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Departments of Cell Biology (M.M., N.B.-J.) and Obstetrics and Gynecology (M.Z.), University of Cincinnati, College of Medicine, Cincinnati, Ohio 45267
Address all correspondence and requests for reprints to: Dr. Nira Ben-Jonathan, Department of Cell Biology, 3125 Eden Avenue, Cincinnati, Ohio 45267-0521. E-mail: Nira.Ben-Jonathan{at}uc.edu.
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Abstract |
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Introduction |
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TopAbstractIntroductionSubjects and MethodsResultsDiscussionReferences |
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PRL in humans is synthesized to a variable degree in many extrapituitary sites (2). Although an identical PRL protein is produced by all sites, its expression and release are differentially regulated. As illustrated in Fig. 1
, pituitary PRL is regulated by a proximal promoter that requires the Pit-1 transcription factor for transactivation (3). This promoter is inhibited by dopamine and stimulated by estrogens, neuropeptides, and growth factors (4). In contrast, PRL expression in the decidua (5), myometrium (6), and lymphocytes (7) is driven by a superdistal promoter, located 5.8 kb upstream of the pituitary start site. This promoter is silenced in the pituitary, does not bind Pit-1, and is not affected by dopamine, estrogens, or TRH (8). Exon 1a, serving as the alternative transcriptional initiation site, is spliced into exon 1b, yielding an identical coding region as the pituitary transcript, except for an additional 150-bp 5' untranslated region (UTR). The superdistal promoter contains activator protein-1, cAMP-response element binding protein, and CCAAT/enhancer binding protein sites, but the ligands that regulate its expression are poorly defined. Tissue-specific control of PRL release is exemplified by progesterone, which increases PRL synthesis in the endometrium (9), decreases it in the myometrium (10), and does not affect pituitary PRL.
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There is increasing evidence that local PRL is involved in breast cancer (15). PRL is expressed in normal and malignant breast tissues and breast cancer cell lines (16). Its function as a local mitogen is supported by partial suppression of T47D breast cancer cell proliferation by PRL antisense (17), anti-PRL antibodies (18), or PRL antagonists (19, 20). Also, treatment of nude mice carrying tumors derived from T47D cells with a PRL antagonist suppresses tumor growth (21), whereas PRL overexpression by MDA-MB-435 breast cancer cells accelerates tumor growth in nude mice and up-regulates the PRL receptor (PRL-R; Ref. 22). The latter suggests that local PRL can form an autocrine/paracrine loop with its receptor, thereby providing growth advantage to tumor cells. Notably, 80–90% of breast carcinomas express the PRL-R, with a higher expression in neoplastic than adjacent tissue (23).
The human breast is composed of small regions of cancer-prone epithelial cells surrounded by a sizable stroma made of adipose and connective tissue. Breast adipose tissue is made of 90–92% adipocytes, 6–7% preadipocytes, and 1–2% endothelial cells (24). Preadipocytes are unipotent fibroblasts that maintain a proliferative capacity until they undergo terminal differentiation to adipocytes in response to adipogenic factors (25). Interactions between the stromal and epithelial compartments are important for the development and morphogenesis of the ductal/alveolar structures and play a critical role in carcinogenesis. Notably, human breast cancer cells develop into much larger tumors and are capable of metastasis when inoculated into the mammary fat pad of nude mice than when implanted sc (26).
Our objective was to examine PRL production/release by human breast explants and determine whether it is regulated by ovarian steroids. The specific aims were to: 1) compare PRL release from glandular and adipose breast explants under serum-free conditions, 2) examine time-related changes in PRL gene expression and de novo synthesis, and 3) determine whether PRL release is regulated by estradiol and/or progesterone. We are reporting de novo synthesis of PRL by the human breast, with higher PRL release by adipose than glandular tissue.
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Subjects and Methods |
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Breast tissue was obtained from women undergoing mastectomy or breast reduction surgery. Their age, race, indication for surgery, and histological diagnosis are detailed in Table 1. Adipose tissue was obtained from 20 patients with an average age of 42.8 ± 3.9 (mean ± SEM) yr, whereas glandular tissue was obtained from 12 patients with an average age of 37.1 ± 5 yr. Informed consent was obtained from all women before surgery, and the study was performed under the approval of the Institutional Review Board of the University of Cincinnati.
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Breast specimens were placed immediately in DMEM/F12 (BD Biosciences, Bedford, MA) on ice. After several washes, blood vessels and connective tissue were removed, and glandular or adipose explants were dissected by gross appearance. Tissue samples (time 0) were fixed in 4% paraformaldehyde for hematoxylin and eosin (H&E) staining. Histological examination of selected slides shows approximately 5% cross-contamination between the two tissue types. Random 1–2 mm3 explants were either frozen at -70 C for RT-PCR analysis (d 0) or placed into 24-well plates (7–8 pieces per well, 3–4 wells per treatment) and incubated with 500 µl phenol red-free DMEM/F12 supplemented with ITS+ (BD Biosciences). Treatments consisted of media alone (control), estradiol (1 or 10 nM), progesterone (10 or 100 nM), or estradiol plus progesterone (10 nM estradiol and 100 nM progesterone). Steroids were dissolved in ethanol and diluted in DMEM/F12 media. The concentration of ethanol in the incubation media did not exceed 0.01%. Explants were incubated at 37 C under 5% CO2 for 10 d. Conditioned media (CM) were collected on d 1, 3, 7, and 10 of incubation and replaced with control or treatment media. The CM samples were immediately combined with an equal amount of Fischer’s treatment media (see next section), and duplicate aliquots were analyzed for PRL by the Nb2 bioassay. At the termination of the experiment, explants were blotted, weighed, and stored at -70 C for later RT-PCR analysis (d 10).
PRL bioassay
The rat Nb2 lymphocyte bioassay was used to verify release of bioactive PRL, taking advantage of its superior sensitivity over RIA. Nb2 lymphocytes (ATCC, Rockville, MD) were grown in Fischer’s leukemia media containing 10% heat-inactivated horse serum, 10% fetal bovine serum (FBS), 50 U/ml penicillin, 50 µg/ml streptomycin, and 5 µM 2-ß-mercaptoethanol. Before the assay, cells were incubated for 24 h in a starvation media prepared with Fischer’s leukemia media containing 10% charcoal-stripped FBS, 1% FBS, and 2-ß-mercaptoethanol. Cells were then seeded in 96-well plates (20,000 cells per well) in treatment media containing 10% heat-inactivated horse serum and 2-ß-mercaptoethanol. Quadruplicate aliquots of human PRL (hPRL) standard [National Institute of Diabetes and Digestive and Kidney Diseases (NIDDK)], amniotic fluid (1:1000) as an internal control, or duplicate aliquots of CM from the explants were added to each well and incubated with the Nb2 cells for 3 d. Cell number was determined by the MTT [3-(4,5-dimethylthiazol-2-yl) 2,5-diphenyltetrazolium bromide] method (27). Briefly, 125 µg MTT in PBS was added to each well and incubated for 2 h. A solubilizer, containing 50% N, N-dimethylformamide, 20% sodium dodecylsulfate, and 60 mM sodium acetate (pH 4.7) was added to each well and incubated overnight in the dark. OD at 570 nm was determined with an EL800 Universal Microplate Reader (Bio-Tek Instruments, Inc., Winooski, VT). The amount of PRL produced by the explants was calculated from the standard curve and was expressed as picograms of PRL per 100 mg wet weigh per 24 h. The assay sensitivity was 1 pg PRL/well.
RT-PCR
Total RNA was isolated from adipose and glandular explants on the day of surgery (d 0) and after 10 d of incubation using Tri Reagent (Molecular Research Center, Inc., Cincinnati, OH). The RNA was reverse-transcribed using Superscript II reverse transcriptase (Invitrogen, San Diego, CA) and random hexamers. PCR was performed on 0.75 µg of cDNA template with the following intron-spanning primers: 1) PRL coding: sense primer, 5'-CCTCTCCTCAGAAAGGTTCAGCG-3'; and antisense, 5'-TGTTGTTGTGGATGATTCGGC-3'; expected product size, 500 bp; 2) PRL noncoding: sense primer, 5'-CAAGAAGAATCGGAACATACAGGCTTT-3'; and antisense, 5'-GTCGATT TTATGTGAAGCCCTGCG-3'; expected product size, 760 bp; 3) PRL-R: sense primer, 5'-GACTTGCTGGTGGAGTATTTA-3'; and antisense, 5'-TGCCTTTCCCTCTTCTC TA-3'; expected product size, 495 bp; and 4) glyceraldehyde-3-phospate dehydrogenase (GAPDH): sense primer, 5'-CAAGTGGGGCGATG CTG-3'; and antisense, 5'-ACAGTCTTCTGGGTGGCAGTG-3'; expected product size, 309 bp. Cycle conditions for each transcript were optimized to fall within a linear range and were as follows: 94 C, 58 C, 72 C at 45 sec each for 35 cycles (28 cycles for GAPDH). Products were separated on a 2% agarose gel containing ethidium bromide and photographed.
Metabolic labeling
Glandular and adipose explants were incubated for 8 d as described above. Media were then replaced with a methionine- and cysteine-free DMEM to which 100 µCi 35S-metabolic labeling reagent containing L-methionine [35S] and L-cysteine [35S] (ICN Pharmaceuticals, Costa Mesa, CA) were added. Jurkat cells, a human lymphoid cell line that produces PRL (28), were plated at 1 x 106 cells per well and served as a positive control. After a 6-h incubation, media were removed, and 100 ng hPRL were added as a cold carrier. Media were concentrated by spin filtering and resuspended in TSA buffer (10 mM Tris-HCl, 0.14 M sodium chloride, and 0.025% sodium azide) containing 1% Triton X-100, 1% BSA, 1 mM iodoacetamide, 5 µg/ml aprotinin, 5 µg/ml leupeptin, and 1 mM phenylmethylsulfonyl fluoride. Breast explants or Jurkat cells were extracted with TSA buffer to which 100 ng of hPRL were added. Samples were precleared by overnight incubation at 4 C with normal rabbit serum (NRS), followed by incubation with protein A Sepharose (Pierce Chemical Co., Rockford, IL) for 2 h. After centrifugation, supernatants were incubated at 4 C for 4 h with anti-hPRL antibodies. These were generated in our laboratory by injecting rabbits with highly purified hPRL (NIDDK). Extensive characterization of these antibodies showed similar avidity and specificity as the anti-hPRL antibodies provided by the NIDDK. After incubation, samples were treated with protein A Sepharose for 2 h, followed by centrifugation at 4 C. The pellet containing the immune complex was resuspended in laemmli loading buffer (BRL, Invitrogen), boiled, and loaded onto 12% SDS-PAGE. After electrophoresis, the gel was fixed in a 10% methanol/10% acetic acid solution, placed in Amplify (Amersham Pharmacia Biotech, Piscataway, NJ) to enhance the signal, and autoradiographed.
Data analysis
When appropriate, results are expressed as the mean ± SEM. Mean values were derived by averaging the PRL results from the individual wells for each patient, with the N representing the number of patients. Statistical differences within groups were determined by using one-way ANOVA followed by post hoc analysis (SigmaStat 4.0, SPSS, Inc., Chicago, IL). P values less than 0.05 were considered statistically significant.
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Results |
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The conditions for Nb2 bioassay for PRL were optimized to increase sensitivity. As shown in Fig. 2, the OD of MTT, reflecting the number of Nb2 cells, showed a curvilinear relationship with the amount of standard hPRL from 1–60 pg/well. Addition of 5, 10, or 20 µl duplicate aliquots of CM, collected from glandular or adipose tissue explants after 1 d of incubation, showed parallelism with the standard curve. A similar dose-response relationship was evident with diluted amniotic fluid samples (data not shown). To verify that increased Nb2 cell proliferation in response to CM was due to PRL only, we used immunoneutralization with anti-hPRL antibodies. Nb2 cells were incubated for 3 d with various aliquots of CM or highly diluted human amniotic fluid samples (as a positive control) alone, together with NRS (1:1000), or with anti-hPRL antibodies (1:1000). Figure 3 shows that addition of PRL antibodies abrogated the Nb2 cell proliferative response to CM from either glandular or adipose explants as well as the amniotic fluid. On the other hand, cell proliferation in the presence of NRS did not differ from control.
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Both glandular and adipose breast explants incubated under serum-free conditions secreted measurable amounts of PRL. In glandular tissue, the average rate of PRL release was 64 pg/100 mg·24 h on the first day, with a significant reduction to 20 pg/100 mg·24 h by d 3 (Fig. 4). In contrast, the rate of 42 pg PRL/100 mg·24 h seen on d 1 of adipose tissue incubation remained unchanged by d 3. From d 3 on, PRL release rate increased in both tissues. In glandular tissue, the release rate was 3- and 4-fold higher on d 7 and 10, respectively, than on d 3 (P < 0.05). In adipose explants, PRL release rate increased in a linear manner from d 3 to 10, achieving an 11-fold higher rate on d 10 than during the first 3 d (P < 0.05). On d 10, the release rate of PRL from adipose tissue was more than 6-fold higher (P < 0.05) than that from glandular tissue. Figure 4 also shows H&E-staining of representative glandular and adipose explants, depicting the differences in cell size and morphology between adipocytes and the glandular epithelium.
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Total RNA, extracted from glandular and adipose explants on d 0 and after 10 d in culture, was examined by RT-PCR for relative levels of PRL, extrapituitary PRL (using an upstream primer annealing to exon 1a; Fig. 1
) and PRL-R transcripts; GAPDH was used as a control. Representative results from two patients are shown in Fig. 5. As evident, the PRL transcripts were barely detectable in glandular tissue on d 0 but were clearly seen on d 10. The PRL transcripts in adipose tissue were low on d 0 and increased markedly on d 10. PRL-R expression in adipose tissue was generally lower than that in glandular tissue. Similar results were obtained in samples from four additional patients (data not shown).
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To verify de novo PRL synthesis rather than release from storage depots, we performed metabolic labeling. On d 8 of culture, glandular and adipose explants were incubated with 35S-methionine/cysteine for 6 h, followed by PRL immunoprecipitation, gel electrophoresis, and autoradiography; Jurkat cells expressing PRL were used as a positive control. As shown in Fig. 6, metabolically labeled PRL was detected in both tissue extracts and CM, providing a strong evidence for de novo synthesis of PRL by both types of explants.
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Glandular and adipose explants were incubated with various doses of estradiol and progesterone, and the media were collected and completely replaced on d 1, 3, 7, and 10. As shown in Fig. 7, top panel, progesterone at either 10 or 100 nM significantly attenuated the delayed rise in PRL release from glandular explants between d 3 and 10. A combination of 10 nM estradiol and 100 nM progesterone resulted in a similar inhibition of PRL release (data not shown). On the other hand, estradiol alone at either dose did not significantly alter PRL release from glandular explants. Neither hormone affected the time-dependent rise in PRL release from adipose explants (Fig. 7, bottom panel). Note the differences in the Y axis scale between the adipose and glandular explants.
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Discussion |
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Expecting low rate of PRL release from breast explants, which might not be detected by RIA, we optimized the Nb2 bioassay and achieved a 25- to 50-fold higher sensitivity than RIA. Nb2 cells are immature rat T-lymphocytes that express a unique intermediate form of the PRL-R (30) and depend on PRL for proliferation/survival (31). This assay has replaced the pigeon crop sac assay as the most widely used bioassay for PRL, and its specificity for PRL has been well established (32, 33). By configuring the assay to 96-well plates, the incubation volume was decreased, thus increasing assay efficacy. The 96-well format also enables the use of MTT for measuring cell number by means of an optical microplate reader. By using immunoneutralization, we confirmed that increased Nb2 cell number in response to CM from breast explants is due only to the presence of PRL.
PRL release from glandular explants decreased significantly between d 1 and 3 of incubation, followed by a 4-fold increase (Fig. 7). We believe that the initial decline is due to depletion of prestored PRL, whereas the subsequent increase results from de novo synthesis that is coupled to release. Prestored PRL could originate from local synthesis as well as via uptake and retention of circulating PRL. Uptake is a mechanism by which PRL is transported into fluid compartments such as cerebrospinal fluid (34) and milk (35, 36). Because hPRL binds to heparin (37), it has the capacity to be retained by proteoglycans in the extracellular matrix, increasing its effective concentration in the vicinity of producing/responsive cells. Unlike pituitary PRL, decidual PRL is not stored in secretory granules and is not subjected to calcium-dependent exocytosis (38), and this could also be the case with breast PRL.
Pituitary PRL is under tonic inhibition by dopamine (39). When lactotrophs are placed in culture, removed from hypothalamic inhibition, PRL synthesis and release increase in a time-dependent manner. We believe that such a situation is recapitulated in extrapituitary sites, albeit by a nondopaminergic inhibitory mechanism. Indeed, PRL release from cultured decidual (40) and myometrial (41) cells, dermal fibroblasts (42), and breast tissue (Figs. 4 and 7) increases progressively over time, raising the possibility that PRL is normally kept suppressed by tissue-specific inhibiting factor(s). Because progesterone inhibits PRL release from glandular explants, it may serve as a potential candidate for the in vivo inhibitory agent. However, other local or blood-born factor(s) cannot be ruled out. Inhibition of PRL release from nonmalignant human breast explants by a high dose of progesterone (10 µM) after 48-h incubation was recently reported (43). Interestingly, these authors found that progesterone increased PRL release from malignant explants that were estrogen receptor negative, progesterone receptor positive, but not from other types of explants.
Using RT-PCR, we found noticeable increase in PRL expression in both glandular and adipose tissue after 10 d of incubation, matching the profile of PRL release. The use of primers that anneal to exon 1a showed that PRL in breast tissue is likely transcribed from the decidual start site (Fig. 5). Although these data confirm the similarity of breast PRL transcript to that in other extrapituitary sites (i.e. having a longer 5'UTR sequence), the identity of the superdistal promoter should be verified before examining the control of PRL gene expression in the breast. The detection of metabolically labeled PRL in both CM and cell extracts from glandular and adipose tissue (Fig. 6) provided a further support for de novo synthesis of PRL by the human breast.
Glandular tissue from breast with invasive or metastatic breast cancer was excluded from this study (Table 1). However, we have examined PRL release from tumor explants from five patients and found that it resembled the pattern seen in normal glandular tissue rather than the marked time-dependent rise seen in adipose tissue. Notably, two highly invasive and metastatic tumors showed a high initial PRL release, which was subsequently reduced over time in culture. A larger number of specimens will be needed before a clear pattern of PRL production/release from tumors can be established. Nonetheless, it is of interest that PRL release from cultured leiomyomas is significantly higher than that from the normal myometrium (41), suggesting that tumorigenesis may be associated with increased local PRL production due to inactivation or loss of responsiveness to an inhibitor, an increase of a stimulator, or both. In future studies we will attempt to compare PRL release from tumors, nonmalignant tissue surrounding the tumors, and nonadjacent glandular/adipose tissue.
The most striking, and unexpected, finding of this investigation is the increased PRL production by adipose tissue. When tissues were harvested, PRL mRNA levels were very low, providing a plausible explanation why PRL production by adipose tissue has been overlooked. Because adipose tissue comprises the bulk of the stroma, it potentially constitutes the richest source of breast PRL. Adipose tissue produces and secretes many cytokines and hormones that act locally or at distant sites. We propose that PRL should be added to the list of locally produced cytokines. Based on very preliminary immunohistochemical data, we suspect that PRL may be produced by preadipocytes. If confirmed, PRL production per cell is not insignificant, given that preadipocytes constitute only a minor cell population within adipose tissue. Such cells could be a source of relatively high PRL concentration in the vicinity of tumors that are often surrounded by a layer of adipose-derived fibroblasts that provide structural and biochemical support for breast cancer (44).
The incidence of obesity has been rapidly increasing in the industrial countries and is associated with a wide variety of health problems such as diabetes, heart disease, hypertension, and increased risk of cancer, including breast cancer. Although the focus of our investigation has been on the action of PRL as a local mitogenic factor in breast cancer, the PRL-R is also expressed by adipocytes (45), in which PRL suppresses cytokine-mediated signaling and inhibits insulin-induced leptin release (46). Notably, adult female PRL-R-deficient mice have a 50% reduction in total abdominal fat mass and a 30% reduction in fat mass expressed as a percentage of body weight (47). Thus, adipose-derived PRL can have functions beyond breast cancer, including local action on preadipocytes/adipocytes. Adipocytes exhibit depot- and gender-specific receptor expression, hormonal responsiveness, and aromatase activity (48), indicating that the local microenvironment affects their cellular properties. Therefore, it would be of great interest to determine whether PRL exhibits a similar pattern of expression and release in other fat depots, e.g. visceral and sc, to identify the adipose cell type that produces PRL, and to examine for interaction with adipogenic hormones. Another major issue that should be resolved is whether PRL production by adipose tissue is unique to humans or is also seen in other species.
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Acknowledgments |
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Footnotes |
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M.Z. and M.M. contributed equally to this investigation.
Abbreviations: CM, Conditioned media; FBS, fetal bovine serum; GAPDH, glyceraldehyde-3-phospate dehydrogenase; H&E, hematoxylin and eosin; hPRL, human PRL; MTT, 3-(4,5-dimethylthiazol-2-yl)2,5-diphenyl tetrazolium bromide; NRS, normal rabbit serum; PRL, prolactin; PRL-R, PRL receptor; UTR, untranslated region.
Received August 12, 2002.
Accepted October 18, 2002.
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References |
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-subunit stimulate prolactin production by explant cultures of human leiomyoma and myometrium. Am J Obstet Gynecol 170:677–683[Medline]
and interleukin 11 secreted by malignant breast epithelial cells inhibit adipocyte differentiation by selectively down-regulating CCAAT/enhancer binding protein and peroxisome proliferator-activated receptor 
: mechanism of desmoplastic reaction. Cancer Res 61:2250–2255This article has been cited by other articles:
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  N. Ben-Jonathan, C. R. LaPensee, and E. W. LaPensee What Can We Learn from Rodents about Prolactin in Humans? Endocr. Rev., February 1, 2008; 29(1): 1 - 41. [Abstract] [Full Text] [PDF]
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 M. Kutsukake, R. Ishihara, M. Yoshie, H. Kogo, and K. Tamura Involvement of insulin-like growth factor-binding protein-related protein 1 in decidualization of human endometrial stromal cells Mol. Hum. Reprod., October 1, 2007; 13(10): 737 - 743. [Abstract] [Full Text] [PDF]
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 U. A. White, A. A. Coulter, T. K. Miles, and J. M. Stephens The STAT5A-Mediated Induction of Pyruvate Dehydrogenase Kinase 4 Expression by Prolactin or Growth Hormone in Adipocytes Diabetes, June 1, 2007; 56(6): 1623 - 1629. [Abstract] [Full Text] [PDF]
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 O. Eyal, J.-B. Jomain, C. Kessler, V. Goffin, and S. Handwerger Autocrine Prolactin Inhibits Human Uterine Decidualization: A Novel Role for Prolactin Biol Reprod, May 1, 2007; 76(5): 777 - 783. [Abstract] [Full Text] [PDF]
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 C. R. LaPensee, N. D. Horseman, P. Tso, T. D. Brandebourg, E. R. Hugo, and N. Ben-Jonathan The Prolactin-Deficient Mouse Has an Unaltered Metabolic Phenotype Endocrinology, October 1, 2006; 147(10): 4638 - 4645. [Abstract] [Full Text] [PDF]
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 A. Diogenes, A. M. Patwardhan, N. A. Jeske, N. B. Ruparel, V. Goffin, A. N. Akopian, and K. M. Hargreaves Prolactin Modulates TRPV1 in Female Rat Trigeminal Sensory Neurons J. Neurosci., August 2, 2006; 26(31): 8126 - 8136. [Abstract] [Full Text] [PDF]
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E. R. Hugo, T. D. Brandebourg, C. E. S. Comstock, K. S. Gersin, J. J. Sussman, and N. Ben-Jonathan LS14: A Novel Human Adipocyte Cell Line that Produces Prolactin Endocrinology, January 1, 2006; 147(1): 306 - 313. [Abstract] [Full Text] [PDF]
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V. Goffin, S. Bernichtein, P. Touraine, and P. A. Kelly Development and Potential Clinical Uses of Human Prolactin Receptor Antagonists Endocr. Rev., May 1, 2005; 26(3): 400 - 422. [Abstract] [Full Text] [PDF]
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, higher than d 3 (P < 0.05). Each value is a mean ± SEM. Right, photographs of H&E staining of paraffin-embedded adipose and glandular explants on d 0. 
, Adipocyte; arrow, glandular epithelium. Note the difference in cell size and morphology.
