Identification of PRG1, A Novel Progestin-Responsive Gene with Sequence Homology to 6-Phosphofructo-2-Kinase/Fructose- 2,6-Bisphosphatase

Jenny A. Hamilton, Michelle J. Callaghan, Robert L. Sutherland and Colin K. W. Watts

Cancer Research Program Garvan Institute of Medical Research St. Vincent’s Hospital Sydney, New South Wales 2010, Australia


    ABSTRACT
 TOP
 ABSTRACT
 INTRODUCTION
 RESULTS
 DISCUSSION
 MATERIALS AND METHODS
 REFERENCES
 
To define early molecular targets of progestin action, the differential display technique was used to identify genes with altered levels of expression in T-47D breast cancer cells treated with the synthetic progestin ORG 2058 for 3 h. PRG1 was first isolated as a 200-bp cDNA clone and its progestin regulation confirmed by Northern analysis. Cloning of the complete coding region of PRG1 revealed that it shared a high degree of amino acid sequence identity with isoforms of the enzyme 6-phosphofructo-2-kinase/fructose-2,6-bisphosphatase from several tissues and species. Expression of PRG1 mRNA was observed in several normal breast epithelial and breast cancer cell lines and in a variety of human tissues, with highest expression in the breast, aorta, and brain. In T-47D cells, PRG1 mRNA was rapidly and transiently induced by progestins, expression peaking between 2 and 4 h and returning to control levels by 12 h. Progestin-induced increases in PRG1 mRNA were inhibited by the progestin antagonist RU 486 and occurred via the progesterone receptor. Progestin induction of PRG1 mRNA was also inhibited by actinomycin D but not by cycloheximide. PRG1 is therefore a novel human gene that is directly regulated by progestins via the progesterone receptor.


    INTRODUCTION
 TOP
 ABSTRACT
 INTRODUCTION
 RESULTS
 DISCUSSION
 MATERIALS AND METHODS
 REFERENCES
 
The sex steroid hormone progesterone has two major roles in mammalian physiology. First, progesterone is involved in preparing the uterus for implantation of the fertilized ovum. Second, progestins have proliferative and differentiating effects on mammary epithelium (reviewed in Refs. 1 and 2). Mitotic activity in breast epithelium varies in a cyclic manner through the menstrual cycle, and a role for progesterone in this process is suggested by observations that levels of this hormone and epithelial cell proliferation are both maximal during the late secretory phase (3). Progesterone is essential for lobuloalveolar development and preparation for lactation: when ovulation is established, progesterone, produced by the corpus luteum, stimulates growth of the lobuloalveolar structures and, during pregnancy, promotes branching of the ductal system and differentiation of alveolar cells into secretory cells ready for milk production. The importance of progestin in these processes is clearly illustrated in progesterone receptor (PR) knockout mice, which fail to develop lobuloalveolar structures (4). Some breast tumors retain progesterone responsiveness, and the use of high doses of synthetic progestins are recognized endocrine therapies for PR-positive breast cancers, since in this setting progestins have an antiproliferative effect (5). Progestins also have predominantly growth-inhibitory effects on human breast cancer cell lines in vitro, although under certain conditions they may stimulate growth (Refs. 2 and 6 and references therein). Mechanistic studies have clearly defined both a stimulatory and inhibitory effect of progestins on breast cancer cell cycle progression (6), but the functional consequences of these effects in vivo remain to be defined. This is of considerable importance given the widespread pharmacological usage of progestins in oral contraceptives and in hormone replacement therapy.

The mechanisms underlying the biological effects of progestins in the normal breast and in breast cancer are only partially understood. Progestin action is mediated primarily via the PR, which upon activation by ligand binding interacts with gene promoter sequences containing progesterone responsive elements (PREs) to regulate gene transcription. Very few mammalian genes have been described that are directly regulated by progestins in this manner; examples include c-jun (7), c-fos (8), fatty acid synthetase (9), PR (10, 11), and uteroglobin (12, 13). Of these c-jun and c-fos have roles in control of cell cycle progression. Other progestin-regulated genes with known roles in cell cycle control are c-myc (6, 8), and cyclin D1 (14). PR can be classified with those progestin-regulated genes whose functions are related to steroid and growth factor action and might contribute to the proliferative effects of progestin, at least indirectly. This group includes estrogen receptor (15), retinoic acid receptors (16), epidermal growth factor receptor (6, 17), PRL receptor (18), insulin-like growth factor I receptor (19), insulin receptor (20), epidermal growth factor (21), transforming growth factors {alpha} and ß1 (6, 8, 22, 23), 17ß-hydroxysteroid dehydrogenase (24), and insulin-like growth factor-binding proteins 4 and 5 (25). Other progestin-regulated genes including fatty acid synthetase (9), alkaline phosphatase (26), and lactate dehydrogenase (27) have functions that are important in differentiation effects mediated by progestin. While progestin action ultimately involves changes in the levels of large numbers of mRNAs and proteins, many of these require intermediary de novo protein synthesis. Specific genes that mediate the proliferative effects of progestins are poorly defined, and thus much remains to be learned about genes induced as an acute response to progestin treatment and their role in mediating progestin effects on cell proliferation and differentiation.

The aim of this study was therefore to identify novel progestin-regulated target genes involved in early responses of progestin action in breast cancer cells. We employed a serum-free cell culture system using the T-47D human breast cancer cell line that enables both stimulatory and inhibitory effects of progestins on cell cycle progression to be observed (6). In this system progestins accelerate entry into S phase of cells already progressing through G1 phase, presumably by acting on genes or gene products that are rate limiting for G1 progression. These cells subsequently complete a round of replication and then become growth-arrested early in G1 phase.

We isolated mRNA after 3 h of treatment with the synthetic progestin ORG 2058 (16{alpha}-ethyl-21-hydroxy-19-norpregn-4-en-3,20-dione) and used this as a template for cDNA synthesis and analysis by the differential display technique (28). This time point was chosen as previous studies showed that final commitment to cell cycle progression does not occur until 3 h after initial progestin exposure (6, 14), implying that critical changes in progestin-regulated gene expression are most likely to be detected within this time frame. Using this strategy we now report the identification and characterization of a novel progestin-regulated cDNA from human breast cancer cells, PRG1 (progestin-responsive sequence, Garvan 1), which shares a high degree of sequence homology with the bifunctional enzyme phosphofructo-2-kinase/fructose-2,6-bisphosphatase1 and may link progestin action with the glycolytic pathway.


    RESULTS
 TOP
 ABSTRACT
 INTRODUCTION
 RESULTS
 DISCUSSION
 MATERIALS AND METHODS
 REFERENCES
 
Cloning of a cDNA Identified by Differential Display
The differential display technique was used to identify mRNAs in T-47D human breast cancer cells whose levels of expression had altered in response to treatment with the synthetic progestin ORG 2058 for 3 h. Using the PCR primer combination 5'-T12GG and 5'-CAAACGTCGG, a total of nine cDNA fragments identified by gel electrophoresis were clearly up-regulated (Fig. 1AGo). Preliminary confirmatory screening by Northern analysis showed one of these, designated PIG1, was induced rapidly in the presence of ORG 2058 (Fig. 1BGo) and therefore warranted further characterization.



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Figure 1. Identification of Differentially Expressed cDNAs in T-47D Cells Treated with the Synthetic Progestin ORG 2058

A, Identification of PIG1 by differential display. Total RNA obtained from T-47D cells treated with ORG 2058 or vehicle control (ethanol) for 3 h was used as a template for differential display PCR reactions with 5'-T12GG and 5'-CAAACGTCGG as primers. The PCR products were separated on a 6% polyacrylamide denaturing gel, and the gel was exposed to x-ray film. The arrows indicate PCR products present at a higher level in the progestin-treated (T) compared with control (C) lane. B, Confirmation of the progestin induction of PIG1 by Northern blot analysis. T-47D cells proliferating in insulin-supplemented serum-free medium were treated with 10 nM ORG 2058 (T) or ethanol vehicle (C) for 3 h, and total RNA was harvested for Northern analysis. The Northern blot was probed with the PIG1 fragment.

 
To obtain the complete coding sequence from which PIG1 was derived, a human kidney cDNA library constructed using oligo-dT-primed and random-primed cDNA was screened using the PIG1 fragment. Four cDNAs were isolated after screening 2.85 x 105 recombinants, namely 11.2, 6.3, 3.1, and 19.1 (Fig. 2AGo). Further screening of this library with an oligonucleotide derived from 5'-sequence of 3.1 (5'-ACCGTCATCGTCATGGTGGG-3') and with a 373-bp EcoRI-BglII restriction fragment derived from 3.1 resulted in the isolation of a chimeric clone 9.1 and clone 8.1 (Fig. 2AGo). Screening of a human heart library with the 373-nt EcoRI-BglII restriction fragment resulted in the isolation of clone H7.1 (Fig. 2AGo). Clones 11.2, 6.3, and 3.1 were sequenced in their entirety on both strands, and 350 nucleotides of the 5'-end of clone 19.1 were sequenced on both strands, to give 2887 nucleotides of cDNA sequence, designated PRG1, shown in uppercase letters (Fig. 2BGo). While the 2887 nucleotides of sequence is less than the mRNA size determined from Northern blots, it contains a complete open reading frame (ORF) as discussed below.



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Figure 2. Determination of the PRG1 cDNA Sequence

A, A schematic representation of PRG1 structure with a restriction map for the PRG1 cDNA and the cDNA clones used to derive the PRG1 sequence shown beneath. The initial PCR cDNA fragment identified by differential display was designated PIG1. All the cDNA clones were isolated from a human kidney cDNA library with the exception of H7.1, which was isolated from a human heart cDNA library. Clone 9.1 is a chimeric clone. The cosmid clone containing genomic sequence, part of which overlaps with PRG1 cDNA sequence, was obtained from the Genbank database and is shown above the PRG1 sequence. The numbers refer to distances in nucleotides. B, Nucleotide and deduced amino acid sequence of PRG1. The nucleotide sequence determined from cDNA clones is shown in uppercase letters whereas the nucleotide sequence obtained from the cosmid genomic clone CRI-JC2015 is shown in lowercase letters. The translation termination codon is shown by an asterisk in the amino acid sequence. The in-frame termination codon that precedes the initiating methionine is underlined. The numbers refer to distances in nucleotides.

 
Comparison of the cDNA sequence with the GenBank and EMBL databases revealed a partial overlap with a cosmid clone (CRI-JC2015) (29) containing human genomic sequence from chromosome 10. Nucleotides numbering 1–399 of the cDNA overlap with nucleotides 1671–2070 from the cosmid clone but in the reverse orientation (Fig. 2AGo). The first 1670 nucleotides of the cosmid clone are not present in PRG1 and appear to be intron sequence as the consensus splicing sequence (30) occurs adjacent to PRG1 homologous sequence. Extrapolating backward 27 nucleotides from the 5'-end of the cDNA into the genomic sequence reveals an in-frame stop codon (see Fig. 2BGo) where the genomic sequence is shown in lowercase letters.

Analysis of the PRG1 cDNA sequence identified an ORF containing 520 amino acids encoding a protein very similar to human liver 6-phosphofructo-2-kinase/fructose-2,6-bisphosphatase (PFK-2/FBPase-2) (31) as well as to bovine brain and heart forms of this enzyme (32, 33). PRG1 bears 72% amino acid identity with the human liver PFK-2/FBPase-2 in 447-amino acid overlap and 93% and 74% identity with bovine brain PFK-2/FBPase-2 and bovine heart PFK-2/FBPase-2 in 462-amino acid and 447-amino acid overlap, respectively (Fig. 3Go). The ORF identified in the PRG1 cDNA sequence appears to be complete. The initiation codon is preceded by an in-frame stop codon located 357 nucleotides upstream as identified in the cosmid clone (CRI-JC2015) and is surrounded by a consensus sequence for strong translational initiation (34). In addition, the initiating methionine is within close proximity to initiating methionines of other known related proteins (e.g. Refs. 31, 33, 35, 36) with the exception of the bovine brain PFK-2/FBPase-2, the cDNA sequence of which has only been partially characterized (32).



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Figure 3. Protein Sequence Homologies of Human and Bovine PFK-2/FBPase-2 Isoforms

Amino acid sequence homology between PRG1 and PFK-2/FBPase-2 isoforms from human liver, bovine brain, and bovine heart. An alignment of the amino acid sequences was obtained using the computer programs MacVector 4.5.3 and SeqVu. Identical residues found in three or four of the proteins are boxed. Amino acids are numbered from the first residue. The bovine brain sequence is believed to be incomplete (32). bov., Bovine; hum., human.

 
Analysis of the deduced amino acid sequence of PRG1 using the Prosite Database Release 13.0 revealed several consensus phosphorylation sites for cAMP/cGMP-dependent protein kinases, protein kinase C, casein kinase II, and tyrosine kinases. Of particular interest was a consensus cAMP-dependent protein kinase site at Ser461 and a consensus protein kinase C site at Tyr471 because in the bovine heart isoform cAMP-dependent phosphorylation at Ser466 and protein kinase C phosphorylation at Tyr476 affects kinase activity (37). In addition, the ATP/GTP-binding site signature motif, conserved in all mammalian forms of PFK-2/FBPase-2 (38), was identified at amino acids 42–49 as was the phosphoglycerate mutase family phosphohistidine signature, amino acids 51–60.

Northern Blot Analysis of PRG1 Gene Expression
The expression profile of PRG1 in a panel of human breast cancer and normal breast cell lines was investigated by hybridizing Northern blots of total RNA isolated from one normal breast epithelial cell strain (HMEC 184), two transformed epithelial cell lines (HMEC 184B5 and HBL-100), and 12 breast cancer cell lines (only nine are shown) to a probe from a 1.8-kb cDNA subclone, 6.3 (Fig. 2AGo), which contains 98% of the PRG1 ORF. PRG1 mRNA, a single transcript of ~4.4 kb, was expressed in all the breast cancer and normal breast epithelial cell lines examined (Fig. 4AGo). The highest level of expression was in the T-47D cell line, and the lowest levels were noted in SK-BR-3 (not shown), BT-474, HBL-100, HMEC 184, and HMEC 184B5 cell lines. There was no correlation between PRG1 mRNA expression and estrogen receptor, PR, or glucocorticoid receptor (GR) status (39) although the T-47D cell line, which expresses PR at a level 5-fold higher than the other cell lines, had the highest level of expression (40).



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Figure 4. Expression of PRG1 mRNA in Different Human Tissues and Breast Cancer and Normal Breast Cell Lines

A, Northern blot analysis of total RNA from different human breast cancer and normal breast cell lines. The blot was probed with a 1.8-kb cDNA subclone of PRG1 and an oligonucleotide complementary to 18S rRNA as a loading control. B, Northern blot analysis of poly A+ RNA from human tissues. The blot was hybridized with a 1.8-kb cDNA subclone of PRG1. Molecular sizes of markers are indicated. PBL, Peripheral blood leukocytes. C, Dot blot analysis of poly A+ RNA from human tissues. The blot was hybridized with a 1.8-kb cDNA subclone of PRG1. Row A: 1, whole brain; 2, amygdala; 3, caudate nucleus; 4, cerebellum; 5, cerebral cortex; 6, frontal lobe; 7, hippocampus; 8, medulla oblongata; Row B: 1, occipital lobe; 2, putamen; 3, substantia nigra; 4, temporal lobe; 5, thalamus; 6, subthalamic nucleus; 7, spinal cord; Row C: 1, heart; 2, aorta; 3, skeletal muscle; 4, colon; 5, bladder; 6, uterus; 7, prostate; 8, stomach; Row D; 1, testis; 2, ovary; 3, pancreas; 4, pituitary gland; 5, adrenal gland; 6, thyroid gland; 7, salivary gland; 8, mammary gland; Row E: 1, kidney; 2, liver; 3, small intestine; 4, spleen; 5, thymus; 6, peripheral leukocyte; 7, lymph node; 8, bone marrow; Row F: 1, appendix; 2, lung; 3, trachea; 4, placenta; Row G: 1, fetal brain; 2, fetal heart; 3, fetal kidney; 4, fetal liver; 5, fetal spleen; 6, fetal thymus; 7, fetal lung.

 
The tissue specificity of PRG1 gene expression was investigated by hybridizing Northern blots of poly A+ RNA isolated from a variety of human tissues to a probe made from the subclone 6.3. A 4.4-kb transcript was detected in all the tissues examined (Fig. 4BGo). The apparent abundance of PRG1 mRNA in skeletal muscle shown in Fig. 4BGo is likely the result of uneven mRNA loading because in a second set of human tissue poly A+ Northern blots, skeletal muscle mRNA levels were similar to those of the kidney. In the heart, a band of 1.35 kb was also detected. This band was more easily detected under lower stringency conditions when it was also found to be present in kidney, pancreas, skeletal muscle, and colon (data not shown) and suggests that the cDNA PRG1 probe was cross-reacting with a related sequence. A third transcript of around 9.5 kb was also detected in skeletal muscle.

Hybridization of a human tissue poly A+ RNA dot blot representing a wider range of tissues and loaded by the manufacturer so as to allow a more accurate determination of relative mRNA abundance (Fig. 4CGo) showed that PRG1 expression was highest in mammary gland, various brain tissues, and in the aorta. Moderate levels of expression were seen in most other tissues, although several had relatively low levels of expression, e.g. liver, colon, and bladder.

PRG1 mRNA Is Induced Transiently by Progestin
To examine in detail the kinetics of progestin induction of PRG1, regulation of PRG1 mRNA expression was investigated in T-47D cells cultured in insulin-supplemented serum-free medium and harvested for mRNA at various time points after ORG 2058 treatment (Fig. 5Go). PRG1 mRNA was detected in cells cultured in the absence of ORG 2058. The induction of PRG1 mRNA by ORG 2058 was an early and transient event. Maximal levels of PRG1 induction, 4.4-fold relative to time-matched control in the experiment shown in Fig. 5Go, were observed at 3 h following treatment. The induction at 3 h was typically between 3 and 4.4-fold relative to time-matched controls. After 6 h mRNA levels had decreased and by 12 h had returned almost to control levels. A more detailed analysis of early time points showed that maximal levels were reached by 2 h and sustained until 4 h (data not shown). The increase in PRG1 preceded increases in the proportion of cells in S phase (data not shown), which typically occur around 10 h in this system (6).



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Figure 5. Regulation of PRG1 mRNA Expression by the Synthetic Progestin ORG 2058

T-47D cells proliferating in insulin-supplemented serum-free medium were treated with 10 nM ORG 2058 (closed circles) or ethanol vehicle (open circles), and total RNA was harvested for Northern analysis. The Northern blot shown in panel A was probed with a 1.8-kb cDNA subclone of PRG1 and with an oligonucleotide complementary to 18S rRNA as a control for loading. Positions of the 28S and 18S ribosomal bands are indicated. Values in panel B were obtained by densitometric analysis of the autoradiograph and are expressed relative to the control at 0 h.

 
Induction of PRG1 mRNA in Breast Cancer Cell Lines Is Mediated via the PR
To determine whether the effects of ORG 2058 on PRG1 expression were likely to be mediated by the PR, we examined the effects of other synthetic progestins and the antiprogestin RU 486 (17ß-hydroxy-11ß-(4-methylaminophenyl)-17{alpha}-(1-propynyl)-estra-4,9-diene-3-one) in a variety of breast cancer cell lines. T-47D cells, growing exponentially in medium containing 5% FCS, were treated in parallel with the synthetic progestins ORG 2058, R5020 (17{alpha}-21-dimethyl-19-norpregn-4,9-diene-3,20-dione), and MPA (17{alpha}-acetoxy-6{alpha}-methyl-4-pregn-4-en-3,20-di-one) at 10 nM and mRNA harvested at 3 h. All three synthetic progestins induced PRG1 mRNA between 2- and 2.5-fold above control levels (data not shown). PRG1 mRNA was also induced by the synthetic progestin livial [ORG OD-14, (7{alpha},17{alpha})-17-hydroxy-7-methyl-19-norpregn-5(10)-en-20-yn-3-one, 10 nM] in T-47D cells growing in the presence of 5% charcoal-treated FCS as discussed later. The effect of ORG 2058 on PRG1 mRNA in another PR-positive cell line (MCF-7) and in a PR-negative cell line (MDA-MB-231) were investigated. ORG 2058 increased PRG1 mRNA in MCF-7 cells approximately 2-fold above control mRNA levels at 3 h (Fig. 6AGo). In contrast, in MDA-MB-231 cells the levels of PRG1 mRNA in the presence of ORG 2058 were decreased approximately 30% below control mRNA levels at 3 h, with recovery at 6 h (Fig. 6BGo).



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Figure 6. Regulation of PRG1 mRNA Expression in MCF-7 and MDA-MB-231 Cells by ORG 2058 and Dexamethasone

A, MCF-7 (PR +ve, GR +ve) and B, MDA-MB-231 (PR -ve, GR +ve) cells proliferating in RPMI 1640 medium supplemented with 5% FCS were treated with ORG 2058 (10 nM), dexamethasone (100 nM), or ethanol vehicle and harvested for Northern analysis at 3 h and 6 h. Densitometric analysis of the Northern blot hybridized with a 1.8-kb cDNA subclone of PRG1 is presented expressed relative to control at 0 h and adjusted for loading. The data in panel B were obtained from the mean of two experiments. C, MDA-MB-231 cells were transiently transfected with pMSG-CAT and treated with dexamethasone (100 nM) ± RU 486 (100 nM), and ORG 2058 (10 nM) ± RU 486 (100 nM), or ethanol vehicle (control) for 48 h before harvesting and determination of CAT activity. Results are expressed as fold-induction over control levels of CAT activity and are the mean of triplicate determinations.

 
Figure 6Go also demonstrates another important aspect of PRG1 regulation. Given that the consensus sequence for the glucocorticoid and progesterone response elements is similar (41), one might expect that glucocorticoids could regulate PRG1 expression via the GR. The GR could also have a role in mediating progestin effects, given the receptor cross-reactivity of some progestins for both PR and GR. The MCF-7 and T-47D cell lines express GR, although at low levels (42), and treatment with the synthetic glucocorticoid dexamethasone (9-fluro-11,17,21-trihydroxy-16-methylpregn-1,4-diene-3,20-dione, 100 nM) for either 3 or 6 h produced no increase in PRG1 mRNA (Fig. 6AGo and data not shown). Using these lines we have previously found that dexamethasone is also unable to induce either expression of epidermal growth factor receptor (a glucocorticoid-responsive gene in breast cancer cell lines) or chloramphenicol acetyltransferase (CAT) activity from a transiently transfected reporter construct, pMSG-CAT (which contains glucocorticoid- and progestin-responsive sequences from the MMTV promoter) (42, 43). These results, indicating an apparent absence of functional GR in these cell lines, argue against a GR-mediated mechanism for progestin regulation of PRG1. To establish whether the GR can regulate PRG1 expression in a glucocorticoid-responsive cell line, these experiments were repeated in MDA-MB-231 cells, which contain high levels of GR but no PR (42). In these cells, dexamethasone reduced PRG1 mRNA to approximately 60% of control levels at 3 h with some recovery evident at 6 h, suggesting a small glucocorticoid effect, but opposite to that of progestins in PR-positive cell lines. GR functionality in these cells was confirmed by transient transfection with pMSG-CAT (Fig. 6CGo), which showed 4- to 5-fold induction of CAT activity after treatment with dexamethasone that could be inhibited by the glucocorticoid/progestin antagonist RU 486. ORG 2058 (10 nM) did not induce CAT activity, indicating that this progestin does not cross-react with the GR at this concentration and that the small ORG 2058 effect on PRG1 expression in these cells is unlikely to be mediated via the GR.

Additional evidence for the involvement of PR in mediating progestin effects on PRG1 was obtained using the antagonist RU 486, which acts as an antiprogestin by competitively inhibiting the binding of progestins to the PR (44). Its effect on the induction of PRG1 mRNA was investigated by treatment of T-47D cells with ORG 2058 (10 nM) and RU 486 (100 nM) either alone or simultaneously in serum-free medium supplemented with insulin. The cells were harvested 3 h after the treatment, when PRG1 mRNA levels were at a maximum. Simultaneous administration of ORG 2058 and RU 486 led to complete inhibition of progestin-induced PRG1 expression, while treatment with RU 486 alone had no effect on mRNA levels (Fig. 7AGo). Similar effects were seen in MCF-7 cells grown in the presence of serum although the antagonist effect of RU 486 was not quite as pronounced (Fig. 7BGo). Together, these data are consistent with the progestin effect being mediated via the PR and not the GR.



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Figure 7. Antagonism of ORG 2058 Induction of PRG1 mRNA by the Antiprogestin RU 486

A, T-47D cells proliferating in insulin-supplemented serum-free medium were treated with ORG 2058 (10 nM), RU 486 (100 nM), the two compounds simultaneously (ORG 2058 + RU 486), or ethanol vehicle and harvested for Northern analysis at 3 h. The Northern blot was probed with the PIG1 fragment of PRG1 and with an oligonucleotide complementary to 18S rRNA as a control for loading. The graph represents densitometric analysis of the autoradiograph expressed relative to the control at 3 h. B, MCF-7 cells proliferating in medium supplemented with 5% FCS were treated with ORG 2058 (10 nM), ORG 2058 + RU 486 (100 nM) simultaneously, or ethanol vehicle and harvested for Northern analysis at 3 h. The Northern blot was probed with a 1.8-kb cDNA subclone of PRG1 and with an oligonucleotide complementary to 18S rRNA as a control for loading.

 
Because progestin effects can potentially be mediated by the androgen receptor (AR) present in breast cancer cell lines (39, 45), the effect of the antiandrogen hydroxyflutamide ({alpha},{alpha},{alpha}-trifluoro-2-methyl-4'-nitro-m-lactotoluidide, 1 µM) on ORG 2058 (10 nM) induction of PRG1 was examined. No antagonism was observed (data not shown), an indication that progestins do not mediate their effects on PRG1 induction via the AR. In addition, 5{alpha}-dihydrotestosterone (1–100 nM), with or without a 100-fold molar excess of hydroxyflutamide, did not induce PRG1 above basal levels, further evidence that this gene is not AR-regulated in breast cancer cells (data not shown).

PRG1 Induction by Progestin Does Not Require de Novo Protein Synthesis
To distinguish between direct activation of PRG1 transcription by the PR or indirect activation via the synthesis of intermediary proteins, T-47D cells were treated with ORG 2058 (10 nM) in the presence of the protein synthesis inhibitor, cycloheximide (20 µg/ml). Cycloheximide failed to block the progestin-mediated induction of PRG1 mRNA. Treatment with cycloheximide alone resulted in an increase of PRG1 mRNA to a level similar to that achieved by ORG 2058 alone, while in the presence of both ORG 2058 and cycloheximide there was ’superinduction’ of PRG1 mRNA (Fig. 8AGo). The magnitude of this induction, 12-fold in the experiment shown in Fig. 8Go, was larger than expected from a combination of the responses to the individual compounds. Such superinduction involving protein synthesis inhibitors is characteristic of genes such as ß- and {gamma}-actin, c-fos, and c-myc, which are induced early (0–2 h) after stimulation of cells by mitogens (46, 47, 48). At time points up to 6 h, the transcription inhibitor actinomycin D (5 µg/ml) prevented induction of PRG1 mRNA in T-47D cells treated with the synthetic progestins livial (Fig. 8BGo) or ORG 2058 (Fig. 8CGo). Together, these data suggest that the induction of PRG1 mRNA is due to a direct effect of the PR on PRG1 transcription and does not require de novo protein synthesis.



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Figure 8. Effect of Cycloheximide and Actinomycin D on Progestin Induction of PRG1 mRNA

A, T-47D cells proliferating in insulin-supplemented serum-free medium were treated with ORG 2058 (10 nM), cycloheximide (CHX, 20 µg/ml), ORG 2058, and CHX simultaneously or ethanol vehicle and harvested for Northern analysis at 3 h. The Northern blot was probed with the PIG1 fragment of PRG1 and with an oligonucleotide complementary to 18S rRNA as a loading control. The graph represents densitometric analysis of the autoradiograph expressed relative to the control at 3 h. B, T-47D cells proliferating in medium supple mented with 5% charcoal-treated FCS were treated with livial (10 nM) and ethanol vehicle in the presence and absence of actinomycin D (5 µg/ml) and harvested for Northern analysis. The Northern blot was probed with a 1.8-kb cDNA subclone of PRG1. C, Ethanol control; L, livial. C, T-47D cells were treated as in panel B above, except that ORG 2058 was substituted for livial. C, Ethanol control.

 

    DISCUSSION
 TOP
 ABSTRACT
 INTRODUCTION
 RESULTS
 DISCUSSION
 MATERIALS AND METHODS
 REFERENCES
 
Despite the biological importance of progestins, the patterns of gene expression mediating proliferative and other responses to these steroids are not understood in detail. To define early molecular targets of progestin action, the differential display technique was used to identify genes with altered levels of expression in T-47D human breast cancer cells treated with the synthetic progestin ORG 2058 for 3 h. A cDNA fragment, PIG1, cloned using this method, was used to screen Northern blots and identified a 4.4-kb mRNA species that was induced in response to a variety of synthetic progestins within 3 h. Compilation of DNA sequence data from several cDNA clones generated a 2.8-kb cDNA, PRG1. The ORF of PRG1 contains 520 amino acids and encodes a protein sharing a high degree of identity with 6-phosphofructo-2-kinase/fructose-2,6-bisphosphatase (PFK-2/FBPase-2) isoforms from many species, including that from human liver. The human liver isoform of PFK-2/FBPase-2 (PFKFB1) has been assigned to chromosome X (49) and on this basis, and because of significant differences throughout the nucleotide and amino acid sequences, is clearly distinct from PRG1, the sequence of which partially overlaps with the cosmid clone, CRI-JC2015, derived from a genomic sequence from chromosome 10 (29). A locus for an additional, uncharacterized human gene that may correspond to the rat heart isoform (PFKFB2) has been identified on chromosome 1 (49).

Six isoforms of PFK-2/FBPase-2 have been identified in mammalian tissues. They are referred to as F (fetal)-, L (liver)-, M (muscle)-, H (heart)-, T (testis)-, and B (brain)-type. Although multiple genes exist in other species, e.g. the bovine H- and B-isoforms, only one human gene, the L-isoform, has been cloned and functionally characterized. The primary differences between mammalian isoforms of this enzyme are the length of the amino- and carboxy-terminal regions and the composition of these regions particularly with regard to the number of protein phosphorylation sites (reviewed in Ref.38). Phosphorylation reciprocally affects the enzyme’s kinase and bisphosphatase activity. Although the ORF of PRG1 shares the highest degree of identity with the bovine brain PFK-2/FBPase-2, its expression is not restricted to neural tissue but is detectable in all tissues examined, although at a wide range of levels. In addition, the 3'-end of the PRG1 ORF sequence has greater similarity with bovine heart PFK-2/FBPase-2 than with the bovine brain isoform. On the basis of the above evidence, it is likely that PRG1 represents the third known human gene of the PFK-2/FBPase-2 family. A comparison of their respective tissue distributions suggests PRG1 has a much more widespread role than the L-isoform of PFK-2/FBPase-2 whose expression among the tissues shown in Fig. 4BGo is limited to liver and skeletal muscle. It is also apparent that despite high levels in the mammary gland and progestin-responsive breast cancer cells, PRG1 expression is not confined to progestin target tissues, where, in fact, expression can be low, e.g. in uterus. Functional analysis of PRG1 protein will be necessary to determine whether the proteins produced in the breast, breast tumors, and other tissues have a similar function and whether PRG1 is the homolog of isoforms identified in other species or has unique properties.

The progestin regulation of PRG1 mRNA was studied in some detail, and Northern analysis of PRG1 mRNA levels over a 24-h time period showed a rapid and transient induction by ORG 2058 that peaked at 3 h and returned to control levels by 12 h. Several lines of evidence are consistent with the view that this response is mediated by the PR. First, the induction of PRG1 mRNA occurred in the presence of three other synthetic progestins, R5020, MPA, and livial. Second, although these progestins potentially have some cross-reactivity with the GR (50, 51), the progestin induction does not appear to be mediated by this receptor in T-47D or MCF-7 cells as the GR is nonfunctional in these lines. There was also no evidence for the involvement of AR in mediating progestin induction. Third, the progestin antagonist RU 486 inhibited the progestin induction of PRG1 mRNA. In MDA-MB-231 cells expressing high levels of functional GR (39), PRG1 was not strongly glucocorticoid-regulated, i.e. dexamethasone caused only a moderate decrease in PRG1 mRNA levels. Interestingly, ORG 2058 had a similar but weaker effect in this cell line. The mechanism by which this occurs is not known but does not appear to involve cross-reactivity of ORG 2058 with the GR.

The progestin induction of PRG1 mRNA was not prevented by the presence of the protein synthesis inhibitor cycloheximide but was blocked by the transcription inhibitor actinomycin D. This strongly suggests that induction of PRG1 by progestin is a transcriptional effect of ligand-activated PR on the PRG1 gene. A computer search of DNA sequence from the cosmid clone CRI-JC2015, encompassing 3000 bp from the initiating methionine in a 5' direction, for an optimal PRE sequence as determined by in vitro studies (52), revealed several PRE-like sequences. The PR could be acting by classic mechanisms on these sequences, i.e. by direct binding, or indirectly, for example through protein-protein interactions with other transcription factors. Identification of the precise mechanism involved in progestin regulation of this gene will require cloning of the PRG1 promoter and characterization of the putative PREs or other DNA-binding motifs responsible for progestin induction. This will enable comparisons to be made with recognized PREs (52, 53, 54, 55) and provide further insight into the nature of these elements.

PFK-2/FBPase-2 is a bifunctional enzyme that catalyzes the synthesis and degradation of fructose-2,6-bisphosphate, a molecule that is a potent stimulator of 6-phosphofructo-1-kinase (PFK-1) and therefore has a role in control of glycolysis (reviewed in Ref.56). The tentative assignment of a functional role for PRG1 as a PFK-2/FBPase-2 isoform suggests a link between progestin effects and glycolytic control. Because PRG1 induction by progestins precedes increases in the S phase fraction of cell populations, it is possible that PRG1 has a role in glycolytic control during the initiation of cell cycle progression. There is some evidence of such a role for PFK-2/FBPase-2 under these conditions. An important metabolic response to growth stimulation by mitogens is an increase in glucose utilization, suggesting that glycolytic rates are regulated to coordinate with DNA synthesis (e.g. Refs. 57–60). Furthermore the rat F-type mRNA of PFK-2/FBPase-2 is induced around the time of the G1/S transition in Rat-1 fibroblasts after epidermal growth factor or serum stimulation of quiescent cells (61).

In summary, this study reports the identification and cloning of PRG1, a new human gene with strong sequence homology to PFK-2/FBPase-2, thus identifying a third human gene for this enzyme. PRG1 is transcriptionally regulated by progestins, with induction peaking at 3 h. This induction is not prevented by cycloheximide, and therefore PRG1 is one of the few human genes demonstrated to be directly regulated by the PR. This gene may prove to be important in advancing our understanding of progestin effects on cell growth and differentiation and gene transcription. Studies are currently underway to investigate the functional activity of the protein product of PRG1 and its role in progestin action in human breast cancer cell lines.


    MATERIALS AND METHODS
 TOP
 ABSTRACT
 INTRODUCTION
 RESULTS
 DISCUSSION
 MATERIALS AND METHODS
 REFERENCES
 
Reagents
Steroids and growth factors were obtained from the following sources: ORG 2058, Amersham Australia (Castle Hill, Australia); R5020, Du Pont Ltd, North Ryde, Australia; MPA, Dr. Dudley Jacobs of Upjohn Pty Ltd, Sydney, Australia; livial, Dr. Willem Schoonen of Organon International (Oss, The Netherlands); RU 486, Dr. John-Pierre Raynaud of Roussel-Uclaf (Romainville, France); 5{alpha}-dihydrotestosterone (5{alpha}-androstan-17ß-ol-3-one) and dexamethasone, Sigma Chemical Co. (St. Louis, MO); hydroxyflutamide, SCH16423, Schering Corp. (Kenilworth, NJ); human transferrin, Sigma Chemical Co.; human insulin, Actrapid, CSL-Novo, North Rocks, Australia. Steroids were stored at -20 C as 1000-fold-concentrated stock solutions in absolute ethanol. Cycloheximide (Calbiochem-Behring Corp., La Jolla, CA) was dissolved at 20 mg/ml in water and filter sterilized. Actinomycin D (Cosmegen, Merck Sharp and Dohme Research Pharmaceuticals, Rahway, NJ) was dissolved at 0.5 mg/ml in sterile water and used immediately. Tissue culture reagents were purchased from standard sources.

Cell Culture
The sources and maintenance of the human breast cancer cell lines used in this study were as described previously (62). 184 and 184B5 normal breast epithelial cells were the kind gift of Dr. M. Stampfer (University of California, Berkeley, CA) and were maintained in mammary epithelial growth medium (Clonetics, San Diego, CA). Tissue culture experiments in serum-free medium were performed as previously described (6, 14). Briefly, T-47D cells were taken from stock cultures and passaged for 6 days in phenol red-free RPMI medium supplemented with 10% charcoal-treated FCS (6, 14). During this time the cells received two changes of medium at 1- to 3-day intervals. The cells were replated into replicate 150-cm2 flasks in medium containing 10% charcoal-treated FCS, and the medium was replaced with serum-free medium on the next 2 days. Serum-free medium was supplemented with 300 nM human transferrin. In experiments involving ORG 2058, the final serum-free medium contained 10 µg/ml (1.7 µM) human insulin. Three days after completion of these pretreatments, steroid, steroid antagonist, or cycloheximide were added. Control flasks received vehicle to the same final concentration. Cell cycle phase distribution was determined by analytical DNA flow cytometry, as previously described (6, 63). Tissue culture experiments in serum-containing medium were performed as previously described (63). The experiments with livial or ORG 2058 and actinomycin D were as for experiments in serum-containing medium except that the medium contained 5% charcoal-treated FCS. Actinomycin D was added at the same time as livial or ORG 2058.

RNA Isolation and Northern Analysis
Cells harvested from triplicate 150-cm2 flasks were pooled and RNA extracted by a guanidinium-isothiocyanate-cesium chloride procedure, and Northern analysis was performed as previously described with 20 µg total RNA per lane (6, 11) The membranes were hybridized overnight (50 C) with probes labeled with [{alpha}-32P]dCTP (Amersham Australia Pty Ltd, Castle Hill, Australia) using the Random Prime Labeling Kit (Promega, Sydney, Australia). The membranes were washed at a highest stringency of 0.2 x SSC (30 mM NaCl, 3 mM sodium citrate [pH 7.0]) + 1% SDS at 65 C and exposed to Kodak X-OMAT or BIOMAX film at -70 C. Human multiple tissue Northern blots and Master dot blots (Clontech Laboratories Inc, Palo Alto, CA) were hybridized under conditions recommended by the manufacturer. The mRNA abundance was quantitated by densitometric analysis of autoradiographs using Molecular Dynamics Densitometer and software (Molecular Dynamics, Sunnyvale, CA). The accuracy of loading was estimated by hybridizing membranes with a [{gamma}-32P]ATP end-labeled oligonucleotide complementary to 18S ribosomal RNA (rRNA) (39, 64).

Transient Transfection with pMSG-CAT
Transfections of MDA-MB-231 cells with pMSG-CAT (AMRAD Pharmacia Biotech, Melbourne, Australia) were carried out as previously described (43) with 40 µg plasmid per 150-cm2 flask without glycerol shock. Cells were treated with dexamethasone (100 nM) or ORG 2058 (10 nM) with or without RU 486 (100 nM) for 48 h before harvesting for CAT assays (43).

Differential Display
Differential display was carried out as described (65). Total RNA, 200 ng, obtained from T-47D cells treated with the synthetic progestin ORG 2058 for 3 h or from T-47D cells treated with ethanol control was reverse transcribed with 5'-T12GG as the primer. The cDNA products were amplified by the PCR using 5'-T12GG and 5'-CAAACGTCGG primers. The PCR products were separated on a 6% polyacrylamide denaturing sequencing gel. The PCR product of interest was excised from the gel, reamplified by PCR, and cloned into the pGEM-T vector (Promega, Madison, WI). DNA sequencing was performed by the dideoxy chain termination method using T7 DNA polymerase (AMRAD Pharmacia Biotech) and Sequenase 2.0 kit (Bresatec, Adelaide, Australia) or by cycle sequencing using the fmol® DNA Cycle Sequencing System (Promega). Sequence database searches were performed at the National Center for Biotechnology Information (NCBI) using the Basic Local Alignment Search Tool (BLAST) network service.

Library Screening
To obtain additional cDNA sequence, {lambda} cDNA libraries derived from human kidney (Clontech; 2.85 x 105 pfu) and human heart (Stratagene, La Jolla, CA; 6 x 105 pfu) were screened using the 32P-labeled differential display cDNA fragment as a probe under stringent hybridization conditions. Seven strongly hybridizing clones were isolated and excised for sequencing using bacterial strain XL1-Blue. Sequencing was performed as described above. Amino acid sequence alignments were performed using the computer programs MacVector 4.5.3 (Eastman Kodak Co., Rochester, NY) and SeqVu (Garvan Institute of Medical Research, Sydney, Australia).


    ACKNOWLEDGMENTS
 
The authors would like to acknowledge members of the Cancer Research Program for assistance with these studies including Andrea Brady, Chris Lee, Kimberley Sweeney, Fiona Hammond, Amanda Russell, and members of the Neurobiology Research Program, Yvonne Hort, Herbert Herzog, and John Shine, for helpful discussions.


    FOOTNOTES
 
Address requests for reprints to: Colin K. W. Watts, Garvan Institute of Medical Research, St. Vincent’s Hospital, Sydney, New South Wales 2010, Australia.

This work was supported by the National Health and Medical Research Council of Australia (NHMRC) and the Kathleen Cuningham Foundation for Breast Cancer Research.

1 The nucleotide sequence data reported in this paper appears in the GSDB and NCBI nucleotide databases with the accession numbers GSDB:S:75650 and L77662. PRG1 has been assigned the gene symbol PFKFB3. Back

Received for publication October 1, 1996. Revision received December 30, 1996. Accepted for publication January 15, 1997.


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