©1995 by The American Society for Biochemistry and Molecular Biology, Inc.
An Initiator Element Is Required for Maximal Human Chorionic Somatomammotropin Gene Promoter and Enhancer Function (*)

(Received for publication, May 10, 1994; and in revised form, October 7, 1994)

Shi-Wen Jiang Allan R. Shepard Norman L. Eberhardt (§)

From the Endocrine Research Unit, Departments of Medicine and Biochemistry/Molecular Biology, Mayo Clinic, Rochester, Minnesota 55905

ABSTRACT
INTRODUCTION
MATERIALS AND METHODS
RESULTS
DISCUSSION
FOOTNOTES
ACKNOWLEDGEMENTS
REFERENCES

ABSTRACT

Previous studies have indicated that cell-specific expression of the human chorionic somatomammotropin (hCS) gene may be mediated by a placental-specific enhancer (CSEn). In the current studies, we have analyzed the promoter elements that are required for enhancer and promoter function in choriocarcinoma cells (BeWo). Mutation of both hCS GHF1 sites had no effect on promoter or enhancer activity. In contrast, mutation of the Sp1 site diminished basal and CSEn-stimulated transcription by approx75% and approx56%, respectively, indicating that Sp1 was necessary but not sufficient for maximal basal and enhancer-mediated transcription. Deletion and site-specific mutation of the proximal promoter region indicated that the TATA box and an initiator site (InrE) located between nucleotides -15/+1 of the hCS promoter were required for maximal promoter and enhancer function. Mutations of the InrE were associated with reduced basal and enhancer-stimulated activities and altered transcription initiation sites. A protein of 70-kDa mass, that was preferentially expressed in human choriocarcinoma cells (BeWo and JEG-3), bound specifically to the InrE. The data suggest that an initiator present in high concentrations in placental cells contributes to the control of cell-specific hCS gene expression at the promoter level and is required for maximal enhancer function.


INTRODUCTION

Human chorionic somatomammotropin (hCS (^1)or placental lactogen, hPL) belongs to the human growth hormone (hGH) gene family which includes prolactin (hPRL) (Miller and Eberhardt, 1983). Both hPRL and hGH are synthesized in the anterior pituitary by lactotrophes and somatotrophes, respectively, whereas hCS is expressed in human placental syncytiotrophoblasts. GH is an essential hormone involved in normal growth, development, and homeostatic regulation. Both hPRL and hCS influence mammary gland differentiation and lactation in pregnancy (Handwerger, 1991). CS has been postulated to be a maternal/fetal growth hormone and may increase fetal glucose availability via its lipolytic/insulin antagonist activity (Handwerger, 1991). Lowered hCS production has been associated with abnormal pregnancy in some but not all studies (Borody and Carlton, 1981; Nielsen et al., 1979). Recent studies indicate that placental lactogens from rat, mouse and human regulate their homologous pancreatic islets directly, suggesting that these hormones are the primary mediators of increased islet beta cell function observed in normal pregnancy and may be involved in gestational diabetes (Brelje et al., 1993). A CS receptor which binds CS, but not GH or PRL, with high affinity has been isolated from a variety of fetal and maternal tissues, suggesting that CS mediates direct and selective functions, including fetal growth promotion (Freemark et al., 1993; Freemark and Comer, 1989).

Although the exonic, intronic and flanking regions of the hCS and hGH genes are highly homologous (geq93.5% sequence identity) due to gene duplication and gene conversion events (Hirt et al., 1987; Miller and Eberhardt, 1983), the mechanisms controlling their cell-specific expression are quite different. Pituitary hGH gene expression is directed by the cell-specific transcription factor GHF1/Pit1 that binds to the proximal hGH promoter (Bodner and Karin, 1987; Ingraham et al., 1988). Conservation of the GHF1 bindign sites in the hCS promoter allows efficient expression of this gene in pituitary (GC) cells (Nachtigal et al., 1989), indicating that complex control of these genes is required to maintain cell-specific expression. In the case of the hCS gene, a placental-specific enhancer, designated CSEn, located approx3 kb down-stream of the hCS-2 gene appears to be responsible for its expression in placenta (Fitzpatrick et al., 1990; Jacquemin et al., 1994; Jiang and Eberhardt, 1994; Walker et al., 1990). CSEn is a classical enhancer that functions in a distance- and orientation-independent manner (Jiang and Eberhardt, 1994). Its activity is limited to cells of placental origin (BeWo and JEG-3 cells) with the exception of a minor activity in pituitary 18-54,SF cells (Rogers et al., 1986). The enhancer is inactive in HeLa, pituitary GC and HepG2 cells (Fitzpatrick et al., 1990; Jiang and Eberhardt, 1994; Walker et al., 1990). We and others have shown that CSEn is composed of multiple DNA elements that interact cooperatively and are homologous to the SV40 GT-IIC and SphI/SphII enhansons (Jacquemin et al., 1994; Jiang and Eberhardt, 1994). The latter data suggest that CSEn function may be controlled by TEF-1 (Xiao et al., 1991), raising the question of how cell specificity is achieved. CSEn appears to function in a promoter-independent manner and functions equally well with the hGH, and herpes simplex thymidine kinase promoters and somewhat less efficiently with the Rous sarcoma virus (RSV) minimal promoter (Jiang and Eberhardt, 1994), suggesting that CSEn can cooperate with different sets of promoter elements or interacts with common basal transcription factors.

In the current studies we have examined the promoter elements present on the hCS 5`-flanking region (5`-FR) that interact with the enhancer. Previous studies indicated that Sp1 played a major role in hCS promoter basal and enhancer-stimulated activities (Fitzpatrick et al., 1990). Our studies confirmed the importance of Sp1 in mediating basal promoter activity and indicate that Sp1 is a necessary but not sufficient factor for maintaining maximal enhancer activity. Mutagenesis studies indicate that proximal elements, including the TATA box and an initiator element (InrE) located between nt -15/+1 of the hCS promoter, act cooperatively with Sp1 to achieve full basal and enhancer activity. Gel shift and UV cross-linking experiments indicate that a 70-kDa nuclear factor that is preferentially expressed in choriocarcinoma cells (BeWo and JEG-3), but is either absent or expressed at low levels in a variety of other cells, binds specifically to the InrE. Mutations on the InrE sequences not only result in major reductions of basal- and enhancer-stimulated activity, but also alter transcription start site selection. The data suggest that cell-specific expression of the hCS gene is mediated in part by an initiator that is preferentially expressed in placental cells.


MATERIALS AND METHODS

Cell Culture

BeWo, JEG-3, COS, and HeLa cells were purchased from American Type Culture Collection. SV40 transformed human fibroblasts (GM0637E) and osteosarcoma cells (MG63) were provided by Dr. Cheryl Conover (Mayo Clinic). HeLa, COS, MG63, GM0637E, JEG-3, and rat anterior pituitary tumor (GC) cells were maintained in Dulbecco's modified Eagle's medium (Life Technologies, Inc.); BeWo cells were grown in RPMI 1640 (Life Technologies, Inc.). All media were supplemented with 10% fetal bovine serum (Whittaker), 100 units/ml penicillin (Life Technologies, Inc.), 100 µg/ml (Life Technologies, Inc.), and 2 mML-glutamine (Life Technologies, Inc.). Cells were maintained at 37 °C in an atmosphere containing 5% CO(2) and 100% humidity.

Cell Transfection and Luciferase Assays

Cells grown to confluence in T175 flasks were rinsed with 10 ml of PBS (phosphate-buffered saline, Life Technologies, Inc.) and harvested by treatment with 5 ml 1 times trypsin (Life Technologies, Inc.). Cells were transfected by electroporation and plated as described previously (Jiang and Eberhardt, 1994). After 18-20 h of incubation, cells were washed twice with 5 ml of cold PBS. Cells were scraped off carefully in 1.5 ml of cold PBS. After centrifugation at 5,000 rpm for 1 min, the PBS was aspirated and the cells were resuspended in 200 µl of lysis buffer (100 mM K(2)HPO(4) (pH 7.8), 1 mM DTT). Lysis was accomplished by three cycles of freezing on dry ice and thawing at 37 °C in a water bath. Following centrifugation at 13,000 rpm for 20 s, the supernatants were saved for luciferase and protein assays as described previously (Jiang and Eberhardt, 1994). Data are expressed as relative light units/µg of protein.

Data Analysis

All data groups were analyzed by analysis of variance (ANOVA) to determine if the effect being monitored, e.g. promoter mutation, among the data was significant at the p < 0.05 level. The results of the ANOVA are provided in the figure legends. For all experimental groups which satisfied the initial ANOVA criterion, individual comparisons were performed against the control using Student's t tests with the use of a Bonferroni inequality to compensate for error introduced due to the multiple comparison analyses (Snedecor and Cochran, 1980). The controls for the comparisons are designated in the figure legends. In cases where the differences in means were very large and the variances were not equivalent, the logarithmical transformed (log) data were analyzed.

Plasmid Constructions

The plasmids pA(3)LUC (Maxwell et al., 1989) and EnApA(3)LUC, containing CSEn, have been described previously (Jiang and Eberhardt, 1994). Site specific deletions and mutations of proximal (p) and distal (d) GHF1 and Sp1 elements were created by inverted PCR mutagenesis (Hemsley et al., 1989; Shepard et al., 1994) with the hCS 5`-FR cloned into the SalI/BamHI sites of pUC8 as template. Oligonucleotides used for the PCR reactions are shown in Table 1. Mutant PCR products were flush-ended with Klenow, 5`-end phosphorylated with T4 polynucleotide kinase, ligated and transformed into Escherichia coli HB101. Individual colonies were screened for correct plasmid mutations by restriction digestion and dideoxy chain termination sequencing (U. S. Biochemical Corp.). The pdGHF1 mutant was made by digesting the dGHF1 mutant with NsiI, treating with T4 polymerase in the presence of dNTPs and religating the modified plasmid. To transfer these mutations to the luciferase vector, the SalI/MscI region of wild-type EnAhCSp.LUC was substituted by the corresponding hCS 5`-FR fragments removed from pUC18 using the same restriction enzymes.



The truncated promoters were generated by digestion of EnAhCSp.LUC with SalI/PstI and SalI/MscI followed by T4 Polymerase treatment and religation to yield EnA282CSp.LUC and EnA36CSp.LUC, respectively. For the EnA19CSp.LUC construct, EnAhCSp.LUC was digested with ApaI, treated with T4 polymerase in the presence of dNTPs, and finally digested with BglI and BstXI. The ApaI (blunted)/BstXI fragment containing the proximal 19 bp of hCS 5`-FR and the entire LUC gene was subcloned into SmaI/BstXI digested pA(3)LUC, generating 19CSp.LUC. The CSEn fragment was inserted at the BglII site to generate EnA19CSp.LUC. For the construction of 493CSDeltaAPAp.LUC, 493CSp.LUC was digested by ApaI and the ends were flushed with T4 polymerase treatment. SalI linkers were added and the hCS 5`-FR fragment was isolated and ligated into the SalI site of EnApA(3).LUC. SalI/MscI digestion and religation of 493CSDeltaAPAp.LUC gave the plasmid containing a TATA box with substitution of the sequences 3` to the TATA box, designated 36CSDeltaAPAp.LUC. In 493CSDeltaTATAp.LUC, a 19-bp region including the TATA box was removed from the hCS 5`-FR by substituting the MscI/BstXI region of EnAhCSp.LUC with the ApaI (blunt)/BstXI fragment of the same plasmid.

Site-specific mutagenesis of the sequences downstream from the TATA box were created by inverted PCR mutagenesis utilizing the primers listed in Table 1. To improve the efficiency of PCR, a small plasmid, pEB.LUC, which contains 493 bp of the hCS 5`-FR and a fragment of the 5`-region of the luciferase gene, was used as template. The CSEn mutant EM1 (Jiang and Eberhardt, 1994), which was engineered to contain a PstI site was cut with PstI and religated to remove the bulk of the enhancer, the three polyadenylation stops in pA(3)LUC and nt -493/-282 of the hCS 5`-FR. The resulting plasmid was subjected to EcoRI/BamHI digestion, T(4) Polymerase treatment and religation to yield the 3.7-kb pEB.LUC which removed the 3`-end of the luciferase gene. Following inverted PCR mutagenesis with the oligonucleotide primers shown in Table 1, the MscI/NarI fragments were removed from these mutants by restriction digestion and subcloned into MscI/NarI digested EnA-hCSp.LUC generating EnAPPM1-, EnAPPM2-, EnAPPM3-, and EnAPPM4-493CSp.LUC. Plasmids without the hCS enhancer were generated from the plasmids described above by BglII digestion and religation. To authenticate the transcription initiation site in primer extension assays, the plasmid hCSp.Nar/LUC, containing a 139 bp ``stuffer'' sequence in the 5` end of the luciferase gene, was constructed. The plasmid pBluescript (Stratagene) was digested with HpaII/TaqI and the small DNA fragments (50-200 bp) were isolated and ligated to NarI digested hCSp.LUC. One construct, containing a 139-bp pBluescript ``stuffer'' was sequenced and designated hCSp.Nar/LUC.

Gel Shift Assays

Double-stranded oligonucleotides (Table 1) designated in the figure legends were used as probes. The upper strand oligonucleotide (50 ng) was 5`-end labeled with [-P]ATP and polynucleotide kinase. After the labeling reaction, a slight excess of lower strand oligonucleotide was added and annealed to the upper strand by heating to 85 °C for 3 min followed by slow cooling to room temperature. DNA probes were purified from the reaction mixture by passing through a Bio-Gel P-60 column. The sequences of the various double-stranded oligonucleotide competitors are listed in Table 1. Crude nuclear extracts were isolated from cultured cells according to Dignam et al.(1983) or Shapiro et al.(1988). Protein (5 µg) was diluted to 16 µl with buffer D (20 mM Hepes (pH 7.9), 100 mM KCl, 20% glycerol, 0.2 mM EDTA, 0.5 mM phenylmethylsulfonyl fluoride, 0.5 mM DTT. MgCl(2) and poly(dI-dC) in a volume of 8 µl were added to a final concentration of 10.0 mM and 0.1 mg/ml, respectively. After incubation at room temperature for 15 min, 10,000 to 20,000 cpm of probe in a volume of 2 µl was added to the reaction mixture and incubation continued for 15 min. The samples were loaded immediately onto a 4.0% polyacrylamide gel which had been prerun for 1 h. Gels were electrophoresed in 0.5 times TBE at 4 °C for 2 h at 220 V (constant voltage). After electrophoresis, gels were dried in vacuo and exposed to Kodak x-ray film at -80 °C with intensifying screens for 16-48 h.

UV Cross-linking Assays

To estimate the molecular mass of the protein binding to the proximal promoter, a gel shift assay with PCB/MCB probe (Table 1) and GC or BeWo nuclear extracts was performed, except that 0.4 mg/ml poly(dI-dC), 15 µg protein and 50,000 cpm of probe were included in each reaction. After electrophoresis, the wet gel was placed on ice and exposed to UV light for 1 h in a UV Stratalinker 1800 (Stratagene). The gel was covered by a layer of Saran Wrap and exposed to x-ray film at 4 °C for 1.5 h to visualize the DNA-protein complex. The region containing the DNA-protein complex was removed from the gel, and the gel slice was soaked in 200 µl of 2 times SDS-polyacrylamide gel electrophoresis sample buffer (10 mM TrisbulletHCl (pH 6.8), 5.0% SDS, 20.0% glycerol, 200 mM DTT, and 0.1% bromphenol blue) at 4 °C for 1 h. Gel slices were then carefully placed into wells of an 8.0% SDS-polyacrylamide gel and prestained protein standards (Bio-Rad) were simultaneously electrophoresed to estimate the molecular mass. After electrophoresis, the gel was dried and exposed to x-ray film at -80 °C for 16 h with intensifying screens.

Primer Extension Reactions

BeWo cells from four standard electroporations were pooled and plated onto 10-cm dishes. After 18 h, total RNA was extracted from transfected cells according to Xie and Rothblum(1991). The quality of RNA was checked on non-denaturing 1.0% agarose gels to confirm the presence of 18 and 28 S rRNA. Single-stranded pA03 primer was 5`-end labeled and purified following the same procedures as used in preparation of the gel shift probes. Primer extensions were carried out according to Vaccaro et al.(1990) with minor modifications. Briefly, 20 µg of total RNA was ethanol precipitated, washed with 70% ethanol, and redissolved in 10 µl of hybridization buffer (100 mM TrisbulletHCl (pH 7.6), 300 mM KCl, 1 mM EDTA, 10 mM DTT). Radiolabeled primer (5 times 10^5 cpm, pA03) was added and mixed well with the RNA. The mixtures were heated to 85 °C for 5 min and subsequently incubated at 58 °C for 30 min. After hybridization, samples were quickly chilled on ice, and 10 µl of reverse transcription buffer (100 mM TrisbulletHCl (pH 7.6), 20 mM MgCl(2), 1 mM dNTPs, and 60 µg/ml actinomycin D) were added. After addition of 10 units of Moloney murine leukemia virus reverse transcriptase, samples were incubated for 1 h at 37 °C. The volume of each sample was brought up to 120 µl with 1 times TE buffer (10 mM TrisbulletHCl (pH 7.5), 1 mM EDTA). Proteins were eliminated by phenol-chloroform extraction. The extended DNA fragments were recovered by ethanol precipitation and the DNA pellets were washed with 70% ethanol and redissolved in 6.0 µl of 1 times TE and 4.0 µl of sequence loading buffer (1 mg/ml xylene cyanol FF, 1 mg/ml bromphenol blue, 10 mM EDTA (pH 8.0), 50% formamide). The samples were heated to 95 °C for 2 min, chilled on ice, and loaded on an 8% acrylamide/8 M urea gel. The dideoxy sequencing reactions generated from pA03 primers and hCSp.LUC template were loaded as markers. Autoradiography was carried out at -80 °C with intensifying screens for 2-3 days.


RESULTS

Sp1 Modulates Basal and hCS Enhancer-stimulated hCS Promoter Activity, but Is Not Essential for Enhancer Activity

Previous studies of hCS 5`-FR deletion mutants fused to the chloramphenicol acetyltransferase gene provided evidence that Sp1 or an Sp1-like transcriptional factor was important for transcription regulation of the hCS enhancer in placental cells (Fitzpatrick et al., 1990). Since the hCS and hGH promoters are homologous (approx96.5% sequence identity), our previous data had demonstrated that CSEn stimulated both the hGH and hCS promoters equally in BeWo cells (Jiang and Eberhardt, 1994), and previous data have implicated both Sp1 and GHF1 as important modulators of hCS promoter function in GC cells, (^2)we wished to re-examine the hCS promoter elements that were important for its basal and enhancer-stimulated activity in placental cells. Although, GHF1 is not expressed in placental cells, (^3)the possibility that some ubiquitous Oct-related factor might bind to the GHF1 sites and modulate hCS promoter function could not be excluded. We therefore examined hCS promoter constructs containing site-specific mutations in the Sp1, pGHF1, dGHF1, and pdGHF1 binding sites (Fig. 1A) in placental cells. The abilities of the wild-type and mutant hCS promoter in the presence and absence of CSEn to drive transcription of a luciferase reporter gene were tested in transiently transfected BeWo cells. Modifications of the pGHF1, dGHF1, or both (pdGHF1) binding sites had no significant effects on the basal or enhancer-stimulated activities (Fig. 1B), indicating that the binding of Oct-related factors to GHF1 sites is not involved in regulating hCS promoter activity in BeWo cells. In contrast, mutation of the Sp1 site resulted in a approx75% decrease in basal activity (Fig. 1B). Importantly, mutation of the Sp1 site only reduced the relative CSEn activity by approx56% (Fig. 1C; p < 0.01), indicating that promoter factors in addition to Sp1 were essential for CSEn function.


Figure 1: Sp1 is required for maximal CSEn activity; however, additional promoter factors are essential for CSEn function. A, diagram of the hCS 5`-flanking region, highlighting the Sp1 and proximal (p) and distal (d) GHF1 binding sites (sequences enclosed in boxes). The mutagenesis scheme for altering the Sp1 and GHF1 sites is shown below the sequence. Colons designate deleted nucleotides. B, the data reflect basal- and enhancer-stimulated activity [(light units/µg protein ± S.E.) times 10] of wild-type hCSp.LUC and the various Sp1 and GHF1 mutants in transiently transfected BeWo cells. Individual p values for significance were derived as described under ``Materials and Methods.'' ANOVA analysis for the effect of mutation was significant at the p = 0.0001 level. C, the data in A plotted as fold stimulation. ANOVA analysis for the effect of mutation was significant at the p = 0.017 level.



Because the hCS Sp1 site (GGGAGG) is a variant of the canonical site (GGGCGG) and has been reported to have a lower binding affinity for Sp1 (Letovsky and Dynan, 1989), we examined whether reversion of the Sp1 site to the canonical sequence could result in a stronger promoter and increased enhancer-stimulated activity. As shown in Fig. 1, B and C, the Sp1UP mutation did increase promoter activity 1.6-fold (statistically insignificant). However, there was no corresponding effect of the Sp1UP mutation on enhancer activity (Fig. 1, B and C). These experiments suggest that the levels of Sp1 in BeWo cells may be sufficiently high to maintain a relatively high occupancy at the Sp1 site despite the anticipated lower affinity of the GGGAGG binding site (Letovsky and Dynan, 1989). These results confirm the findings of Fitzpatrick et al.(1990) which indicate that Sp1 modulates hCS basal- and enhancer-stimulated promoter activity. Nevertheless, since the enhancer activity was not eliminated upon mutation of the Sp1 site, the data suggest that other promoter factors might be involved in mediating enhancer function.

Proximal Elements Are Essential for hCS Promoter and Enhancer Function

Since the results above suggested that other promoter factors might be required for enhancer function, we examined additional hCS 5`-FR deletion mutants in transiently transfected BeWo cells (Fig. 2A). Deletion of hCS 5`-FR to nt -282 (282CSp.LUC and EnA282CSp.LUC) had no impact on promoter or enhancer-stimulated activity (Fig. 2, B and C), indicating that sequences upstream of the PstI site were not required for enhancer activity and confirming the data of Fitzpatrick et al.(1990). Further deletion down to nt -36 (36CSp.LUC and EnA36CSp.LUC) generated a core promoter containing the TATA box and resulted in significantly decreased (15% of control) basal activity (Fig. 2B). However, CSEn stimulated the activity of this minimal promoter 8.8-fold (70% of wild-type, Fig. 2C), a value not significantly different from the wild-type promoter. These data suggest that proximal promoter elements mediate the enhancer activity. Deletion of the TATA box (19CSp.LUC and EnA19CSp.LUC) resulted in a very low basal activity (3.7% wild type, Fig. 2B) and CSEn-simulated activity was markedly decreased (20% wild-type, p <0.01, Fig. 2C), suggesting that the TATA box might be required for enhancer function. Nevertheless, a 19-bp deletion of the TATA box (493CSDeltaTATAp.LUC) in the context of an otherwise normal 493CS 5`-FR resulted in a promoter with 28% of wild-type basal activity (Fig. 2B) and unaffected enhancer-stimulated activity (103% wild-type, Fig. 2C). Since this latter construct contains the Sp1 site, it is possible that Sp1 may compensate for the loss of function conferred by the TATA box.


Figure 2: The TATA element and sequences downstream are required for maximal CSEn function. A, diagram and sequence of the proximal hCS 5`-flanking region with mutagenesis strategy. Colons designate deleted nucleotides. B, basal- and enhancer-stimulated activity ((light units/µg of protein ± S.E.) times 10) of wild-type hCSp.LUC and the various proximal promoter mutants. ANOVA analysis for the effect of mutation was significant at the p = 0.0001 level. C, the data in B plotted as fold CSEN-stimulated activity. ANOVA analysis for the effect of mutation was significant at the p = 0.0001 level. D, basal- and enhancer-stimulated activity ((light units/µg of protein ± S.E.) times 10) of wild-type hCSp.LUC and the mutants of the InrE region. ANOVA analysis for the effect of mutation was significant at the p = 0.0001 level. E, the data in D plotted as fold CSEn-stimulated activity. ANOVA analysis for the effect of mutation was significant at the p = 0.0005 level.



Removal of sequences downstream of the TATA box (nt -19/+1, 493CSAPADeltap.LUC and EnA493CSAPADeltap.LUC) had a significant effect on the basal activity (35% reduction) but decreased enhancer-stimulated activity by 60% (Fig. 2, B and C) in the context of a normal 493CS 5`-FR containing the Sp1 site, suggesting that sequences around the transcription initiation site might be required for promoter activity. Deletion of the Sp1 site in this latter construct (36CSAPADelta and EnA36CSAPADelta) resulted in a marked reduction of basal activity (4.5% wild-type, Fig. 2B) and enhancer-stimulated activity (15.4% wild-type, Fig. 2C). Thus in the presence of the Sp1 binding site, the sequences downstream of the TATA box appear to be more important for mediating promoter function than the TATA box itself. As in the case with the Sp1 mutant, mutation of the proximal promoter elements did not result in a complete loss of enhancer activity, suggesting that no single promoter element was essential for enhancer function.

Sequences Near the Transcription Initiation Site Are Required for hCS Promoter Function

To examine the role of sequences between the TATA box and transcription initiation site in mediating enhancer function in more detail, proximal promoter block mutants (PPM1-PPM4, Fig. 2A) were created and tested for basal and enhancer stimulated activity in BeWo cells. Mutation of the sequences just 3` of the TATA box (PPM1, nt -20/-12) did not affect basal or enhancer stimulated activity significantly (Fig. 2, D and E). In contrast, mutation of the sequences between -15/-6 (PPM2), -10/-1 (PPM3), and -5/+5 (PPM4) reduced basal activity by 48, 62, and 58%, respectively. The enhancer-stimulated activity of mutant PPM2 (60% wild-type activity) was not significantly reduced; however, the enhancer-stimulated activities of PPM3 and PPM4 were reduced by 57 and 63%, respectively (Fig. 2, D and E). These data suggest that a transcription initiator element (InrE) located between nt -15/+1 is required for efficient promoter and maximal enhancer activity.

A Nuclear Factor Preferentially Expressed in Choriocarcinoma Cells Binds to the InrE to Form a Distinct DNA-Protein Complex

We examined whether the hCS InrE could be recognized by nuclear factors utilizing a gel shift assay. A P-end-labeled PCB/MCB probe (Table 1) containing nt -20/+1 of the hCS 5`-FR was incubated with BeWo, GC, HeLa, SV40 transformed fibroblasts (GM0637E), osteosarcoma cells (MG63), COS, and JEG-3 cell nuclear extracts, and the free and protein-bound probes were separated on non-denaturing gels. Nuclear proteins from GC, HeLa, GM0637E, and COS cells formed several prominent, nearly identical DNA-protein complex patterns (H2-H6, Fig. 3A) which had a significantly lower mobility than that formed with BeWo or JEG-3 cells (H1, Fig. 3A). The different patterns observed with BeWo and JEG-3 compared to the other cell nuclear proteins suggested that a specific nuclear factor that is preferentially expressed in placental cells may bind to the InrE. Lesser amounts of H1 complexes appear to be present in MG63 and some of the other cell types, indicating that the protein forming this complex may not be restricted to placental cells. Since the complex H1 migrated more rapidly than complexes from the other cell types, we examined the possibility that it might be generated by proteolysis by examining the effect of extended incubation, freeze-thaw and mixing experiments (Fig. 3B). Extended incubation at 37 °C or subjecting extracts to multiple freeze-thaw cycles had no effect on the complex migration patterns. In addition, mixing BeWo with GC or HeLa extracts produced a pattern that was the sum of the individual extracts. Finally, multiple independent preparations of nuclear extracts exhibited identical behavior, so that the complex patterns do not appear to be the result of proteolysis or due to the binding of additional proteins present in GC or HeLa cells that are not present in BeWo cells.


Figure 3: A nuclear protein that is preferentially expressed in choriocarcinoma cells binds to the hCS InrE. A, gel shift analysis of protein-DNA complexes (H1-H6) formed with a P-labeled DNA fragment (PCB/MCB) containing nucleotides -20/+1 of the hCS 5`-flanking DNA and nuclear extracts from BeWo, GC, HeLa, GM0637E (SV40 transformed human fibroblasts), MG63 (osteosarcoma), COS, and JEG-3 cells. B, effect of extended incubation (INC, 37 °C for 20 min), freeze-thaw (FT, four cycles), and mixing 4 µg each of BeWo and GC or HeLa cell nuclear extracts on the profiles of the complexes H1, H2, and H4-H6.



The specificity of the protein-DNA complexes formed with BeWo cell nuclear proteins was characterized further by oligonucleotide binding and competition experiments. For these experiments three additional oligonucleotides containing mutations within a 4-5 bp region from nt -12/+1 were examined (Fig. 4A). Both PCB/MCB-MUT1 and PCB/MCB-WT oligonucleotides formed complex H1 equally well, whereas PCB/MCB-MUT2 and PCB/MCB-MUT3 failed to form complex H1 (Fig. 4A). Similarly, PCB/MCB-MUT1 and PCB/MCB-WT, but not PCB/MCB-MUT2 and PCB/MCB-MUT3, competed for H1 complex formation to PCB/MCB (Fig. 4B). Similar results were obtained with competition experiments with DNA fragments corresponding to the proximal promoter mutants (PPM1-PPM4) that were employed in the functional studies (data not shown). These experiments indicate that the binding to the proximal promoter is sequence-specific and that the recognition site is localized between nt -12/+1. Thus the sequences that are important for mediating basal and enhancer-stimulated activity co-localize with the sequences required for formation of the BeWo nuclear protein-DNA complex. Taken together, these data suggest that a factor binding at the InrE just downstream from the TATA box is required for hCS promoter function.


Figure 4: The InrE-binding protein present in choriocarcinoma cells recognizes specific sequences located between nucleotides -12/-1 of the hCS promoter. A, binding of BeWo cell nuclear extract to P-labeled PCB/MCB and P-labeled mutant oligonucleotides, PCB/MCB-MUT1, PCB/MCB-MUT2, and PCB/MCB-MUT3. The sequences of the oligonucleotides are shown to the right of the gel shift experiment. B, competition analysis of the BeWo nuclear protein-DNA complex H1 formed with the P-labeled PCB/MCB probe and with the wild-type and mutant oligonucleotides PCB/MCB-MUT1 (MUT1), PCB/MCB-MUT2 (MUT2), and PCB/MCB-MUT3 (MUT3). The fold excess (weight) of the competing oligonucleotides is given (FOLD).



A Protein of 70-kDa Mass Binds to the InrE in the Proximal hCS Promoter

To estimate the size of the protein binding to the InrE, DNA-protein cross-linking procedures were employed. Following UV irradiation of a gel containing [P]DNA-labeled complexes H1 from BeWo cells and H2 from GC cells (Fig. 3A), the complexes were analyzed by SDS-PAGE. A single band of 70 kDa was visualized from the BeWo protein-DNA complex (Fig. 5). A different pattern, however, composed of a very minor amount of a 70-kDa band and two major lower bands of 30 and 20 kDa, respectively, were observed from the GC cell H2 complex. These data indicate that the proximal hCS promoter binds distinctly different proteins and suggest that the 70-kDa BeWo nuclear protein that binds the InrE may account in part for the cell-specific expression of this promoter in placental cells.


Figure 5: A BeWo cellular protein of 70 kDa binds to the hCS 5`-flanking region in the vicinity of the InrE. BeWo and GC cell nuclear protein-DNA complexes formed with P-labeled PCB/MCB DNA were UV cross-linked as described under ``Materials and Methods,'' and the products were separated on SDS-polyacrylamide gels and visualized by autoradiography. The molecular mass of the DNA fragment was approx20 kDa.



Mutations in the hCS InrE Result in Alteration of Transcriptional Start Sites

One possible role of the factor binding at the InrE may be to direct the transcription initiation site. We therefore compared the transcriptional start sites on the wild-type and mutated hCS promoters to provide information about the roles of the InrE and Sp1 in directing transcription initiation. Initially, the start site used by the wild-type hCS promoter was determined by primer extension assay (Fig. 6A). To establish the authenticity of the start site on the hCSp.LUC gene, a 139-bp ``stuffer'' fragment was inserted at the NarI site between the pA03 primer and the hCS promoter to yield the plasmid hCSp.Nar/LUC. Primer extension of RNA derived from this plasmid should generate a transcript 139 bp longer than the transcript from the hCSp.LUC gene, allowing a verification of the transcript termination site. In addition, RNA isolated from mock- and promoterless pA(3)LUC-transfected cells were included as negative controls. As shown in Fig. 6A, the same initiation site (marked +1) as identified previously by Selby et al.(1984) from human placental RNA was utilized in BeWo cells. As expected, RNA from hCSp.Nar/LUC-transfected cells yielded an extension product shifted 139 bp further upstream, confirming the authenticity of the initiation site.


Figure 6: Sequences near the hCS gene InrE are required for accurate transcription initiation site selection and define a probable initiator element (InrE). A, primer extension analysis of RNA from BeWo cells transfected with the wild-type hCSp.LUC, pALUC (promoterless control) and hCSp.Nar/LUC genes (containing a 139-bp insert to assess specificity of the transcription initiation site). Cellular RNA was from mock-transfected BeWo cells. A and G represent dideoxynucleotide sequencing reactions utilizing the same primer used for primer extension with the wild-type hCSp.LUC plasmid DNA. B, primer extension analysis of total RNA from BeWo cells transfected with 493CSp.Nar/LUC and 36CSp.Nar/LUC. C and T represent dideoxynucleotide sequencing reactions utilizing the same primer used for primer extension. C, primer extension analysis of total cellular RNA (PPM1-PPM4) from BeWo cells transfected with the 493CSPPM1-PPM4p.LUC genes. T and C represent dideoxynucleotide sequencing reactions utilizing the same primer used for primer extension. D, sequences and transcription initiation site selection in the various hCS proximal promoter mutants derived from the data in C.



Initiation sites from truncated and mutated promoters are shown in Fig. 6, B and C, and are summarized in Fig. 6D. Deletion of the upstream sequences, including the Sp1 site, to nt -36 (36CSp.Nar/LUC) did not alter the transcription initiation site (Fig. 6B), supporting the concept that initiation site selection is solely determined by the core promoter. The PPM1 and PPM2 mutations just upstream from the region required for InrE binding by the BeWo cell nuclear protein did not significantly affect the use of the authentic start site (Fig. 6C). However, the PPM3 and PPM4 mutations between nt -10/+5 shifted the transcriptional start sites 2 and 3 nt downstream, respectively (Fig. 6C). Thus loss of initiation site focus is correlated with the effects of these proximal promoter mutations on basal- and enhancer-stimulated promoter activity and factor binding at the InrE, providing additional evidence that the InrE-binding factor is involved in transcription initiation. These results suggest that Inr elements are required for accurate transcription initiation from the CS promoter in placental cells.


DISCUSSION

The hCS enhancer is composed of multipartite enhansons which bear striking similarities with SV40 enhansons (Jacquemin et al., 1994; Jiang and Eberhardt, 1994), particularly the GT-IIC and SphI/SphII enhansons. The unique location of CSEn at the extreme 3`-end of the hGH/hCS gene cluster, placing it in proximity to the placenta-expressed genes, and the highly restricted function of this enhancer to placental cells suggest that CSEn might play a dominant role in the control of hCS gene expression. Nevertheless, the similarity of CSEn to the SV40 enhancer, and the possibility that it might be controlled by the ubiquitous factor TEF-1 (Jacquemin et al., 1994; Jiang and Eberhardt, 1994; Walker et al., 1990) that binds the GT-IIC and SphI/SphII enhansons (Davidson et al., 1988; Xiao et al., 1991), raise questions about possible cell-specific control mechanisms. For example, if CSEn is controlled by TEF-1, why is it not functional in cell types which are known to have abundant TEF-1 activity? Previous data had also shown that the ubiquitous Sp1 played an important role in mediating enhancer function (Fitzpatrick et al., 1990), providing additional questions about how this enhancer could be controlled in a cell-specific manner. Consequently, we decided to study the interaction of the enhancer with the hCS promoter to ascertain if clues about the cell-specific mechanism could be obtained. We reasoned that the enhancer stimulatory effect results from the synergistic action of enhancer and promoter elements and that cell-specific factors might be dependent on such interactions. In the current studies, we provide evidence that in addition to Sp1, the TATA box and an initiator element (InrE) located near the transcription initiation site are required for optimal enhancer activity. The InrE binding factor may be a cell-specific factor since it is preferentially expressed in choriocarcinoma cells (BeWo and JEG-3 cells), but is absent or present in very low concentrations in a variety of other cell types (Fig. 3).

Previous studies had indicated that Sp1 appeared to play a dominant role in CSEn function in Jar and JEG-3 cells (Fitzpatrick et al., 1990). Since Sp1 serves as an enhanson in several enhancers, including those for bovine papilloma virus (Li et al., 1991), chicken histone H5 (Sun et al., 1992) and mouse pro-alpha2(I) collagen genes (Pogulis and Freytag, 1993), it was possible that Sp1 was an integral and essential component of enhancer function. We confirmed the earlier observations that Sp1 contributes in a major way to hCS promoter activity (Fig. 1B) in human placental cells (Fitzpatrick et al., 1990). However, our studies demonstrate that CSEn maintains significant, albeit reduced, enhancer activity in the absence of the Sp1 site (Fig. 1C), suggesting that CSEn interactions with other parts of the basal transcription machinery are required. Sp1 does not participate in the selection of the transcription initiation site, since a core hCS promoter without Sp1 binding site can direct faithful initiation in placental cells (Fig. 6B). Similar results, demonstrating that neither the number nor orientation of Sp1 binding sites affected initiation site selection on other TATA box-containing promoters, have been observed (Jones and Tjian, 1985; Segal and Berk, 1991). Thus the role of Sp1 can be distinguished from those factors that direct initiation site selection. Importantly, as discussed below, the evidence indicates that hCS promoter function requires an InrE-binding factor which is involved in transcription initiation.

Support for the concept that factors in addition to Sp1 were required for optimal enhancer-stimulated activity was obtained in subsequent mutational studies of the hCS proximal promoter (Fig. 2) which indicated that the TATA element and downstream sequences played an important role in hCS promoter and enhancer function. Direct evidence for an initiator factor was obtained from gel shift experiments ( Fig. 3and Fig. 4) and UV cross-linking experiments (Fig. 5) which demonstrate that a BeWo cell-specific protein of 70 kDa binds to the hCS promoter near the InrE. The specificity of the hCS InrE-protein interaction was established by a series of binding and competition experiments with wild-type and mutated oligonucleotides (Fig. 4). Site-specific mutation of the region between nt -15/+1 resulted in significantly reduced basal- and enhancer-stimulated activity (Fig. 2, D and E) and resulted in altered transcription initiation site selection (Fig. 6C), providing additional evidence that an InrE was required for basal- and enhancer-mediated hCS promoter activity. Since the InrE-binding ability and initiation site selection are strongly correlated, we propose that the 70-kDa protein binding to the hCS InrE might play a crucial role in the formation of the initiation complex.

The requirement of the DNA sequences around the InrE for faithful and efficient transcription is well known (Nakatani et al., 1990; Smale and Baltimore, 1989; Talkington and Leder, 1982). Several families of InrEs based on consensus sequences have been described (reviewed in Weis and Reinberg(1992)). These families include the TdT-InrE (YYCAYYYYY^6), PBGD-InrE (CA^1 and ^5TCCTGGTTAC^14), DHFR-InrE (ATTTCGCGCCAAACTT^5), the ribosomal protein InrE (CTTCCCTTTTCC^8) and the adeno-associated virus p5-InrE (CTCCATTTT). All of these sequences are pyrimidine rich (>63%), but there is no obvious relationship in the sequences between nt -15/+1 of the hCS promoter (38% pyrimidine content) with any of these InrE families, suggesting that the hCS InrE may be a unique element. Javahery et al.(1994) have provided evidence that many reported Inrs fit the consensus YYA(+1)N(T/A)YY at the transcription initiation site; however, this does not fit the hCS initiation site (CTA^1GGA), providing additional evidence that the hCS InrE may be unique. It is interesting that the region containing the hCS InrE contains an imperfect palindrome (AGAGACCGGCTCT) and that the InrE binding protein fails to bind to oligonucleotides containing mutations in the region -8/-5 and -5/-1; however, the significance of this palindromic structure is unknown.

By footprinting and gel shifting assays, several InrEs have been shown to interact with specific nuclear proteins (Du et al., 1993; Means and Farnman, 1990; Nakatani et al., 1990; Purnell and Gilmour, 1993). Two different InrE-binding factors from HeLa cells, YY1 (adeno-associated virus p5-InrE-binding factor) (Shi et al., 1991) and TFII-I (TdT-InrE-binding factor) (Roy et al., 1991) have been extensively characterized, demonstrating that InrE function is dependent on unique transcription factors. Although InrEs are essential for the function of TATA-less promoters, they also have been shown to function on TATA-containing promoters. Cooperative interactions between TATA and Inr elements may be important for the functioning of several promoters (Conaway et al., 1990; Nakatani et al., 1990; Purnell and Gilmour, 1993; Smale and Baltimore, 1989; Tokunaga et al., 1984), but not the major histocompatibility complex E1 promoter (Mantovani et al., 1993). Such cooperative interactions appear to be important for the hCS promoter (Fig. 2C).

Although the exact mechanism(s) involved in InrE-mediated transcription are not known, evidence has been obtained that TBP is required for TATA- and InrE-mediated transcription. Smale et al.(1990) demonstrated that a TFIID-containing fraction restored TdT-InrE-mediated transcription in a heat-inactivated nuclear extract. Subsequently, Carcamo et al.(1991) showed that transcription from the Ad-ML promoter lacking the TATA element was TBP-dependent and Pugh and Tjian(1991) demonstrated that polyclonal anti-TBP antibodies inhibited transcription from both TATA-containing and TATA-less promoters. It has been proposed that InrE-binding proteins may tether TBP to the transcription initiation complex through protein-protein interactions (Mantovani et al., 1993; Pugh and Tjian, 1991). On promoters like hCS containing both TATA and Inr elements, TBP may be more stably associated with the initiation complex by binding to the TATA element and by its interaction with the InrE-binding protein. This could explain why mutation or deletion of the TATA or Inr elements result in reduction of hCS promoter function and enhancer-stimulated activity. Accordingly, our data are consistent with a model in which the TATA and Inr elements form a basic initiation complex which directs a low level, faithful transcription whose initiation frequency is increased by Sp1 and the other enhancer elements.

Our results indicate that no single, specific promoter element is responsible for enhancer activity. All of the promoter elements that were found to be necessary for hCS enhancer stimulation also made significant contributions to the basal promoter activity. Thus the enhancer may interact with ubiquitous transcription factors that form part of the basal transcriptional machinery. This idea is supported by the data showing that the enhancer functions with a core promoter containing the TATA and Inr elements (Fig. 2C) and has an absolute requirement for a promoter, since the enhancer cannot initiate transcription by itself (Fig. 6A). These results are consistent with our earlier observation that CSEn can stimulate cell-specific transcription from the heterologous thymidine kinase and RSV promoters (Jiang and Eberhardt, 1994), which lack a structure similar to the hCS InrE. Thus the InrE-binding factor may contribute to hCS promoter efficiency which provides for optimal enhancer modulation of promoter activity. Although there is no evidence for a direct interaction between the InrE-binding protein and the enhancer complex, it cannot be excluded that the InrE-binding protein may be utilized in promoters which lack the InrE binding site by basal factor- or enhancer-mediated tethering mechanisms (Zenzie-Gregory et al., 1992). Although these data do not completely explain cell-specific hCS gene expression, the data provide the first evidence for an InrE in any member of the human growth hormone gene family, provide evidence that the InrE accounts in part for cell-specific expression of the hCS promoter and demonstrate that this element is essential for maximal basal- and enhancer-mediated hCS transcription in placental cells.


FOOTNOTES

*
This work was supported by National Institutes of Health Grant DK41206 (to N. L. E.). The costs of publication of this article were defrayed in part by the payment of page charges. This article must therefore by hereby marked ``advertisement'' in accordance with 18 U.S.C. Section 1734 solely to indicate this fact.

§
To whom correspondence should be addressed: 4-407 Alfred, SMH, Mayo Clinic, Rochester, MN 55905. Tel.: 507-255-6554; Fax: 507-255-4828; eberhardt{at}mayo.edu.

(^1)
The abbreviations used are: hCS, human chorionic somatomammotropin (also known as placental lactogen); CSEn and En, placental-specific chorionic somatomammotropin gene enhancer associated with the hGH/hCS gene locus; hGH, human growth hormone; rGH, rat growth hormone; GHF1/Pit1, pituitary-specific transcription factor; InrE, transcription initiator element; DTT, dithiothreitol: LUC, luciferase; 5`-FR, 5`-flanking region; ANOVA, analysis of variance; RSV, Rous sarcoma virus; PCR, polymerase chain reaction; PBS, phosphate-buffered saline; bp, base pair(s); nt, nucleotide(s).

(^2)
A. R. Shepard and N. L. Eberhardt, unpublished results.

(^3)
S.-W. Jiang, unpublished results.


ACKNOWLEDGEMENTS

We thank Dr. Peter Cattini for providing the plasmid containing the 242-bp CSEn fragment, Dr. William Wood for providing the luciferase vector pALUC, Dr. Cheryl Conover for providing the GM0637E and MG63 cells, and Mary Craddock for secretarial and editorial help with the preparation of the manuscript.


REFERENCES

  1. Bodner, M., and Karin, M. (1987) Cell 50, 267-275 [Medline] [Order article via Infotrieve]
  2. Borody, I. B., and Carlton, M. A. (1981) Br. J. Obstet. Gynaecol. 88, 447-449 [Medline] [Order article via Infotrieve]
  3. Brelje, T. C., Scharp, D. W., Lacy, P. E., Ogren, L., Talamantes, F., Robertson, M., Friesen, H. G., and Sorenson, R. L. (1993) Endocrinology 132, 879-887 [Abstract]
  4. Carcamo, J., Buckbinder, L., and Reinberg, D. (1991) Proc. Natl. Acad. Sci. U. S. A. 88, 8052-8056 [Abstract]
  5. Conaway, J. W., Travis, E., and Conaway, R. C. (1990) J. Biol. Chem. 265, 7564-7569 [Abstract/Free Full Text]
  6. Davidson, I., Xiao, J. H., Rosales, R., Staub, A., and Chambon, P. (1988) Cell 54, 931-942 [Medline] [Order article via Infotrieve]
  7. Dignam, J. D., Lebovitz, R. M., and Roeder, R. G. (1983) Nucleic Acids Res. 11, 1475-1489 [Abstract]
  8. Du, H., Roy, A. L., and Roeder, R. G. (1993) EMBO J. 12, 501-511 [Abstract]
  9. Fitzpatrick, S. L., Walker, W. H., and Saunders, G. F. (1990) Mol. Endocrinol. 4, 1815-1826 [Abstract]
  10. Freemark, M., and Comer, M. (1989) J. Clin. Invest. 83, 883-889 [Medline] [Order article via Infotrieve]
  11. Freemark, M., Kirk, K., Pihoker, C., Robertson, M. C., Shiu, R. P. C., and Driscoll, P. (1993) Endocrinology 133, 1830-1842 [Abstract]
  12. Handwerger, S. (1991) Endocr. Rev. 12, 329-336 [Abstract]
  13. Hemsley, A., Arnheim, N,, Toney, M., D., Cortopassi, G., and Galas, D. J. (1989) Nucleic Acids Res. 17, 6545-6551 [Abstract]
  14. Hirt, H.. Kimelman, J., Birnbaum, M. J., Chen, E. Y., Seeburg, P. H., Eberhardt, N. L., and Barta, A. (1987) DNA (N. Y.) 6, 59-70 [Medline] [Order article via Infotrieve]
  15. Ingraham, H. A., Chen, R., Mangalam, H. J., Elsholtz, H. P., Flynn, S. E., Lin, C. R., Simmons, D. M., Swanson, L., and Rosenfeld, M. G. (1988) Cell 55, 519-529 [Medline] [Order article via Infotrieve]
  16. Jacquemin, P., Oury, C., Peers, B., Morin, A., Belayew, A., and Martial, J. A. (1994) Mol Cell. Biol. 14, 93-103 [Abstract]
  17. Javahery, R., Khachi, A., Lo, K., Zenzie-Gregory, B., and Smale, S. T. (1994) Mol. Cell. Biol. 14, 116-127 [Abstract]
  18. Jiang, S.-W., and Eberhardt, N. L. (1994) J. Biol. Chem. 269, 10384-10392 [Abstract/Free Full Text]
  19. Jones, K. A., and Tjian, R. (1985) Nature 317, 179-182 [Medline] [Order article via Infotrieve]
  20. Letovsky, J., and Dynan, W. S. (1989) Nucleic Acids Res. 17, 2639-2653 [Abstract]
  21. Li, R., Knight, J. D., Jackson, S. P., Tjian, R., and Botchan, M. R. (1991) Cell 65, 493-505 [Medline] [Order article via Infotrieve]
  22. Mantovani, R., Tora, L., Moncollin, V., Egly, J. M., Benoist, C., and Mathis, D. (1993) Nucleic Acids Res. 21, 4873-4878 [Abstract]
  23. Maxwell, I. H., Harrison, G. S., Wood, W. M., and Maxwell, F. (1989) BioTechniques 7, 276-280 [Medline] [Order article via Infotrieve]
  24. Means, A., and Farnman, P. J. (1990) Mol. Cell. Biol. 10, 653-661 [Medline] [Order article via Infotrieve]
  25. Miller, W. L., and Eberhardt, N. L. (1983) Endocr. Rev. 4, 97-130 [Medline] [Order article via Infotrieve]
  26. Nachtigal, M. W., Nickel, B. E., Klassen, M. E., Zhang, W. G., Eberhardt, N. L., and Cattini, P. A. (1989) Nucleic Acids Res. 17, 4327-4337 [Abstract]
  27. Nakatani, Y., Horikoshi, M., Brenner, M., Yamamoto, T., Besnard, F., Roeder, R. G., and Freese, E. (1990) Nature 348, 86-88 [CrossRef][Medline] [Order article via Infotrieve]
  28. Nielsen, P. V., Pedersen, J., and Kampmann, E. M. (1979) Am. J. Obstet. Gynecol. 135, 322-326 [Medline] [Order article via Infotrieve]
  29. Pogulis, R. G., and Freytag, S. O. (1993) J. Biol. Chem. 268, 2493-2499 [Abstract/Free Full Text]
  30. Pugh, B. F., and Tjian, R. (1991) Genes & Dev. 5, 1935-1945
  31. Purnell, B. A., and Gilmour, D. S. (1993) Mol. Cell. Biol. 13, 2593-2603 [Abstract]
  32. Rogers, B. L., Sobnosky, M. G., and Saunders, G. F. (1986). Nucleic Acids Res. 14, 7647-7659 [Abstract]
  33. Roy, A. L., Meisterernst, M., Pognoec, P., and Roeder, R. (1991) Nature 354, 245-248 [CrossRef][Medline] [Order article via Infotrieve]
  34. Segal, R., and Berk, A. J. (1991) J. Biol. Chem. 266, 20406-20411 [Abstract/Free Full Text]
  35. Selby, M., Barta, A., Baxter, J. D., Bell, G. I., and Eberhardt, N. L. (1984) J. Biol. Chem. 259, 13131-13138 [Abstract/Free Full Text]
  36. Shapiro, D. J., Sharp, P. A., Wahli, W. W., and Keller, M. J. (1988) DNA (N. Y.) 7, 47-55 [Medline] [Order article via Infotrieve]
  37. Shepard, A. R., Zhang W., and Eberhardt, N. L. (1994) J. Biol. Chem. 269, 1804-1814 [Abstract/Free Full Text]
  38. Shi, Y., Seto E., Chang, L.-S., and Shenk, T. (1991) Cell 67, 377-388 [Medline] [Order article via Infotrieve]
  39. Smale, S. T., and Baltimore, D. (1989) Cell 57, 103-113 [Medline] [Order article via Infotrieve]
  40. Smale, S. T., Schmidt, M. C., Berk, A. J., and Baltimore, D. (1990) Proc. Natl. Acad. Sci. U. S. A. 87, 4509-4513 [Abstract]
  41. Snedecor, G. W., and Cochran, W. G. (1980) Statistical Methods , 7th Ed, p. 116, The Iowa State University Press, Ames, IA _
  42. Sun, J.-M., Penner, C. G., and Davie, J. R. (1992) Nucleic Acids Res. 20, 6385-6392 [Abstract]
  43. Talkington, C., and Leder, P. (1982) Nature 298, 192-195 [Medline] [Order article via Infotrieve]
  44. Tokunaga, K., Hirose, S., and Suzuki, Y. (1984) Nucleic Acids Res. 12, 1543-1558 [Abstract]
  45. Vaccaro, M., Pawlak, A., and Jost, J.-P. (1990) Proc. Natl. Acad. Sci. U. S. A. 87, 3047-3051 [Abstract]
  46. Walker, W. H., Fitzpatrick, S. L., and Saunders, G. F. (1990) J. Biol. Chem. 265, 12940-12948 [Abstract/Free Full Text]
  47. Weis, L., and Reinberg, D. (1992) FASEB J. 6, 3300-3309 [Abstract/Free Full Text]
  48. Xiao, J. H., Davidson, I., Matthes, H., Garnier, J. M., and Chambon, P. (1991) Cell 65, 551-568 [Medline] [Order article via Infotrieve]
  49. Xie, W.-Q., and Rothblum L. I. (1991) BioTechniques 11, 325-327
  50. Zenzie-Gregory, B., O'Shea-Greenfield, A., and Smale, S. T. (1992) J. Biol. Chem. 267, 2823-2830 [Abstract/Free Full Text]

©1995 by The American Society for Biochemistry and Molecular Biology, Inc.