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American Journal of Physiology - Endocrinology and Metabolism logoLink to American Journal of Physiology - Endocrinology and Metabolism
. 2009 Aug 18;297(5):E1089–E1096. doi: 10.1152/ajpendo.00209.2009

Progesterone stimulates mitochondrial activity with subsequent inhibition of apoptosis in MCF-10A benign breast epithelial cells

Millie A Behera 1,*, Qunsheng Dai 1,*, Rachana Garde 1, Carrie Saner 1, Emily Jungheim 1, Thomas M Price 1,✉
PMCID: PMC2781356  PMID: 19690070

Abstract

The effects of progesterone on breast epithelial cells remain poorly defined with observations showing both proliferative and antiproliferative effects. As an example, progesterone levels correlate with increased epithelial cell proliferation, but there is discordance between the dividing cells and the cells with nuclear progesterone receptor expression. The release of paracrine growth factors from nuclear receptor-positive cells has been postulated as a mechanism, since in vitro studies show a lack of growth effect by progesterone in breast epithelial cells lacking nuclear receptors. This study examined possible nongenomic effects of progesterone in breast epithelia by using MCF-10A cells known to lack nuclear progesterone receptor expression. Treatment for 30–60 min with progesterone or the progestin, R5020, increased mitochondrial activity as shown by an increase in mitochondrial membrane potential (hyperpolarization) with a concordant increase in total cellular ATP. The reaction was inhibited by a specific progesterone receptor antagonist and not affected by the translation inhibitor cycloheximide. Progestin treatment inhibited apoptosis induced by activation of the FasL pathway, as shown by a decrease in sub-G1 cell fraction during fluorescence-activated cell sorting and a decrease in caspase 3/7 levels. Progestin treatment did not alter the cell cycle over 48 h. Our study demonstrates a nongenomic action of progesterone on benign breast epithelial cells, resulting in enhanced cellular respiration and protection from apoptosis.

Keywords: progesterone, MCF-10A cells, mitochondria, apoptosis, cellular respiration

previous studies have shown a discordance between the proliferative action of progesterone in the breast and the content of nuclear progesterone receptors (PR-B and PR-A) in breast epithelial cells. As examples, the mitotic rate of breast epithelial cells is greatest in the progesterone-dominant luteal phase (44). Exogenous administration of estrogen plus progestin results in greater breast epithelial proliferation than estrogen alone (15, 18). The progesterone receptor knockout (PRKO) mouse, an excellent model for the function of nuclear PR in mammary gland development, supports a proliferative role. PR-B is primarily responsible for gland development, with PRKO-B mice showing less extensive ductal development and lack of alveolar terminal end buds (7, 29). In contrast to these physiological effects, very few breast epithelial cells appear to express nuclear PR. In immunocytochemical studies of human breast tissue, from 6 to 13% of ductal epithelial cells express PR (6, 37), with the two PR isoforms, B and A, typically localizing in the same cell (14). Immunocytochemical analysis of proliferation by detection of Ki67 antigen in human breast or autoradiographs of [3H]thymidine incorporation in rodent mammary samples show nuclear PR-expressing cells to be nonproliferating. Adjacent cells lacking nuclear PR expression have greater mitotic activity (23). This disassociation between nuclear PR expression and proliferation led to the hypothesis that nuclear PR-expressing cells regulate proliferation of adjacent cells via the control of paracrine factors (4).

This hypothesis makes the supposition that progesterone has no effect on breast ductal cells lacking nuclear PR expression. This supposition is supported by the lack of progesterone-induced proliferation of breast explants in nude mice (5), of immortalized breast epithelial cells lacking nuclear PR expression (25), and in breast cancer cell lines lacking nuclear PR (40). In contrast to these observations, recent studies with the immortalized nuclear PR-negative breast epithelial cell line, MCF-10A, show a progestin-induced growth response when combined with tyrosine kinase growth factors (24, 25). Because proliferation is a complex action requiring activation of many cellular pathways, we hypothesized that progesterone/progestin may have metabolic effects via nongenomic regulation.

In this study, we used the MCF-10A cell line to analyze the effect of progesterone/progestin on cellular respiration and the resulting influence on apoptosis and cell cycle. Despite the lack of a nuclear PR, these cells demonstrate a clear response to progesterone/progestin with an increase in mitochondrial membrane potential and cellular ATP production. This increase in cellular respiration correlates with a protective effect against Fas-mediated apoptosis.

METHODS

Cell lines.

The MCF-10A breast epithelial cell line was obtained from the American Type Culture Collection (Manassas, VA). Cells were grown in Mammary Epithelial Growth Medium supplemented with 100 ng/ml cholera toxin. MCF-10A cells were obtained varying from 99 to 116 passages. Cells were tested during a maximum of six serial passages.

Mitochondrial membrane potential.

Mitochondrial membrane potential (MMP) was determined with 5,5′,6,6′-tetrachloro-1,1′,3,3′-tetraethylbenzimidazolecarbocyanide iodine (JC-1) dye (Invitrogen). Cells (125,000/well) were grown overnight in Costar 48-well plates (Corning, Corning, NY) in media and then placed in modified Krebs-Ringer-HEPES buffer (KRH) containing 25 mM Na-HEPES, 115 mM NaCl, 5 mM KCl, 1 mM KH2PO4, 1.2 mM MgSO4, 0.5 mM CaCl2, and 5 mM glucose at pH 7.4 for 2 h before experimentation. Cells were pretreated for 30–60 min with ligands, including progesterone or R5020 (Perkin-Elmer, Wellesley, MA) in the absence or presence of the PR antagonist RTI-6413-049b (41). In other experiments, cells were treated with dexamethasone (Sigma) with or without the human glucocorticoid receptor (hGR) inhibitor AL082D06 (Ligand Pharmaceuticals). In some experiments, the translation inhibitor cycloheximide (5 μg/ml final concentration) was added when the cells were placed in buffer. Control experiments included treatment with pyruvate, ADP, and A-23187. Cells were then treated with 2.5 μg/ml JC-1 for 10 min. With the use of a Tecan Safire multichannel plate reader, cells were excited at 488 nm, and emission was determined at 529 and 585 nm.

Total cellular ATP determination.

ATP was determined in MCF-10A cells grown in 96-well plates, plated at a density of 50,000 cells/well. Cells were grown overnight in media and then placed in modified KRH buffer for 2 h before experimentation. Cells were treated for 2 h with R5020 with or without the addition of the PR antagonist RTI-6413-049b. ATP was determined with a bioluminescent assay (Sigma). This assay is based on the conversion of ATP + Luciferin → adenyl-luciferin + pyrophosphate (PPi), by the enzyme firefly luciferase and subsequent reaction of adenyl-luciferin + O2 → oxyluciferin + AMP + CO2 + light. For the reaction, 100 μl of sample is added to 100 μl of reaction mix containing luciferase, luciferin, MgSO4, dithiothreitol, EDTA, BSA, and tricine buffer, and immediately analyzed in a luminometer. Cells are first permeabilized as to allow escape of ATP by a Somatic Cell Releasing Agent (Sigma-Aldrich). Light emission was determined in a luminometer.

Identically treated wells (3/condition) in the same plate were used for total protein determination. Cells were lysed with the addition of 1 N NaOH. Protein content was then determined on an aliquot of the lysate with a Bradford reagent using known concentrations of BSA for a standard curve.

Induction of apoptosis.

MCF-10A cells were plated in six-well plates for FACS analysis or 96-well plates for caspase analysis. At 60% confluence, apoptosis was induced by treating with an activating mouse monoclonal anti-Fas antibody (IgM, 0.5 μg/ml, clone CH11; Upstate Biotechnology, Lake Placid, NY), with or without R5020. Controls included cells treated with ethanol vehicle and cells treated with normal mouse IgM (Santa Cruz Biotechnology, Santa Cruz, CA). Cells were treated for 24 h for caspase analysis and for 30 h for fluorescence-activated cell sorter (FACS) analysis at 37°C in 5% CO2.

FACS.

Flow cytometry was used to assess DNA content as a surrogate measure for cell cycle phase. At the end of the treatment period, cells were harvested by trypsinization and resuspended in cold Hanks' balanced salt solution (Invitrogen, Carlsbad, CA). Cells were fixed in ethanol and stored at −20°C for at least 18 h. Fixed cells were resuspended in 500 μl of PBS containing 100 μg/ml propidium iodide (Sigma) and 100 μg of RNase A (Sigma). The cells were then analyzed on a FACS Calibur (Becton-Dickinson) with the DNA data, and the cell cycle stages were analyzed using ModFit LT 3.0 software. Within each experiment, samples were analyzed in duplicate.

Caspase assay.

Caspase 3/7 activity was determined by an enzymatic fluorescent assay. At the end of the 24-h treatment period, 100 μl of freshly mixed Apo-ONE Caspase-3/7 Reagent was added to each well of the plate following the manufacturer's protocol (Promega, Madison, WI). Cell lysis was enhanced by a freeze-thaw cycle two times at −70°C and room temperature (RT). Plates were shaken at 300–500 revolutions/min for 2 h at RT. Fluorescence of each well was determined by excitation at 499 nm and emission at 521 nm using a Tecan Safire multichannel plate reader.

RT-PCR analysis of kallikrein 10 expression.

MCF-10A cells were treated with 10−6 M dexamethasone with or without 10−6 M AL082D06 or 10−6 M AL082D06 alone for 48 h and ethanol and dimethyl sulfoxide as a control. Total RNA was extracted from treated cells using TRIzol reagent according to the manufacturer's instruction (Invitrogen). Total RNA (1 μg) was used for first-strand cDNA synthesis by SuperScript III RT with an oligo(dT) primer, and semiquantitative PCR was performed for amplifying both kallikrein 10 and glyceraldehydes-3-phosphate dehydrogenase (GAPDH) gene under the same PCR condition, 25 cycles each, as described by the manufacturer's instructions (Invitrogen). The primers for kallikrein 10 were as follows: 5′-GGAAACAAGCCACTGTGGGC-3′ (forward) and 5′-GAGGATGCCTTGGAGGGTCTC-3′ (reverse), and for GAPDH were 5′-GAAGGTGAAGGTCGGAGTC-3′ (forward) and 5′-GAAGATGGTGATGGGATTTC-3′ (reverse). RT-PCR products were 467 and 225 bp.

Cycloheximide inhibition of transforming growth factor-β1 induction of inhibitor of DNA-binding protein 1.

MCF-10A cells were pretreated for 2 h with KRH + glucose buffer with or without 5 μg/ml cycloheximide. Transforming growth factor (TGF)-β1 (5 ng/ml) was added for 60 min. Total protein was isolated by centrifugation in PBS with protease inhibitors (PI), resuspension in RIPA/PI buffer, and sonicated. Protein concentration was determined as above. For Western blot analysis, 50 μg protein were separated on a 4–15% polyacrylamide gel in Tris-glycine-SDS buffer. Protein was transferred to a polyvinylidene difluoride membrane in Tris-glycine buffer. The membrane was blocked with 5% milk for 1 h at RT, incubated with 1:500 dilution of rabbit polyclonal anti-inhibitor of DNA-binding protein 1 (Id-1) antibody (Santa Cruz Biotechnology) overnight at 4°C, and then incubated with goat anti-rabbit secondary antibody (1:2,000) for 1 h at RT. The membrane was developed with ECL Reagent (Amersham, Buckinghamshire, UK) per the manufacturer's recommendation and exposed to film.

Statistical analysis.

Analyses of MMP, ATP, and caspase activity were performed in multiwell plates. One experiment represented the results of one plate, and experiments were performed in duplicate or triplicate as noted in the legends for Figs. 1–6. FACS analysis was performed with treatments in individual tubes for a total of three experiments. Numbers in Figs. 1–6 show the total number of replicates in the experiments. Repeated-measures analysis was performed with SPSS software for experiments in which replicate analyses were performed at different times. In groups showing an overall significant difference, Tukey testing was used to evaluate differences between treatment groups. Statistical significance was considered a P ≤ 0.05. Results are expressed as means ± SE.

Fig. 1.

Fig. 1.

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Mitochondrial membrane potential (MMP) determined by 5,5′,6,6′-tetrachloro-1,1′,3,3′-tetraethylbenzimidazolecarbocyanide iodine (JC-1) fluorescent emission. A: MCF10-A cells were placed in buffer for 2 h and then treated for 60 min with varying concentrations of progesterone. JC-1 was excited at 488 nm, and emission was determined at 529 and 585 nm. The 585-to-529 nm ratio represents membrane potential, with a dose-dependent hyperpolarization evident. B: with identical methods, a dose-dependent hyperpolarization was seen with the specific progesterone receptor (PR) agonist R5020. C: R5020 (10−8 M)-induced hyperpolarization was completely inhibited by concomitant treatment with the PR antagonist RTI-6413-049b (10−6 M). D: as a control, the increase in MMP with R5020 was compared with 10−5 M ADP. Nos. in bars represent total observations. For A–C, assays were performed in triplicate, while in duplicate for D. Significant differences include 10−6 M vs. ethanol (EtOH) (P = 0.005), 10−6 vs. 10−8 M (P = 0.045) (A) and 10−6 vs. 10−8 M (P = 0.031), 10−6 M vs. EtOH (P < 0.001), 10−7 M vs. EtOH (P < .001), and 10−8 M vs. EtOH (P = 0.052) (B). C: R5020 (R) 10−8 M vs. R + RTI-6413-049b (RTI) (P < 0.001), R 10−8 M vs. RTI (P < 0.001), and R 10−8 M vs. EtOH (P < 0.001). D: ADP vs. EtOH (P < 0.001), R 10−6 M vs. EtOH (P < 0.001), and R 10−7 M vs. EtOH (P = 0.003).

Fig. 2.

Fig. 2.

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Evaluation of MMP changes with cycloheximide (CHX) pretreatment. MCF-10A cells were placed in buffer for 2 h with or without 5 μg/ml CHX and then treated for 30 min with 10−6 M R5020. Pretreatment with CHX did not inhibit the R5020-induced hyperpolarization. Nos. in bars represent total observations, and assays were performed in triplicate. Significant differences include R vs. CHX (P < 0.001), R vs. EtOH (P < 0.001), R + CHX vs. CHX (P < 0.001), R + CHX vs. EtOH (P < 0.001), and CHX vs. EtOH (P = 0.04).

Fig. 3.

Fig. 3.

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Comparison of MMP changes with glucocorticoid treatment. A: MCF-10A cells were placed in buffer for 2 h and then treated for 60 min with dexamethasone at concentrations of 10−6 to 10−8 M. Nos. in bars represent total observations, and assays were performed in triplicate. There was no significant overall difference in the treatments. B: MCF-10A cells were treated as above with R5020 (10−6 M) in the presence or absence of AL082D06 (D06). No diminution of the R5020 increase in MMP was noted. Because of a limited amount of available AL082D06, this assay was only performed one time. A: no significant difference in the group. B: significant differences include R vs. D06 (P < 0.001), R vs. dimethyl sulfoxide (DMSO) + EtOH (P < 0.001), R vs. DMSO (P < 0.001), R vs. EtOH (P < 0.001), R + D06 vs. D06 (P < 0.001), R + D06 vs. DMSO + EtOH (P < 0.001), R + D06 vs. DMSO (P < 0.001), and R + D06 vs. EtOH (P < 0.001).

Fig. 4.

Fig. 4.

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ATP determination by bioluminescent assay in MCF-10A cells. A: cells were placed in buffer for 2 h and then treated for 2 h with R5020 (10−6 M) with or without the PR antagonist RTI-6413-049b (10−5 M). Total ATP levels were greater in R5020-treated cells compared with controls and antagonist-treated cells. B: to ensure that changes in ATP were not because of a difference in cell number, total protein levels were determined by Bradford analysis. Nos. in bars represent total observations. Assays were performed in triplicate. Significant differences include R vs. R + RTI (P < 0.001), R vs. RTI (P < 0.001), and R vs. EtOH (P < 0.001) (A) and no significant difference among groups (B).

Fig. 5.

Fig. 5.

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Percentage of sub-G1 cells after treatment with Fas antibody and R5020. Cells were treated in media for 30 h with 0.5 μg/ml activating Fas antibody and R5020 at 10−6 and 10−8 M. Cells in the sub-G1 fraction were determined by fluorescence-activated cell sorter (FACS) analysis. R5020 inhibited Fas antibody-induced apoptosis. Nos. in bars represent total observations. Assays were performed in triplicate. Controls included cells treated with EtOH, normal mouse IgM, and untreated. Significance differences include: FasL vs. FasL + R 10−6 M (P = 0.002), FasL vs. FasL + R 10−8 M (P < 0.001), FasL vs. R 10−6 M (P < 0.001), FasL vs. R 10−8 M (P < 0.001), FasL vs. UT (P < 0.001), FasL vs. IgM (P < 0.001), FasL vs. EtOH (P < 0.001), FasL + R 10−6 M vs. R 10−6 M (P < 0.001), FasL + R 10−6 M vs. R 10−8 M (P < 0.001), FasL + R 10−6 M vs. UT (P < 0.001), FasL + R 10−6 M vs. IgM (P < 0.001), FasL + R 10−6 M vs. EtOH (P < 0.001), FasL + R 10−8 M vs. R 10−6 M (P < 0.001), FasL + R 10−8 M vs. R 10−8 M (P < 0.001), FasL + R 10−8 M vs. UT (P < 0.001), FasL + R 10−8 M vs. IgM (P < 0.001), FasL + R 10−8 M vs. EtOH (P < 0.001).

Fig. 6.

Fig. 6.

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Caspase 3/7 activity after treatment with activating Fas antibody and R5020. A: results of cells treated in media for 24 h with 0.5 μg/ml activating Fas antibody and differing concentrations of R5020. Caspase 3/7 activity was determined by fluorescent assay. A dose-dependent decrease in activity is seen with R5020. B: results of cells treated in the same manner with activating Fas antibody with or without R5020 (10−7 M) and the PR antagonist RTI-6413-049b (10−7 M). Addition of RTI-6413-049b prevented R5020 inhibition of FasL-induced caspase activation. Nos. in bars represent total observations. Assays in A were performed in triplicate and those in B in duplicate. Significant differences include: FasL vs. FasL + R 10−6 M (P < 0.001), FasL vs. FasL + R 10−7 M (P < 0.001), FasL vs. R 10−6 M (P < 0.001), FasL vs. EtOH + IgM (P < 0.001), FasL vs. untreated (P < 0.001), FasL + R 10−6 M vs. FasL + R 10−8 M (P < 0.001), FasL + R 10−6 M vs. EtOH + IgM (P < 0.001), FasL + R 10−6 M vs. untreated (P < 0.001), FasL + R 10−7 M vs. FasL + R 10−8 M (P = 0.01), FasL + R 10−7 M vs. R 10−6 M (P < 0.001), FasL + R 10−7 M vs. EtOH + IgM (P < 0.001), FasL + R 10−7 M vs. untreated (P < 0.001) (A) and FasL vs. FasL + R 10−7 M (P < 0.001), FasL vs. R 10−7 M + RTI (P < 0.001), FasL vs. EtOH + IgM + DMSO (P < 0.001), FasL + R 10−7 M vs. FasL + R 10−7 M + RTI (P = 0.004), FasL + R 10−7 M vs. R 10−7 M + RTI (P < 0.001), FasL + R 10−7 M vs. EtOH + IgM + DMSO (P < 0.001), FasL + R 10−7 M + RTI vs. R 10−7 M + RTI (P < 0.001), FasL + R 10−7 M + RTI vs. EtOH + IgM + DMSO (P < 0.001) (B).

RESULTS

Figure 1 shows the effect of a 60-min treatment with progesterone and the synthetic progestin, R5020, on MMP in MCF-10A breast epithelial cells. MMP was determined with the cationic dye JC-1. JC-1 manifests a membrane potential accumulation in the mitochondria with a shift from green to red fluorescent emission. The ratio of 585/529 nm reflects MMP, with a decrease in the ratio showing depolarization and an increase in the ratio consistent with hyperpolarization. A dose-dependent increase in MMP (hyperpolarization) is seen after treating cells with either progesterone or R5020. The reaction is inhibited by the concomitant addition of the PR antagonist RTI-6413-049b. As a positive control, cells were treated with 10−5 M ADP, a known substrate for mitochondrial ATP production. The ADP-induced increase in MMP was similar in magnitude to 10−6 M R5020. As an additional control, cells treated with the calcium ionophore A-23187 (20 μM) showed a significant mitochondrial membrane depolarization [Supplemental Fig. 1 (Supplemental data for this article may be found on the American Journal of Physiology - Endocrinology and Metabolism web site.)].

Figure 2 shows the lack of inhibition of MMP hyperpolarization by R5020 (10−6 M) when cells were pretreated with cycloheximide (5 μg/ml). Cycloheximide was added 2 h before the addition of the R5020. To ensure the dose of cycloheximide was adequate to inhibit protein synthesis, cells were treated in the same manner with TGF-β1, and inhibitor of DNA-binding protein 1 (Id-1) levels were determined. Supplemental Fig. 2 shows inhibition of Id-1 expression with a 2-h cycloheximide pretreatment.

Because of the potential activation of a glucocorticoid receptor (GR) by progestins, the effect of dexamethasone was evaluated. Figure 3A shows no significant difference in MMP after a 60-min exposure to 10−6 to 10−8 M dexamethasone relative to control. For further analysis, the MMP assay was performed with 10−6 M R5020 in the presence or absence of the specific GR antagonist AL082D06. No significant diminution of the R5020 effect was seen. As an additional control, AL082D06 was demonstrated to antagonize dexamethasone-mediated inhibition of kallikrein 10 transcript levels in MCF-10A cells. As shown in Supplemental Fig. 3, 10−6 M AL082D06 prevented the inhibition of kallikrein 10 expression by 10−6 M dexamethasone.

We also analyzed for a potential androgen effect on MMP. Dihydrotestosterone at concentrations of 10−6, 10−7, and 10−8 M showed no effect on MMP (data not shown).

We noted that the 585- to 529-nm ratio of the JC-1 experiments varied between assays, sometimes being more than one and other times less than one. All assays showed a progesterone/progestin-induced increase in 585-nm emission, consistent with increased mitochondrial JC-1 aggregates, and a decrease in 529-nm emission, consistent with decreased cytoplasmic monomers, thus indicating increased MMP. The difference in assays with ratios of greater than one vs. less than one depended upon the distribution of JC-1 within the cell. Assays with ratios of greater than one had a greater accumulation of JC-1 in the mitochondria compared with the cytoplasm, whereas assays with ratios less than one had greater accumulation of JC-1 in the cytoplasm compared with the mitochondria. This suggests a difference in baseline mitochondrial activity but does not contradict the finding of increased MMP with progesterone/progestin treatment. We were not able to determine the cause of different baseline mitochondrial activity levels.

To verify that mitochondrial hyperpolarization was associated with increased cellular respiration, total cellular ATP was measured. Figure 4 shows an increase in total cellular ATP after a 2-h treatment with R5020; the increase was inhibited by cotreatment with the PR antagonist RTI-6413-049b. The change in total ATP is not likely because of a difference in cell number, since total cellular protein was unchanged.

The consequence of the progesterone-induced change in cellular respiration upon Fas ligand-induced apoptosis was determined. Figure 5 compares the sub-G1 cell fraction with FACS in cells cotreated with R5020 and FasL-activating antibody after 30 h. A dose-dependent inhibition of FAS-induced apoptosis was demonstrated.

To further corroborate this observation, caspase 3/7 activity was measured after 24 h of cotreatment with R5020 and FasL-activating antibody. Figure 6A shows a dose-dependent inhibition of caspase activity with progestin treatment. Figure 6B shows inhibition of the progestin effect on caspase activation by simultaneous addition of the PR antagonist RTI-6413-049b.

Next, we investigated an effect of R5020 on cell cycle dynamics (Supplemental Fig. 4). With the use of nonsynchronized cells, FACS analysis every 6 h from 6 to 48 h showed no statistically significant difference in the percentage of cells in G1, S, or G2/M phases between control and treated cells. Additionally, cells synchronized by serum starvation were treated by adding back serum with or without R5020 (10−6 M). There was no difference in the percentage of cells in G1, S, or G2/M phases between control and treated cells after 41 h (data not shown).

Last, we sought to provide further evidence of a causal relationship between increased MMP and cell survival using the oxidative phosphorylation substrate pyruvate. Cells treated with 1, 2, and 4 mM sodium pyruvate for 60 min showed an increase in MMP at the 4 mM dose. Subsequently, cells treated for 24 h with 4 mM sodium pyruvate and activating Fas antibody showed decreased caspase 3/7 activity compared with activating Fas antibody alone (Supplemental Fig. 5).

DISCUSSION

We sought to perform experiments in an immortalized cell line that was similar to normal ER/PR (estrogen receptor/progesterone receptor) negative breast epithelial cells. Unlike breast cancer cells, MCF-10A breast epithelial cells show no abnormality of expression of oncogenes, including c-erb-2 and c-HA-ras-1, and no expression of SV40 T-antigen. Likewise, these cells lack anchorage-independent growth and will not survive in an athymic mouse model. Karyotype analysis shows mild aneuploidy of 48, XX, 3p-, 9p+, 6p+, +18, and +16 (45). Previous investigators have identified neither nuclear PR protein in MCF-10A cells by Western blot analysis (34) nor nuclear PR transcript by RT-PCR (8). Our work has shown the same findings, with no evidence of a nuclear PR transcript by RT-PCR or protein by Western blot analysis (unpublished data).

Our studies showed a definite effect of progesterone/progestins on MCF-10A cells despite the absence of a nuclear PR. A rapid dose-dependent increase in MMP was seen with both natural progesterone and a synthetic progestin at physiological doses. This response was inhibited by a specific PR antagonist, RTI 6413-049b. Unlike RU-486, RTI 6413-049b lacks any PR agonist effects, lacks binding of ERα, ERβ, and AR, and has much less antiglucocorticoid effects (41). Lack of inhibition of the response by cycloheximide supports a mechanism not dependent upon protein synthesis. The dose of cycloheximide used in this study was previously shown to inhibit protein synthesis in MCF-10A cells (36). We corroborated this finding by demonstrating inhibition of TGF-β1-induced expression of Id-1, a known transcriptional induction in MCF-10A cells (28).

The question is raised as to whether the progesterone effect could be related to a glucocorticoid action, since GR are found in these cells (27). Evidence against this includes the response to R5020, which has little or no reaction with the GR. Using a glucocorticoid response element transfection assay in 1470.2 cells (a mouse mammary carcinoma cell line), no activation of endogenous GR was seen at a dose of 30 nM R5020 (13). In our studies, an effect on MMP was seen at the lowest dose of 10 nM R5020. Additionally, we observed no increase in MMP after 60 min treatment with dexamethasone with a maximum concentration of 1 μM. A specific GR antagonist, AL082D06, showed no change in the R5020 response. In studies with a hGR expression vector and MMTL:Luciferase reporter, there was no activation of the GR by AL082D06 from concentrations of 10−5 to 10−11 M. A dose of 1 μM AL082D06 completely inhibited GR activation by dexamethasone at 3 × 10−10 M. Last, AL082D06 had no detectable binding to the PR (32). We demonstrated activity of AL082D06 by the antagonism of a known dexamethasone response, namely the inhibition of kalkrien 10 transcription (27).

An increase in MMP may be the result of 1) an increase in proton pumping by increased activity of respiratory enzyme complexes and consequential ATP production, 2) a transient block of electron transfer, 3) a transient block of proton use by enzymes within the mitochondria, and 4) a change in the mitochondrial pH leading to a change in membrane potential (54). Our observation of a concordant increase in total cellular ATP supports the initial mechanism of an increase in respiratory enzyme activity. These studies are inadequate to completely define the mechanism whereby progesterone increases cellular respiration. Possibilities include an increase in glycolysis resulting in increased mitochondrial substrate and/or a direct effect on mitochondrial enzyme activity. Further metabolomic studies will be necessary to delineate these mechanisms.

Next, we determined the consequence of increased mitochondrial activity in the presence of an apoptosis activator, FasL (1). Most commonly, apoptosis is associated with mitochondrial membrane depolarization. FasL classically activates the extrinsic apoptotic pathway. Yet, cross talk exists in which FasL activation of caspase 8 results in cleavage of Bid. Cleaved Bid translocates to the mitochondrion with coordinate depolarization of the MMP (3, 48). As with all cellular energy-requiring processes, ATP is required for early steps of apoptosis such as protein kinase activities and for dynein-mediated movement of mitochondria and other organelles (43). Thus a transient increase in MMP and ATP production may be seen before apoptosis.

These studies show that the progesterone-induced increase in mitochondrial activity is not a precursor to apoptosis, but rather is protective. Inhibition of apoptosis by progestin has been previously demonstrated in breast cancer cells, but always with emphasis on cells expressing nuclear PR. Moore and colleagues (33) showed inhibition of cell death induced by serum starvation or chemotherapeutic agents by 1 nM R5020 after 6 days in T47D cells. Surprisingly, a similar but less robust effect was seen in the MDA-MB-231 cell line, known to lack expression of nuclear PR (20). The authors contributed this to a possible GR effect, but our observations suggest another possibility.

We provide further evidence that an increase in mitochondrial activity may be protective against apoptosis by treating cells with sodium pyruvate. Sodium pyruvate increased MMP and inhibited caspase 3/7 activation due to FasL. Even though pyruvate increases oxidative phosphorylation, the multiple interconnections between metabolic pathways such as glycolysis, gluconeogenesis, and oxidative phosphorylation make it impossible to solely isolate mitochondrial function. Extracellularly added pyruvate has multiple actions, including activation of gylcolysis, alcohol oxidation, and activation of pyruvate dehydrogenase complex, increasing tricarboxylic acid cycle substrates (46).

The energy status of the cell determines the chance of cell death vs. survival. As an example, a transient decrease in ATP concentration triggers apoptosis. A threefold decline is seen in HeLa cells treated for 3 h with oligomycin to block oxidative phosphorylation and deoxyglucose to inhibit glycolysis. Despite reversal of ATP levels with subsequent removal of deoxyglucose, >40% of cells had undergone apoptosis by 48 h (21, 43). In contrast, enhanced ATP production decreases apoptosis. Enhanced glucose uptake with subsequent increased glycolysis decreases apoptosis induced under several experimental conditions, including hypoxia (10, 42), growth factor withdrawal (22), and tumor necrosis factor-α stimulation (2). The contribution of ATP production from oxidative phosphorylation, compared with glycolysis, has been less well defined with conflicting data. In rat neonatal cardiac myocytes, hypoxia-induced apoptosis was inhibited by glucose but not other substrates such as lactate, pyruvate, and propionate (30). Yet, in another study, apoptosis induced in mast cells by growth factor withdrawal was blocked by the oxidative phosphorylation substrates glutamine and pyruvate (22). Differences in the results may lie in different cell types or the different treatments to induce apoptosis. For example, hypoxia results in a decline in activity of oxidative phosphorylation enzymes, which may not be seen with other methods of apoptosis induction (35).

Despite an increase in cellular respiration, we did not see evidence of proliferation or change in cell cycle distribution with progestin treatment. This agrees with many publications showing that progesterone alone does not induce proliferation in MCF-10A cells (9, 25, 51). The energy status of the cell is key in the decision to progress from the G1 to the S phase of the cycle. Inhibiting cellular ATP production by lowering glucose (19) or by inhibiting oxidative phosphorylation (39, 47) results in a block from G1-to-S transition. The ATP levels of the cell are sensed by AMP-activated protein kinase (AMPK). As ATP levels fall, AMP levels rise, activating AMPK with subsequent activation of p53. p53 acts to reduce levels of cyclin E, the rate-limiting protein for G1-to-S transition (31). Our studies show that an increase in cellular respiration alone is not adequate to stimulate proliferation in these cells.

Recently, a synergistic increase in proliferation by a progestin in combination with a tyrosine kinase growth factor was demonstrated in MCF-10A cells (24, 25). The explanation for this observation may involve the progestin-induced enhanced cellular respiration discussed above.

The receptor responsible for mediating a progesterone increase in mitochondrial activity remains to be determined. Our studies support that this is not due to cross-reactivity with the GR, but does not reveal the PR responsible. Also, it is not clear if this is a direct action on the mitochondrion or a secondary increase in enzyme activity because of increased substrate.

Candidate receptors for nongenomic progesterone action remain poorly defined. A G protein-coupled receptor [membrane (m) PR] was originally cloned from the sea trout ovary and was shown to modulate mitogen-activated protein kinase (MAPK) activity when transfected to MDA-MB-231 breast cancer cells lacking nuclear PR expression (53). Three human homologs (mPR α, β and γ) have been identified (52). Enthusiasm for mPR was dampened when another study showed the protein localized to the endoplasmic reticulum, not the plasma membrane, and could not demonstrate specific progestin binding nor MAPK activation (26).

We have previously cloned and expressed a novel, truncated progesterone receptor, named PR-M (38). The predicted protein structure of PR-M from cDNA sequence includes a novel NH2-terminal mitochondrial localization signal followed by the same hinge and hormone-binding domains found in the nuclear PR. Data presented thus far only in abstracts has shown mitochondrial localization by several techniques, including confocal imaging of a green fluorescent protein-tagged recombinant protein, Western blot analysis of purified human heart mitochondrial protein, and Western blot analysis after cellular fractionation of nuclear PR-negative T47D-Y cells (11). In addition, previous studies showed expression of PR-M in MCF-10A cells (12). Further studies are needed to determine the role of PR-M in the control of mitochondrial activity.

PR-C is another NH2-terminal truncated variant with the translation start site proposed to be Met-595 located within the DNA-binding domain (DBD), believed to result in a 60-kDa protein. The predicted protein sequence is characterized by a defective DBD followed by an intact hinge and hormone-binding domain. In vitro expression has shown cytoplasmic localization with ligand-induced nuclear migration (49, 50). Thus far, nongenomic actions of PR-C have not been proposed.

Other PR splice variant transcripts have been identified in various tissues. These include PR-S and PR-T originally cloned from a human testicular cDNA library with transcripts demonstrated in human endometrium (16, 17). The transcripts are characterized by novel noncoding 5′ exons (S, T) derived upstream of exon 1 of the nuclear PR followed by sequence for exons 4–8. To date, publication of data supporting translation into a functional protein is lacking.

Using MCF-10A cells as a model, our studies yield further evidence of a progesterone effect on nuclear PR-negative breast epithelial cells. This challenges the dogma that all effects of progesterone on breast epithelial cells lacking nuclear PR expression are indirect, via paracrine actions from adjacent PR-positive cells. We postulate that progesterone increases cellular respiration in nuclear PR-negative breast epithelial cells. Depending on other physiological factors, progesterone-stimulated ATP production may contribute to prevention of cell death and/or act cooperatively with growth factor-stimulated signaling (35).

GRANTS

This work was supported by National Institute of Child Health and Human Development Grants 1R03HD-052770-01 and the Charles B. Hammond, MD, Foundation to T. M. Price.

ACKNOWLEDGMENTS

We thank Dr. Donald McDonnell for the gift of RTI-6413-049b and Ligand Pharmaceuticals for the gift of AL082D06. We also thank Dr. Ricardo Pietroban, Dimple Rajgor, and the Research on Research Group of Duke University for statistical analysis. We are indebted to Dr. Richard Auten for providing necessary technical assistance for this study.

Current addresses: R. Garde, MD:15001 Shady Grove Reproductive Science Center, 15001 Shady Grove Rd., Ste. 400, Rockville, MD 20850; and E. Jungheim, MD:Washington University, 4921 Parkview Pl., Ste. 6C, St. Louis, MO 63110.

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