, developed by Van Harmelen et al. (23) and modified by the addition of a retinoid X receptor ligand, fatty acids, and a triglyceride. Under these conditions, a differentiation rate of 50–70% as early as d 10 of differentiation was achieved, as judged by lipid accumulation (Fig. 2). Differentiation was also confirmed by a marked increase in GPDH activity on d 10 of adipogenesis.
Comparison of gene expression in LS14 cells and primary adipocytes before and after differentiation
Conventional RT-PCR was used to compare the expression of adipose-specific genes (Fig. 3) and selected cytokines/hormones and receptors (Fig. 4) in LS14 and primary visceral adipocytes. The adipose-specific genes, aP2, glucose transporter 4, hormone-sensitive lipase, and lipoprotein lipase, were similarly induced during differentiation of LS14 and primary adipocytes. Peroxisome proliferator-activating receptor was robustly expressed in both cell types before and after differentiation, whereas the expression of uncoupling protein 1 was more apparent in primary than in LS14 cells, with a low detection of preadipocyte factor 1 in both cell types (Fig. 3). The expression of adiponectin, leptin, and angiotensinogen was similar in differentiated LS14 and primary adipocytes (Fig. 4), whereas IL-6 and TNF were more highly expressed in primary cells, especially after differentiation. There was no evidence for 1- and 3-adrenergic receptors (AR) expression in differentiated LS14 cells, with the expression of 2AR primarily observed in nondifferentiated cells (Fig. 4). Notably, the expression of both PRL and PRLR was evident in both cell types before and after differentiation.
Changes in PRL and PRLR expression throughout LS14 differentiation
The expression of both PRL and PRLR during differentiation of LS14 cells was quantified using real-time RT-PCR. As shown in Fig. 5, upper right panel, PRL expression increased 8- and 22-fold on d 1 and 3 of differentiation, followed by a decline on d 7 and 10. In contrast, PRLR expression was reduced 7- and 17-fold on d 1 and 3 of differentiation, returning to predifferentiation levels on d 7 and 10 (Fig. 5, lower right panel).
PRL release during LS14 differentiation
To verify that not only PRL expression but also PRL release was altered during LS14 differentiation, we used the sensitive Nb2 bioassay. Because dexamethasone, included in BAM, is apoptotic for Nb2 cells (24, 25), the protocol was modified so as to remove the glucocorticoids. For that, LS14 cells were incubated in adipogenic medium as described, then washed free of the medium on the designated days of differentiation, followed by incubation for 24 h in DMEM/Ham’s F-12 containing 2% charcoal-stripped serum. CM from this incubation was analyzed for PRL by the Nb2 bioassay. As evident in Fig. 6, left panel, PRL release was unchanged on d 1, increased 2- and 3-fold on d 3 and 7, and was markedly reduced thereafter. To validate assay specificity for PRL, Nb2 cells were coincubated with CM aliquots from LS14 on d 7 of differentiation, either alone or together with anti-hPRL IgG or rabbit IgG. As shown in Fig. 6, right panel, the Nb2 cell proliferative response to CM was completely abolished by hPRL antibodies, confirming that all of the Nb2 proliferative response to CM was due to PRL.
Changes in IL-6 expression during differentiation and effects of exogenous PRL on IL-6 release
As determined by real-time PCR, IL-6 expression decreased considerably throughout LS-14 differentiation, reaching 100- and 200-fold decreases on d 3 and 10, respectively (Fig. 7, left panel). Addition of exogenous PRL to nondifferentiated LS14 cells resulted in a dose-dependent inhibition of IL-6 release, as determined by ELISA (Fig. 7, right panel). IL-6 levels were reduced by 40% and 60% after incubation with 5 and 125 ng/ml PRL, respectively.
Discussion
This report describes the generation and characterization of a new, spontaneously immortalized, human adipocyte cell line that produces and responds to PRL and is capable of terminal differentiation. This cell line possesses many of the morphological and biochemical characteristics of primary human adipocytes and thus fills a major shortage in suitable cellular models of human adipocyte biology and adipogenesis. Although in this report we placed an emphasis on PRL production and functions, LS14 cells can be used for investigating many aspects of human adipocyte homeostasis, including responsiveness to hormones, cytokines, and metabolites; adipogenesis; and the regulation and functions of cytokines/adipokines.
Human preadipocytes immortalized by transforming viruses (4, 26) or liposarcoma-derived tumor cells (5, 6, 7) provide an abundant supply of homogeneous cells and complement studies using primary cells. The LS14 clone was derived from a low grade, recurrent, metastatic liposarcoma. Liposarcomas are a soft tissue malignancy with an incidence of 15% of all sarcomas (27). Localized liposarcomas are treated with surgery, whereas metastatic liposarcomas are normally unresponsive to chemotherapy and result in poor prognosis (28). The origin of the LS14 clone is unclear, but may be a dedifferentiated cell or a stem cell. This clone has been maintained in culture for over 100 generations. Morphologically, the cells are much larger than primary preadipocytes, with many cells displaying multiple nuclei. They have a longer doubling time (96 h) than that of freshly isolated primary human preadipocytes and require a more complex culture medium composition to support adequate cell growth. The growth rate of LS14 cells can be extensively altered by the inclusion of different growth factors and hormones. Although the current culture medium composition may not be optimal for rapid cell growth, it reflects a compromise between a practical doubling time and cell competency for undergoing terminal differentiation.
Much of the current knowledge of adipogenesis is based on studies with murine preadipocyte cell lines. The most widely used are 3T3-L1 and 3T3-F422A cells, which were derived from disaggregated Swiss 3T3 mouse embryos (29, 30). At low density, they can be passaged indefinitely, whereas upon reaching confluence, they can be induced to differentiate by a well-defined adipogenic mixture. Within 24 h, they undergo postconfluent mitosis, followed by growth arrest. They start expressing markers of differentiation by d 3–4 and complete the conversion process by d 7–8 (31). Unlike murine preadipocytes, adipogenesis in LS14 cells does not require serum, but, in fact, is inhibited by serum. During differentiation, these cells also show a weaker adhesion to tissue culture surfaces than primary human and murine cells, and this can be ameliorated by coating the plates with rat tail collagen. The use of fibronectin, Matrigel, poly-D-lysine, and protamine was not as effective as collagen (data not shown).
Under optimal differentiation conditions, more than 60% of LS14 cells undergo terminal differentiation, a percentage similar to that reported for visceral human preadipocytes (32). Terminal differentiation was judged by several morphological, biochemical, and molecular criteria. During differentiation, LS14 cells initially become more compact. By d 7, lipid accumulation becomes apparent, increasing significantly by d 10. Increased lipogenic activity in the differentiated cells is supported by the marked rise in GPDH activity, a critical enzyme for glucose metabolism (33). The higher expression of adipocyte-specific markers, such as adiponectin, leptin, aP2, and lipoprotein lipase in differentiated LS14 cells also confirms their functional maturation. The pattern of expression of most genes, examined by conventional RT-PCR, is similar in LS14 cells and visceral adipocytes before and after differentiation. Notable exceptions are the reduced expression of TNF and IL-6 in differentiated LS14 cells. The expression of 3AR, which is unique to adipose tissue (34), is undetectable in LS14 cells, which do express 1- and 2AR in the undifferentiated state. ARs play an important role in the regulation of lipolysis in adipose tissue (35). Differentiated LS14 cells also show a robust expression of leptin, unlike the murine counterparts, where leptin expression is down-regulated under high levels of insulin and glucocorticoids (36).
We previously reported that adipose tissue was the major source of locally produced PRL within the breast (10). We also found that adipose PRL expression is driven by a superdistal promoter that has been well characterized in several human tissues and cell lines (37, 38, 39). The present results clearly demonstrate the expression of both PRL and its receptor in primary adipocytes as well as LS14 cells. PRL production, at both expression and protein levels, increases markedly during early LS14 differentiation. We speculate that this may be due to the presence of IBMX, a phosphodiesterase inhibitor and cAMP activator, during the first 3 d of differentiation. Induction of the superdistal PRL promoter by cAMP activators has been reported by several investigators (40, 41). Ongoing research examines the effects of catecholamines and other cAMP-activating ligands on the control of PRL expression. The apparent inverse relationship between the expression of PRL and its receptor during LS14 differentiation raises the possibility that PRLR is down-regulated by its ligand.
The spectrum of PRL functions in adipose tissue remains to be determined. Because LS14 cells make their own PRL, we opted to examine the effect of PRL removal on cell growth or differentiation. Theoretically, this can be accomplished in three ways: 1) through sequestration of endogenous PRL, 2), by blocking the PRLR, and 3) by down-regulating the PRL gene. To date, our efforts to sequester PRL, using either anti-hPRL antibodies or a soluble extracellular domain of the PRL receptor protein (42) at concentrations that completely block the ability of PRL to stimulate Nb2 growth, have been unsuccessful. We speculate that the failure to sequester PRL is due to its binding to heparin-containing proteoglycans in the extracellular matrix (43). When bound to heparin, both the antibody and the extracellular domain may have limited access to the hormone. Presently, we are developing a small interference RNA approach to down-regulate the PRL gene.
To date, we have examined the effect of exogenous PRL on LS14 cells. Previous reports indicated interactions between PRL and IL-6 in mouse mammary epithelia cells (44) and suppression of IL-6 expression by PRL in rat decidual cells (45). IL-6 is a pleiotropic cytokine, with 30–40% of its circulating levels originating from adipose tissue (46). In addition to playing a role in inflammation (47), IL-6 is involved in tissue remodeling (45). Production of IL-6 can be induced by TNF, which, in turn, is elevated by infection or tissue damage; thus, IL-6 is an acute phase inflammatory protein (48). Within human adipose tissue depots, spontaneous IL-6 release is 3-fold higher in omental than sc explants, and its release is suppressed by glucocorticoids (49). Addition of exogenous PRL to nondifferentiated LS14 cells resulted in a significant, dose-dependent inhibition of IL-6 release. A similar inhibitory effect of PRL on IL-6 was observed using visceral preadipocytes (data not shown). We also observed a progressive, marked suppression of IL-6 gene expression throughout LS14 differentiation, which could be functionally related to the early elevation of endogenous PRL. We have not been able to confirm the suppression of IL-6 during differentiation at the secretion level due to the fact that IL-6 levels are near the limit of detection for the ELISA used. Preliminary studies have also shown the suppression of leptin release by PRL (data not shown).
In summary, we have established a human adipocyte cell line with many characteristics of primary adipocytes. Unlike most transformed cells that are resistant to differentiation, these cells can undergo considerable morphological and functional differentiation under the appropriate culture conditions. LS14 cells open up new avenues for research on human adipocyte biology and add to the repertoire of nonpituitary, PRL-producing cell lines.
Footnotes
This work was supported by National Institutes of Health Grants ES-0955, ES-012212, and CA-096613; and Training Grant DK59803 (to E.R.H.).
Preliminary results of this investigation were presented at the Obesity and Diabetes Joint Keystone Conference, Banff, Alberta, Canada, March 2004.
First Published Online September 29, 2005
Abbreviations: aP2, Adipose fatty acid-binding protein; AR, -Adrenergic receptor; BAM, basal adipogenesis medium; B2M, 2-microglobulin; CM, conditioned medium; FBS, fetal bovine serum; GPDH, glycerol-3-phosphate dehydrogenase; h, human; IBMX, isobutylmethylxanthine; MTT, [3-(4,5-dimethylthiazol-2-yl)2,5-diphenyl tetrazolium bromide; PRL, prolactin; PRLR, prolactin receptor.
Accepted for publication September 22, 2005.
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Abstract
Adipose tissue is an integral component within the endocrine system. Adipocytes produce numerous bioactive substances, and their dysregulation has serious pathophysiological consequences. We previously reported that human adipose tissue from several depots produces significant amounts of prolactin (PRL). To study locally produced PRL, we sought an acceptable in vitro model. Consequently, we developed an adipocyte cell line derived from a metastatic liposarcoma. The cell line, designated LS14, has been in continuous culture for 2 yr. These cells exhibit many properties of primary preadipocytes, including the ability to undergo terminal differentiation, as judged by morphological alterations, lipid accumulation, and increase in glycerol-3-phosphate dehydrogenase. LS14 cells express many adipose-associated genes, such as adipocyte fatty acid-binding protein (aP2), hormone-sensitive lipase, lipoprotein lipase, preadipocyte factor 1, adiponectin, leptin, and IL-6. Similar to primary adipocytes, LS14 cells also produce and respond to PRL, thus making them an attractive model to study adipose PRL production and function. The expression of PRL was confirmed at the transcriptional level by RT-PCR, and PRL secretion was determined by the Nb2 bioassay. Addition of exogenous PRL to LS14 cells resulted in a dose-dependent inhibition of IL-6 release. In summary, we have established a novel human adipocyte cell line with many characteristics of primary adipocytes. The LS14 cells open up new avenues for research on human adipocyte biology and add to the repertoire of nonpituitary, PRL-producing cell lines.
Introduction
A DIPOSE TISSUE serves as an energy reservoir as well as a highly active endocrine tissue. It responds to and secretes numerous hormones, cytokines, and fat-specific molecules, collectively referred to as adipokines. Adipose-derived secretory molecules have extensive regulatory roles in metabolic homeostasis, with their major effects exerted on the brain, liver, gastrointestinal tract, and muscle (1, 2). Aberrant production of adipokines is associated with the pathophysiology of the metabolic syndrome, diabetes, and cardiovascular diseases (3). The incidence of excess adiposity or obesity has been steadily increasing in the past several decades, promoting active research on the endocrine functions of adipose tissue.
In vitro models for adipose tissue function in humans consist of primary adipocytes and cell lines. The utility of primary cells is limited by their inherent senescence as well as by patient to patient variations. Given the paucity of suitable human adipocyte cell lines (4, 5, 6, 7), most investigators have been using murine adipogenic cell lines such as 3T3-L1 and 3T3–442A. Although capable of high levels of differentiation, murine adipocytes do not accurately represent the full spectrum of hormonal and metabolic characteristics of human adipocytes (8).
Our laboratory, which has long been studying prolactin (PRL) production and function (9), became interested in adipose tissue because of a recent serendipitous observation. Upon investigating PRL production in human breast tissue, we discovered that adipose, rather than glandular, tissue was the major source of locally produced PRL (10). This novel observation led us to examine and then confirm PRL production by sc and visceral adipose depots (11). We recently identified mature adipocytes as the primary source of the hormone (12).
In humans, PRL is produced in multiple nonpituitary sites. Extrapituitary PRL production was first reported in 1978, with the detection of PRL expression in the human decidua (13), followed by the identification of a human lymphoid B cell line that produces PRL (14). Since then, multiple sites of PRL production, including breast, prostate, brain, myometrium, and, most recently, adipose, have been identified (15). Because the prolactin receptor (PRLR) is ubiquitously expressed, most tissues have the capacity to respond to PRL signaling. Given the multiple production sites of PRL and the widespread expression of its receptor, extrapituitary PRL most likely acts as an autocrine or paracrine signaling protein.
Given the difficulty in obtaining sufficient human adipose tissue and the unavoidable variability of clinical specimens, we sought to develop a suitable in vitro model. Because induced cell immortality, e.g. by viral transformation, often results in loss of typical characteristics, we looked for an alternative approach and elected to use a liposarcoma. Liposarcomas are the second most common soft tissue malignancies in humans (16). Although most liposarcomas have defective adipogenesis, this can often be overcome by exposure to exogenous ligands (17). In this study we report the successful generation of a liposarcoma-derived, novel, human adipocyte cell line, LS14. These cells have been in continuous culture since early 2004, exhibit a gene expression pattern similar to that in primary human preadipocytes, and are capable of terminal differentiation. LS14 cells express many adipocyte-specific genes as well as PRL and the PRLR.
Materials and Methods
Cloning of LS14 cells
Tumor tissue was obtained during abdominal surgery in November 2003 from a 52-yr-old man diagnosed with a recurrent liposarcoma. The patient gave informed consent in accordance with the rules and regulations of the institutional review board of University of Cincinnati. Liposarcoma cell cultures were established from a small tumor sample, processed within 30 min of excision. After mincing, fragments were suspended in Hanks’ balanced salt solution containing 1 mg/ml collagenase type I (Worthington Biochemical Corp., Lakewood, NJ) and digested for 12 h at 37 C with gentle agitation. After filtration through a 70-μm pore size nylon filter, the cells were sedimented by centrifugation at 500 x g for 10 min. Cells were seeded into tissue culture flasks (Nunc, Rochester, NY) containing various combinations of tissue culture medium and were incubated at 37 C with 95% relative humidity in 5% CO2. Optimal growth was observed in DMEM/Ham’s F-12 medium (1:1; Mediatech, Herndon, VA) without phenol red, supplemented with 10% fetal bovine serum (FBS; Invitrogen Life Technologies, Inc., Carlsbad, CA), 1% ITS+ additive (BD Clontech, Mountain View, CA), and 100 μg/ml of the antibiotic normocin (Invivogen, San Diego, CA). Cells were subcultured by a brief exposure to trypsin/EDTA (Invitrogen Life Technologies, Inc.). At passage 8, clonal cultures were generated by limiting dilution in 96-well plates. Several single-cell-derived cultures were obtained, of which clone 14, designated LS14 cells, showed the best growth and differentiation properties.
Harvesting of primary visceral human adipocytes
Fresh visceral (omentum) adipose tissue was obtained from morbidly obese patients undergoing gastric bypass surgery. The study was approved by the University of Cincinnati institutional review board, and informed consent was obtained from each patient. Tissue was minced and digested with collagenase I (Worthington Biochemical Corp.) at 200 U/g tissue in Hanks’ balanced salt solution containing 3% (wt/vol) fatty acid-free BSA (U.S. Biological Corp., Swampscott, MA) and 200 nM adenosine. Digestion was performed at 37 C with agitation for 30–40 min. After filtration through 70-μm pore size stainless steel mesh, mature adipocytes were collected from the floating cell layer after centrifugation at 200 x g for 1 min. Preadipocytes were sedimented by centrifugation at 800 x g for 10 min. Cell fractions were kept frozen at –70 C. For RNA extraction, either mature adipocytes or preadipocytes were pooled from several patients.
Cell growth
LS14 cells were plated into 96-well plates (Nunc) at a density of 500 cells/well. After 12–18 h, the plating medium was replaced with growth medium at eight wells per treatment. Cell growth was evaluated on various days by incubation with [3-(4,5-dimethylthiazol-2-yl)2,5-diphenyl tetrazolium bromide (MTT) at a final concentration of 0.5 mg/ml for 2 h. This was followed by aspiration of the medium and solubilization of the formazan product with dimethylsulfoxide. Absorbance was read at 570 nm using a 96-well plate reader (Bio-Tek, Winooski, VT).
Adipogenesis
LS14 cells were suspended in serum-free basal adipogenesis medium (BAM) consisting of phenol red-free DMEM/Ham’s F-12 (1:1) containing 33 μM biotin, 17 μM pantothenic acid, 1 μM human insulin (Sigma-Aldrich Corp., St. Louis, MO), 10 μg/ml apotransferrin (Sigma-Aldrich Corp.), 1 nM T3 (Sigma-Aldrich Corp.), 2 μM rosiglitazone (Kemprotec, Middlesbrough, UK), 200 μM ascorbate phosphate, 4 μM oleic acid/BSA, 4 μM linoleic acid/BSA, and 1 μM tripalmitate/BSA (U.S. Biochemical Corp., Cleveland, OH). Cells were plated at 35,000 cells/cm2 onto 6 well plates coated with type I rat tail collagen (50 μg/ml). After 24 h, adipogenesis was induced by incubation in BAM containing 250 μM isobutylmethylxanthine (IBMX; BIOMOL, Plymouth Meeting, PA). After 72 h of IBMX treatment, cells were incubated in BAM, which was replaced every fourth day throughout differentiation. The progress of differentiation was followed microscopically using Oil Red O staining. Briefly, cells were plated onto collagen-coated glass slides and differentiated as described above. Cells were fixed with 4% paraformaldehyde and stained with 0.1% Oil Red O in 60% triethyl phosphate for 30 min (18). After washing, cells were mounted in glycerol jelly and photographed using a SPOT-2 digital CCD camera mounted on Nikon Microphot microscope (Nikon Corp., Melville, NY).
Glycerol-3-phosphate dehydrogenase (GPDH) activity
GPDH (sn-glycerol-3-phosphate dehydrogenase; EC 1.1.1.8) activity was determined spectrophotometrically by measuring the disappearance of reduced nicotinamide adenine dinucleotide during GPDH-catalyzed reduction of dihydroxyacetone phosphate under zero-order conditions by the method of Wise and Green (19). Enzyme activity was expressed as units per milligram of protein, where 1 U of activity is defined as the oxidation of 1 nmol reduced nicotinamide adenine dinucleotide/min. Proteins was measured by a bicinchoninic acid method (20).
Conventional and real-time RT-PCR
Total RNA was isolated with Tri-Reagent (MRC, Cincinnati, OH), and its concentration and purity were determined spectrophotometrically. First-strand cDNA synthesis was performed using SuperScript II reverse transcriptase (Invitrogen Life Technologies, Inc.), oligo(deoxythymidine)12–18 (Invitrogen Life Technologies, Inc.), 5x first-strand synthesis buffer, 20 mM deoxy-NTPs (Roche, Indianapolis, IN), and RNasin placental ribonuclease inhibitor (Promega Corp., Madison, WI). Conventional PCR was performed using Immomix PCR premix (Bioline, Randolph, MA) and intron-spanning primers (Table 1). Standard cycle conditions were 96 C for 6 min for polymerase activation, followed by 28–34 cycles of 94, 57, and 72 C, each for 45 sec. Products were resolved on a 1.5% agarose gel containing ethidium bromide and photographed.
Quantitative real-time PCR was performed using Immolase heat-activated Taq DNA polymerase (Bioline) and appropriate primers (Table 1). SYBR Green I (Invitrogen Life Technologies, Inc.) was used for fluorometric product detection using a SmartCycler I (Cepheid, Sunnyvale, CA). Cycle parameters were 96 C for 6 min for polymerase activation, followed by 45 cycles of 94 C for 15 sec, 57 C for 15 sec, and 72 C for 30 sec, and finally an optical read stage at 83.5 C for 6 sec. Product purity was confirmed by DNA melting curve analysis and agarose gel electrophoresis. PCR efficiency was determined for each reaction using the LinRegPCR program (21) or, for a given primer set, by cDNA dose-response curve analysis. 2-Microglobulin (B2M) was used as a reference gene. Efficiencies for PRL and B2M were 79% and 97%, respectively. Fold changes in gene expression were calculated from the cycle threshold and efficiency measurements, using the method of Pfaffl et al. (22).
Nb2 bioassay for PRL
PRL release into conditioned medium (CM) was determined using the rat Nb2 lymphoma bioassay as previously described (10). Briefly, starved Nb2 cells were plated in 96-well plates (20,000 cells/well) and incubated with human PRL (National Institute of Diabetes and Digestive and Kidney Diseases, Bethesda, MD) in triplicate, and CM aliquots were incubated in duplicate. After 3 d, cell number was determined by the MTT method. The amount of PRL in the CM was calculated from the standard curve, with a lowest detectable level of 2 pg/well. To verify assay specificity, Nb2 cells were coincubated with CM aliquots and protein A purified anti-human PRL (anti-hPRL) antibodies (Upstate Biotechnology, Inc., Charlottesville, VA) or normal rabbit IgG.
IL-6 measurement
After treatment with hPRL, IL-6 levels in CM were determined by solid-phase sandwich ELISA, using optimized and validated monoclonal antibody pairs to hIL-6 (Cytosets, BioSource International, Camarillo, CA). Bound biotinylated antibodies were detected using avidin/horseradish peroxidase and a fluorometric horseradish peroxidase substrate (QuantaBlue, Pierce Chemical Co., Rockford, IL) and were measured at 325 nm excitation and 420 nm emission on a Gemini fluorescence plate reader (Molecular Devices, Sunnyvale, CA). The detection limit was 20 pg/ml.
Data analysis
When appropriate, values are expressed as the mean ± SEM. The effect of PRL on IL-6 release was analyzed by Student’s t test using the program Kaleidagraph 4.0 (Synergy Software, Reading, PA). Significance was set at P < 0.05.
Results
Optimization of growth and differentiation of LS14 cells
After cloning, we obtained eight cultures. Clone 14, designated LS14, exhibited the best growth and differentiation characteristics and was used to optimize the culture conditions for both growth and differentiation. Although LS14 cells grew faster in medium containing the Clonetics (Cambrex, Walkersville, MD) Smooth Muscle Growth Medium-3 supplements (Fig. 1), they had a lower rate of differentiation (data not shown). Cells grown in 10% FBS alone showed suppressed growth parameters. A medium containing 5% FBS, 5% FetalClone III (HyClone, Logan, UT), 1% ITS+ additive, and bovine pituitary extract (15 μg/ml protein; Invitrogen Life Technologies, Inc.) was selected for the remainder of the studies because it provided the best compromise between cell growth (doubling time of 96 h) and differentiation. Notably, when cultured in BAM, the cells showed lack of growth (Fig. 1).
Induction of differentiation was accomplished using serum-free medium, developed by Van Harmelen et al. (23) and modified by the addition of a retinoid X receptor ligand, fatty acids, and a triglyceride. Under these conditions, a differentiation rate of 50–70% as early as d 10 of differentiation was achieved, as judged by lipid accumulation (Fig. 2). Differentiation was also confirmed by a marked increase in GPDH activity on d 10 of adipogenesis.
Comparison of gene expression in LS14 cells and primary adipocytes before and after differentiation
Conventional RT-PCR was used to compare the expression of adipose-specific genes (Fig. 3) and selected cytokines/hormones and receptors (Fig. 4) in LS14 and primary visceral adipocytes. The adipose-specific genes, aP2, glucose transporter 4, hormone-sensitive lipase, and lipoprotein lipase, were similarly induced during differentiation of LS14 and primary adipocytes. Peroxisome proliferator-activating receptor was robustly expressed in both cell types before and after differentiation, whereas the expression of uncoupling protein 1 was more apparent in primary than in LS14 cells, with a low detection of preadipocyte factor 1 in both cell types (Fig. 3). The expression of adiponectin, leptin, and angiotensinogen was similar in differentiated LS14 and primary adipocytes (Fig. 4), whereas IL-6 and TNF were more highly expressed in primary cells, especially after differentiation. There was no evidence for 1- and 3-adrenergic receptors (AR) expression in differentiated LS14 cells, with the expression of 2AR primarily observed in nondifferentiated cells (Fig. 4). Notably, the expression of both PRL and PRLR was evident in both cell types before and after differentiation.
Changes in PRL and PRLR expression throughout LS14 differentiation
The expression of both PRL and PRLR during differentiation of LS14 cells was quantified using real-time RT-PCR. As shown in Fig. 5, upper right panel, PRL expression increased 8- and 22-fold on d 1 and 3 of differentiation, followed by a decline on d 7 and 10. In contrast, PRLR expression was reduced 7- and 17-fold on d 1 and 3 of differentiation, returning to predifferentiation levels on d 7 and 10 (Fig. 5, lower right panel).
PRL release during LS14 differentiation
To verify that not only PRL expression but also PRL release was altered during LS14 differentiation, we used the sensitive Nb2 bioassay. Because dexamethasone, included in BAM, is apoptotic for Nb2 cells (24, 25), the protocol was modified so as to remove the glucocorticoids. For that, LS14 cells were incubated in adipogenic medium as described, then washed free of the medium on the designated days of differentiation, followed by incubation for 24 h in DMEM/Ham’s F-12 containing 2% charcoal-stripped serum. CM from this incubation was analyzed for PRL by the Nb2 bioassay. As evident in Fig. 6, left panel, PRL release was unchanged on d 1, increased 2- and 3-fold on d 3 and 7, and was markedly reduced thereafter. To validate assay specificity for PRL, Nb2 cells were coincubated with CM aliquots from LS14 on d 7 of differentiation, either alone or together with anti-hPRL IgG or rabbit IgG. As shown in Fig. 6, right panel, the Nb2 cell proliferative response to CM was completely abolished by hPRL antibodies, confirming that all of the Nb2 proliferative response to CM was due to PRL.
Changes in IL-6 expression during differentiation and effects of exogenous PRL on IL-6 release
As determined by real-time PCR, IL-6 expression decreased considerably throughout LS-14 differentiation, reaching 100- and 200-fold decreases on d 3 and 10, respectively (Fig. 7, left panel). Addition of exogenous PRL to nondifferentiated LS14 cells resulted in a dose-dependent inhibition of IL-6 release, as determined by ELISA (Fig. 7, right panel). IL-6 levels were reduced by 40% and 60% after incubation with 5 and 125 ng/ml PRL, respectively.
Discussion
This report describes the generation and characterization of a new, spontaneously immortalized, human adipocyte cell line that produces and responds to PRL and is capable of terminal differentiation. This cell line possesses many of the morphological and biochemical characteristics of primary human adipocytes and thus fills a major shortage in suitable cellular models of human adipocyte biology and adipogenesis. Although in this report we placed an emphasis on PRL production and functions, LS14 cells can be used for investigating many aspects of human adipocyte homeostasis, including responsiveness to hormones, cytokines, and metabolites; adipogenesis; and the regulation and functions of cytokines/adipokines.
Human preadipocytes immortalized by transforming viruses (4, 26) or liposarcoma-derived tumor cells (5, 6, 7) provide an abundant supply of homogeneous cells and complement studies using primary cells. The LS14 clone was derived from a low grade, recurrent, metastatic liposarcoma. Liposarcomas are a soft tissue malignancy with an incidence of 15% of all sarcomas (27). Localized liposarcomas are treated with surgery, whereas metastatic liposarcomas are normally unresponsive to chemotherapy and result in poor prognosis (28). The origin of the LS14 clone is unclear, but may be a dedifferentiated cell or a stem cell. This clone has been maintained in culture for over 100 generations. Morphologically, the cells are much larger than primary preadipocytes, with many cells displaying multiple nuclei. They have a longer doubling time (96 h) than that of freshly isolated primary human preadipocytes and require a more complex culture medium composition to support adequate cell growth. The growth rate of LS14 cells can be extensively altered by the inclusion of different growth factors and hormones. Although the current culture medium composition may not be optimal for rapid cell growth, it reflects a compromise between a practical doubling time and cell competency for undergoing terminal differentiation.
Much of the current knowledge of adipogenesis is based on studies with murine preadipocyte cell lines. The most widely used are 3T3-L1 and 3T3-F422A cells, which were derived from disaggregated Swiss 3T3 mouse embryos (29, 30). At low density, they can be passaged indefinitely, whereas upon reaching confluence, they can be induced to differentiate by a well-defined adipogenic mixture. Within 24 h, they undergo postconfluent mitosis, followed by growth arrest. They start expressing markers of differentiation by d 3–4 and complete the conversion process by d 7–8 (31). Unlike murine preadipocytes, adipogenesis in LS14 cells does not require serum, but, in fact, is inhibited by serum. During differentiation, these cells also show a weaker adhesion to tissue culture surfaces than primary human and murine cells, and this can be ameliorated by coating the plates with rat tail collagen. The use of fibronectin, Matrigel, poly-D-lysine, and protamine was not as effective as collagen (data not shown).
Under optimal differentiation conditions, more than 60% of LS14 cells undergo terminal differentiation, a percentage similar to that reported for visceral human preadipocytes (32). Terminal differentiation was judged by several morphological, biochemical, and molecular criteria. During differentiation, LS14 cells initially become more compact. By d 7, lipid accumulation becomes apparent, increasing significantly by d 10. Increased lipogenic activity in the differentiated cells is supported by the marked rise in GPDH activity, a critical enzyme for glucose metabolism (33). The higher expression of adipocyte-specific markers, such as adiponectin, leptin, aP2, and lipoprotein lipase in differentiated LS14 cells also confirms their functional maturation. The pattern of expression of most genes, examined by conventional RT-PCR, is similar in LS14 cells and visceral adipocytes before and after differentiation. Notable exceptions are the reduced expression of TNF and IL-6 in differentiated LS14 cells. The expression of 3AR, which is unique to adipose tissue (34), is undetectable in LS14 cells, which do express 1- and 2AR in the undifferentiated state. ARs play an important role in the regulation of lipolysis in adipose tissue (35). Differentiated LS14 cells also show a robust expression of leptin, unlike the murine counterparts, where leptin expression is down-regulated under high levels of insulin and glucocorticoids (36).
We previously reported that adipose tissue was the major source of locally produced PRL within the breast (10). We also found that adipose PRL expression is driven by a superdistal promoter that has been well characterized in several human tissues and cell lines (37, 38, 39). The present results clearly demonstrate the expression of both PRL and its receptor in primary adipocytes as well as LS14 cells. PRL production, at both expression and protein levels, increases markedly during early LS14 differentiation. We speculate that this may be due to the presence of IBMX, a phosphodiesterase inhibitor and cAMP activator, during the first 3 d of differentiation. Induction of the superdistal PRL promoter by cAMP activators has been reported by several investigators (40, 41). Ongoing research examines the effects of catecholamines and other cAMP-activating ligands on the control of PRL expression. The apparent inverse relationship between the expression of PRL and its receptor during LS14 differentiation raises the possibility that PRLR is down-regulated by its ligand.
The spectrum of PRL functions in adipose tissue remains to be determined. Because LS14 cells make their own PRL, we opted to examine the effect of PRL removal on cell growth or differentiation. Theoretically, this can be accomplished in three ways: 1) through sequestration of endogenous PRL, 2), by blocking the PRLR, and 3) by down-regulating the PRL gene. To date, our efforts to sequester PRL, using either anti-hPRL antibodies or a soluble extracellular domain of the PRL receptor protein (42) at concentrations that completely block the ability of PRL to stimulate Nb2 growth, have been unsuccessful. We speculate that the failure to sequester PRL is due to its binding to heparin-containing proteoglycans in the extracellular matrix (43). When bound to heparin, both the antibody and the extracellular domain may have limited access to the hormone. Presently, we are developing a small interference RNA approach to down-regulate the PRL gene.
To date, we have examined the effect of exogenous PRL on LS14 cells. Previous reports indicated interactions between PRL and IL-6 in mouse mammary epithelia cells (44) and suppression of IL-6 expression by PRL in rat decidual cells (45). IL-6 is a pleiotropic cytokine, with 30–40% of its circulating levels originating from adipose tissue (46). In addition to playing a role in inflammation (47), IL-6 is involved in tissue remodeling (45). Production of IL-6 can be induced by TNF, which, in turn, is elevated by infection or tissue damage; thus, IL-6 is an acute phase inflammatory protein (48). Within human adipose tissue depots, spontaneous IL-6 release is 3-fold higher in omental than sc explants, and its release is suppressed by glucocorticoids (49). Addition of exogenous PRL to nondifferentiated LS14 cells resulted in a significant, dose-dependent inhibition of IL-6 release. A similar inhibitory effect of PRL on IL-6 was observed using visceral preadipocytes (data not shown). We also observed a progressive, marked suppression of IL-6 gene expression throughout LS14 differentiation, which could be functionally related to the early elevation of endogenous PRL. We have not been able to confirm the suppression of IL-6 during differentiation at the secretion level due to the fact that IL-6 levels are near the limit of detection for the ELISA used. Preliminary studies have also shown the suppression of leptin release by PRL (data not shown).
In summary, we have established a human adipocyte cell line with many characteristics of primary adipocytes. Unlike most transformed cells that are resistant to differentiation, these cells can undergo considerable morphological and functional differentiation under the appropriate culture conditions. LS14 cells open up new avenues for research on human adipocyte biology and add to the repertoire of nonpituitary, PRL-producing cell lines.
Footnotes
This work was supported by National Institutes of Health Grants ES-0955, ES-012212, and CA-096613; and Training Grant DK59803 (to E.R.H.).
Preliminary results of this investigation were presented at the Obesity and Diabetes Joint Keystone Conference, Banff, Alberta, Canada, March 2004.
First Published Online September 29, 2005
Abbreviations: aP2, Adipose fatty acid-binding protein; AR, -Adrenergic receptor; BAM, basal adipogenesis medium; B2M, 2-microglobulin; CM, conditioned medium; FBS, fetal bovine serum; GPDH, glycerol-3-phosphate dehydrogenase; h, human; IBMX, isobutylmethylxanthine; MTT, [3-(4,5-dimethylthiazol-2-yl)2,5-diphenyl tetrazolium bromide; PRL, prolactin; PRLR, prolactin receptor.
Accepted for publication September 22, 2005.
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