INTRODUCTION

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The murine Htf9 locus was isolated by virtue of its association with a CG-rich genomic sequence (1) and mapped to mouse chromosome 16 (2). The locus contains two transcriptional units, Htf9-a and Htf9-c, that are transcribed with opposite polarity from complementary DNA strands (see map in Fig. 1A). The lower strand gene, called Htf9-a, encodes Ran-binding protein 1 (RanBP1)¹ (3, 4), an interacting partner of the Ran GTPase, which is thought to cooperate in control of several processes regulated by the Ran network and including DNA replication, cell cycle progression, mitotic entry and exit, nuclear structure, and nucleo/cytoplasmic transport (reviewed in Refs. 5 and 6). The Htf9-c gene codes for a protein sharing extensive homologies with nucleic acid-modifying enzymes, including bacterial tRNA methyltransferases and yeast DNA endo-exonucleases (7); the evolutionary conservation of the protein sequence in the nucleic acid-interacting domain suggests that the murine Htf9-c gene product may also be implicated in nucleic acid metabolism and/or modifications.

Both Htf9 divergently transcribed genes are expressed at low levels in many tissues and cell types (1, 8) and are up-regulated in proliferating cells: transcription is activated at the G₁/S transition of the cell cycle and peaks in S phase, while being repressed in quiescent tissues and growth-arrested cells (7, 9). Promoter deletion and site-directed mutagenesis assays have revealed that cell cycle expression of both divergently transcribed genes is controlled by separate E2F and Sp1 promoter elements active in each orientation (7).

Both the RanBP1 and Htf9-c transcriptional units are initiated in a genomic sequence that has typical features of the CG-rich class of mammalian promoters, which are mainly, although not exclusively, associated with housekeeping genes. An intriguing feature shared by most promoters of the housekeeping class is the absence of a TATA box, whose function in transcriptional initiation is well established for all three classes of RNA polymerase (reviewed in Refs. 10-12). In most TATA-less promoters the DNA region surrounding the major transcriptional start site(s) plays a crucial role in directing initiation (see Refs. 11 and 13 for reviews). Like many CG-rich promoters, the Htf9 bidirectional promoter harbors no TATA box on either DNA strand; RNA transcription is initiated at multiple sites on each strand; transcription start 1 (TS-1 in Fig. 1B) is the most frequently used start site on both DNA strands (1). Deletion mapping analysis (7, 14) restricted the promoter region required for basal transcription to a 74-bp long region active in both orientations. Within that region, genomic footprinting experiments depicted a site protected by unidentified factor(s), located 20 bp apart from a functional Sp1-binding site and flanking TS-1 (15).

The present study was undertaken to characterize the Htf9 footprinted element (HFE) adjacent to TS-1. In an initial set of experiments, we found that double-stranded HFE interacts with factors belonging to the family of retinoid X receptors (RXRs). In addition, distinct factors recognize each single HFE strand. The distinct DNA binding activities depicted with different forms of the HFE element might have indicated that the Htf9 initiation region was organized either in the double-stranded or in the single-stranded conformation in different cellular conditions. Since both Htf9-associated genes are transcribed in a cell cycle-dependent manner, we sought to determine whether the interaction of factors with HFE varies in relation to proliferation and transcription. Our findings show that the double-stranded nucleoprotein complex is efficiently assembled with extracts from liver cells, in which transcription from the Htf9 locus is extremely low or absent, while single stranded DNA binding activities are abundant in cycling NIH/3T3 fibroblasts, in which both Htf9-associated genes are actively transcribed. In addition, the region surrounding HFE acquires an S1-sensitive conformation in vivo in cycling, but not in growth-arrested, NIH/3T3 cells, where transcriptional repression occurs. Therefore, the Htf9 promoter appears to assume alternative conformations in relation to transcription. These data suggest that factors of the RXR family and the newly identified single-stranded binding proteins participate to transcriptional control of the Htf9-associated genes by interacting with alternative forms of the HFE element in different cellular conditions.

	EXPERIMENTAL PROCEDURES

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Cell Cultures-- Murine NIH/3T3 embryo fibroblasts (ATCC CRL 1658) were grown in Dulbecco's modified Eagle's medium supplemented with 10% (v/v) fetal calf serum under 5% CO₂ at 37 °C. Asynchronously cycling cells were collected from 60-70% confluent cultures. Proliferation arrest was induced by culturing the cells in medium containing 0.5% fetal calf serum for at least 48 h. To obtain S phase-enriched cultures, cell cycle reentry was induced after starvation by adding 15% fetal calf serum and harvesting 15 h after stimulation. Growth arrest and cell cycle progression were monitored by flow cytometry as described (9). Cell samples were analyzed in a FACStar Plus cytofluorimeter using the WinMDI software (10.000 events/sample).

Protein Extract Preparation and Fractionation-- Protein extracts were prepared from both NIH/3T3 fibroblast cultures and from livers from 3-4-week-old black C57 mice after nuclei isolation as described previously (8). All buffers and solutions contained 1 mM phenylmethylsulfonyl fluoride, 0.5 mM dithiothreitol, and 50 µg/ml antipain, leupeptin, chymostatin, and pepstatin A. For chromatographic fractionation, NIH/3T3 nuclei were isolated by centrifugation through a 0.8 M sucrose cushion overlaid by 0.3 M sucrose solution. Proteins were extracted in 0.42 M KCl buffer and sequentially precipitated; pelleted proteins from the 30% ammonium sulfate precipitation were redissolved in D buffer (20 mM Hepes, pH 7.9, 20 mM KCl, 2 mM MgCl₂, 0.2 mM EDTA, 20% glycerol), dialyzed, and again precipitated using 60% ammonium sulfate. Proteins in the supernatant were loaded onto a phosphocellulose column and subjected to chromatography. Recovered fractions were extensively dialyzed against D buffer, subjected to chromatography through DEAE 52 cellulose, and finally recovered by step elution using KCl.

Gel Shift Assays-- Oligonucleotides were synthesized at Genenco Service (Department of Genetics and Molecular Biology, University "La Sapienza," Rome), except for the RXR-binding oligonucleotide (DR-1 configuration) and its mutagenized version (MDR-1), which were from Santa Cruz Biotechnology. Oligonucleotide sequences are indicated in the text. Gel shift reactions were set up with 20-100 pg of [³²P]ATP-labeled oligonucleotide, 5-7 µg of NIH/3T3 cell or liver extract, 1 µg of poly(dI-dC)·poly(dI-dC), 25 mM Hepes (pH 7.6), 10% glycerol, 1 mM dithiothreitol, 0.5 mM EDTA, and 50 mM KCl for 15 min on ice. Slighlty different conditions were used for ssG binding: 1 µg of poly(dI-dC)·poly(dI-dC), 500 ng of nonspecific single-stranded DNA, 20 mM Hepes, pH 7.9, 10% glycerol, 1 mM dithiothreitol, 1 mM EDTA, 0.1 mM EGTA, 60 mM KCl and 0.1% Nonidet-P 40. In certain experiments, sodium deoxycholate (0.8% final concentration) was added either to the protein mixture before addition of the probe, or to the complete reaction, and incubated for 10 min on ice; incubations were terminated by adding Nonidet-P 40 (1.5% final concentration). Where required reactions were preincubated with competitor DNA for 10 min on ice before adding the probe. Supershift reactions were set up as above, except that 2 µl (2 µg) of anti-RXR or anti-retinoic acid receptors (RAR) antibody were added to the binding mixtures and further incubated for 45 min at room temperature. Anti-RXR and anti-RAR specificities are described below.

Western Blot Assays-- Nuclear protein extracts from cycling and growth-arrested NIH/3T3 cells, as well as from liver fractions, were electrophoresed through 10% SDS-polyacrylamide gel, electroblotted in 48 mM Tris-HCl, pH 8.3, 39 mM glycine, 0.037% SDS, and 20% methanol on nitrocellulose membranes and analyzed by Western blotting using either anti-RXR or anti-RAR antibodies (1:1000 dilution). The anti-RXR (N 197, Santa Cruz sc-774) reacts with all murine RXR subtypes (RXR, RXR, and RXR), but not with RAR factors; the anti-RAR (M-454, Santa Cruz sc-773) reacts with all RAR subtypes (₁, ₂, ₁, ₂, ₁, and ₂) but not with RXR factors. Membranes were incubated with horseradish peroxidase-conjugated secondary antibody (1:20.000 dilution), and bands were revealed following the enhanced chemiluminescence protocol (ECL reagents, Amersham Corp.).

Plasmid Constructs-- The pEA-A and pEA-C constructs were generated by inserting the EarI-AluI fragment from the Htf9 promoter (X05830 sequence) in both orientations upstream of the chloramphenicol acetyltransferase (CAT) sequence; the pES-A and pES-C constructs were generated by inserting the EarI-SmaI fragment from the Htf9 promoter in both orientations upstream of the CAT sequence. HFE-mutated constructs carried an oligonucleotide identical to the EarI-AluI fragment, except that HFE was mutated to 5-GAACTCTGATCTCTGGTCTCAGC-3 and 5-GCTGAGACCAGAGATCAGAGTTC-3 on complementary strands (C  T and G  A mutations are underlined). The fragment was cloned in both orientations upstream of the CAT sequence, yielding the pEA-mA and pEA-mC constructs respectively, or ligated to the AluI-SmaI fragment from the Htf9 promoter to yield the pES-mA and pES-mC constructs. In an independent series of constructs, the HFE oligonucleotide was ligated in multiple copies downstream or upstream of the pA10.CAT2 promoter, which carries two copies of the SV40 21-bp repeat (including Sp1 sites) and a TATA box (16).

Transfections-- NIH/3T3 cells were routinely diluted 1:6 the day before transfection. Typical experiments were carried out using 5 × 10⁶ cells and a mixture composed of DOTAP liposome reagent (Boehringer Mannheim), 5 µg of CAT reporter DNA, and 1 µg of pCMV-lacZ plasmid. The medium was changed 6 h after lipofection, and cells were harvested 36-48 h later. In all experiments, mock-transfected cultures were harvested and analyzed by flow cytometry as a control to verify that cells were actively proliferating. Promoter strengths were quantified by immunoenzymatic staining of the CAT protein (CAT enzyme-linked immunosorbent assay kit, Boehringer Mannheim) and normalized relative to the amount of synthesized -galactosidase from the co-transfected plasmid (-galactosidase enzyme-linked immunosorbent assay, Boehringer Mannheim). pSV0 or pCAT promoterless vectors (Promega) were used as negative controls. Four to eight transfections experiments were carried out for each construct.

Northern Blots-- NIH/3T3 cultures were growth-arrested or synchronized in S phase as described above. Total RNA was extracted following the guanidine-acid phenol method, electrophoresed, stained with ethidium bromide to visualize the 28 and 18 S ribosomal bands, and transferred to GeneScreen membranes as reported in detail elsewhere (9). Probes used for Northern hybridizations were gel-purified fragments corresponding to the Htf9-c, Htf9-a/RanBP1, and glyceraldehyde-3-phosphate dehydrogenase coding sequences.

S1 Analysis of Plasmids and Nuclear Chromatin-- Plasmid DNAs were prepared using Qiagen columns, which yield approximately 75% of all DNA molecules in the supercoiled form. Plasmids were subjected to S1 digestion in S1 buffer (3 mM ZnCl₂, 30 mM sodium acetate, pH 4.5, 30 mM NaCl, and 0.2 mM EDTA) for 30 min at 37 °C. Nuclei from growth-arrested or S phase NIH/3T3 cells were resuspended in buffer A (homogenization buffer without sucrose) and digested with increasing amounts of S1 nuclease (Amersham) in S1 buffer as above, except that 300 mM NaCl was used. Digestions were stopped by adding 10 mM EDTA, 10 mM Tris-HCl, pH 7.6, 0.1% SDS, and 0.1 mg/ml proteinase K at 37 °C for 5 h. Genomic DNA was repeatedly extracted with phenol and phenol-chloroform, precipitated, resuspended, digested with EcoRI, electrophoresed, and blotted with conventional methods. Terminal probes for indirect end-labeling experiments were purified from the pL9.2 subclone, which contains the Htf9 bidirectional promoter (X05830 sequence); probes were prepared by double digestion either with EcoRI and HindIII, yielding a 315-bp probe from the 3 region, or with EcoRI and EcoRV, yielding a 485-bp probe from the 5 region of the Htf9 insert, eluted from preparative gels, uniformly labeled, hybridized to the filter-bound DNA, and washed using standard procedures.

	RESULTS

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Mutations within the Footprinted Site Flanking TS-1 Impair Promoter Activity-- In a previous characterization of the Htf9 locus (shown in Fig. 1A), we mapped the full-length bidirectional promoter to a 273-bp fragment (14), which was subsequently found to contain both basal and cell cycle control elements in each orientation. Deletion of the elements responsible for cell cycle control of each gene yielded a 74-bp fragment, called EA to design the EarI-AluI restriction ends (see Fig. 1B), which directed basal transcription in both directions (7). Genomic footprinting identified functional sites of protein interaction in the fragment (15). Among those, a footprint (framed in Fig. 1B) exactly starts at the nucleotide adjacent to the major start site of transcription (TS-1 in Fig. 1B) used for initiation of both the Htf9-a/RanBP1 and Htf9-c mRNA transcripts. We set out to establish whether the sequence defined by the footprint, called Htf9 footprinted element (HFE), identified a functional element.

View larger version (30K): [in this window] [in a new window]   Fig. 1.   The Htf9 bidirectional promoter. A, map of the murine Htf9 locus, showing the 5 ends of the Htf9-a/RanBP1 and Htf9-c genes, the major transcription start site of both genes (vertical arrows), coding sequences (filled boxes), untranslated regions (empty boxes), and introns (lines). The start codon in each open reading frame is marked and the orientation of transcription is arrowed. B, sequence of the Htf9 promoter in the RanBP1 orientation. Footprinted sites are underlined; identified footprinting factors are indicated above the sequence. The HFE footprint is framed. The EarI, SmaI, and AluI sites used in promoter reporter construction are indicated. C, promoter regions assayed in transient experiments; dotted lines represent deleted regions. Mutations within HFE are represented by the crossed box. The graph on the right shows the relativity activity of constructs; values were calculated in pg of CAT enzyme/100 µg of protein extract and normalized relative to the amount of -galactosidase from a cotransfected plasmid. Mean values and S.D. were calculated from four to eight experiments for each construct. pSV0 is the empty vector.

Sequence Requirements for Protein Binding to HFE-- As an initial step to characterize the factors conferring the protection flanking TS-1, an oligonucleotide was synthesized from the footprinted HFE window and incubated with NIH/3T3 cell extracts. A discrete nucleoprotein complex was detected by gel-shift assays (Fig. 2B, lanes 1, 11, and 14). Since the HFE location near the transcription start is similar to that of certain initiators, competition assays were designed with oligonucleotides characterized for their ability to initiate transcription in TATA-less promoters, including the adenovirus Inr initiator element and binding sites for the YY1, USF, and Sp1 factors (see Weis and Reinberg (13) and references therein). A canonical TATA box was also assayed, since transcription initiation involves direct or indirect interaction of the TBP factor with several TATA-less promoters (17-19). None of the tested sequences interfered with HFE complex assembly; the complex was also insensitive to the addition of anti-TFIID antibodies and failed to interact with purified recombinant TBP protein (data not shown). Thus, the HFE complex does neither include TBP nor characterized initiator-binding proteins.

View larger version (48K): [in this window] [in a new window]   Fig. 2.   The HFE site. A, HFE sequence and mutagenized versions; only the strand corresponding to the RanBP1 orientation is shown. Repeats in the wild-type site (WT) are underlined, base substitutions are indicated by asterisks. B, gel shift assays of wild-type HFE with 5 µg of NIH/3T3 extract (lanes 1, 11, and 14) and increasing amounts of wild-type (lanes 2 and 3) or mutated (lanes 4-10) oligonucleotides (50- and 100-fold excess of competitor were used in each set of reactions), after protein extract preincubation with sodium deoxycholate (NaDOC) (lane 12), and after NaDOC addition to the assembled complex (lane 13).

HFE Is Recognized by Members of the RXR Family-- The repeated structure and sequence of HFE are similar to those contained in gene promoters regulated by retinoic acid, known as retinoic acid response elements, that are composed of a direct repetition of the PuGGTCA motif and are recognized by members of two large families of transcription factors, RARs and RXRs. Searching transcription factor data bases with the HFE sequence depicted the highest degree of raw homology with the H2RII element in the major histocompatibility complex class I gene promoter (Fig. 3A), a target of the RXR subtype of retinoid X receptors (20). Unlabeled H2RII interfered with the HFE complex assembly (Fig. 3A, lanes 6-9) almost as effectively as the homologous competitor (Fig. 3, A, lanes 2-5, and C). Reciprocal experiments using labeled H2RII as the probe showed that homologous competitions were highly effective (Fig. 3A, lanes 10-14), while competition by unlabeled HFE required a 50-fold excess to achieve 50% inhibition and plateaued thereafter (Fig. 3, A, lanes 15-18, and B). The finding that H2RII was an effective competitor of HFE, while HFE only partially competed for factors binding to the H2RII probe, suggests that H2RII and HFE are bound by both common and specific partners. We next used a canonical RXR-binding oligonucleotide in which PuGGTCA repeats are separated by one-nucleotide spacer (DR-1 configuration), as in HFE; extensive characterization of DR-1 had shown it to be the preferred target sequence of RXR factors (21). Heterologous competitions for factors between DR-1 and HFE were now complete (Fig. 3D); thus, the binding properties of HFE are indistinguishable from those of the DR-1 oligonucleotide. Together, these results implicate RXR factors in binding to HFE.

View larger version (45K): [in this window] [in a new window]   Fig. 3.   HFE competes with RXR-binding sites. A, HFE and major histocompatibility complex (MHC)-H2RII sequences are shown at the top; single repeats are framed. Below, competition assays of HFE with 5 µg of NIH/3T3 cell extract (lane 1) and increasing amounts of unlabeled HFE (lanes 2-5) or H2RII site (lanes 6-9), and reciprocal assays using the H2RII probe (lane 10) with increasing amounts of unlabeled H2RII (lanes 11-14) or HFE (lanes 15-18) DNA. B, the graph shows the ratio of complexed/total input DNA calculated from microdensitometric scanning of H2RII gel shifts in the presence of the indicated amounts of homologous or heterologous competitor. C, complexed/total input DNA ratios calculated after microdensitometric scanning of HFE gel-shifts in the presence of the indicated amounts of homologous or heterologous competitor. D, top, HFE and DR-1 sequences; repeats are framed. Below, competitive inhibition of both the HFE (lane 1) and DR-1 (lane 6) complexes formed with NIH/3T3 extracts (5 µg) by preincubation with a 50-fold excess of unlabeled HFE (lanes 2 and 7) or DR-1 (lanes 3 and 8) sites, but not with mutagenized versions of either HFE (M3, lanes 4 and 9) or DR-1 (MDR1, lanes 5 and 10).

Distinct RXR-containing Complexes Are Assembled with Extracts from Liver and NIH/3T3 Fibroblasts-- Since in previous work the strongest HFE in vitro footprints were detected using mouse liver extracts (8), the study of HFE-binding factors was further pursued using such extracts. We found that the HFE complex with liver nuclear extracts had higher abundance, and lower electrophoretic mobility, than seen using NIH/3T3 extracts (Fig. 4, compare lanes 1 and 7). Addition of an anti-RXR antibody recognizing all three RXR subtypes (, , and ) to the HFE reaction with liver extracts supershifted the complex in a manner comparable to that seen with DR-1 (Fig. 4, lanes 2 and 5). An antibody recognizing all members (₁, ₂, ₁, ₂, ₁, and ₂) of the RAR family did neither affect the HFE nor the DR-1 complex (Fig. 4, lanes 3 and 6). When immunoassays were carried out using the same amount of NIH/3T3 nuclear extract, no clear supershift was detected with either HFE or DR-1 probes; however, the amount of assembled complex with both probes decreased with the addition of both anti-RXR (Fig. 4, lanes 8, 9, 13, and 14) and anti-RAR (Fig. 4, lanes 10, 11, 15, and 16) antibodies, suggesting that component(s) of both the RXR and RAR families interacted with HFE and DR-1 in NIH/3T3 cells. Thus, the differences in abundance, mobility, and antibody reactivity indicate that specific complexes are formed by both probes depending on the extract source.

View larger version (105K): [in this window] [in a new window]   Fig. 4.   Double-stranded HFE forms distinct complexes in different cell types. The HFE (lane 1) and DR-1 (lane 4) complexes formed with liver extracts are supershifted (arrow) by anti-RXR (lanes 2 and 5) but not anti-RAR (lanes 3 and 6) antibodies; neither HFE (lane 7) nor DR-1 (lane 12) complexes formed with NIH/3T3 extracts are visibly supershifted, yet decrease, with increasing amounts of either anti-RXR (lanes 8, 9, 13, and 14) or anti-RAR (lanes 10, 11, 15, and 16) antibodies. Reactions were set up using 105 cpm of labeled probes and 6 µg of nuclear extract from either source.

View larger version (28K): [in this window] [in a new window]   Fig. 5.   Cofractionation of the HFE-binding activity and RXR factors from liver extracts. A, schematic of the procedure used for fractionation of the HFE-binding activity from liver extracts. AS, ammonium sulfate. B, gel-shift assays of HFE with eluted fractions (indicated above each lane). 4 µg of protein from nuclear extract (NE) and phosphocellulose fractions (PC), and 2 µg of DEAE 52 (DE) fractions were used. The addition of homologous competitor DNA is indicated by +. C, Western blot assays of protein from liver fractions and from growth-arrested or cycling NIH/3T3 cells with anti-RXR (top panel) and anti-RAR (lower panel) antibodies.

Distinct Proteins Bind to Each Strand of HFE in NIH/3T3 Fibroblasts-- Since the region surrounding the transcription start of both Htf9 genes can be expected to open up during transcription and expose single-stranded DNA templates to transcriptional complexes, we wondered whether single DNA strands interacted with proteins. The G-rich and C-rich strands of the HFE oligonucleotide were separately incubated with NIH/3T3 extracts and their ability to form nucleoprotein complexes was assessed in gel-shift assays. All forms of the site (i.e. G-rich, C-rich, and double-stranded) assembled discrete complexes (Fig. 6A). The complex formed with the C-rich strand was designed ssC (single-stranded C-rich DNA-containing complex). The G-rich strand formed two predominant complexes, ssG1 and ssG2 (single-stranded G-rich DNA-containing complexes 1 and 2) and two fainter complexes of lower mobility. The specificity of the complexes was assessed in competition experiments. The G-rich strand was firstly examined: the assays in Fig. 6B (lanes 1-10) show that none of the competitor DNAs inhibited the faint complexes, which therefore reflect nonspecific associations. Both ssG1 and ssG2 were instead inhibited by preincubation with homologous (lanes 3 and 4), but not with double-stranded (lanes 8-10), DNA. The complementary strand partially competed for protein(s) (lanes 5-7); however, the excess of C-rich DNA required to inhibit ssG1 and ssG2 formation was at least 5-fold higher than that used in the homologous competition (compare lane 3 to 5, and 4 to 6). Competition experiments were then set up between the C-rich strand and homologous, reverse complementary or double-stranded DNA: ssC assembly was inhibited by preincubation with homologous DNA (lanes 13 and 14), while remaining unaffected in the presence of double-stranded (lanes 18-20) HFE. Precompetition with G-rich DNA affected the formation of ssC in a dose-dependent manner (lanes 15-17); however, the inhibition remained partial even with the largest competitor excess (200-fold). To further establish whether double- and single-stranded complexes were independently assembled, HFE and the C-rich strand were simultaneously assayed. As shown in Fig. 6C, double-stranded competitor DNA selectively inhibited the HFE complex (lanes 3-5), while the C-rich strand competed for only ssC protein(s) (lanes 6-8), thus confirming the specificity of the binding events. Conclusive evidence that the factors binding to HFE and to each DNA strand are chemically distinct was obtained from NIH/3T3 extract fractionation. While HFE-binding activities were eluted in the 0.5 M KCl fraction, ssG and ssC proteins fractionated with the phosphocellulose flow-through (data not shown). Thus, distinct proteins bind to HFE and interact either with each DNA strand, or with the double-stranded form, of the element.

View larger version (39K): [in this window] [in a new window]   Fig. 6.   Distinct factors bind the double-stranded and single-stranded HFE oligonucleotides. A, gel shift assays with NIH/3T3 extracts and the G-rich (lane 2), C-rich (lane 4), or double-stranded (lane 6) probes; lanes 1, 3, and 5, probes alone. B, competition assays between single-stranded oligonucleotides. Lanes 1-10, gel shift of the G-rich strand alone (lane 1), after incubation with NIH/3T3 extracts (lane 2), and in the presence of 50- and 100-fold molar excess of homologous strand (lanes 3 and 4), 50-, 100-, and 200-fold excess of C-rich strand (lanes 5-7), and 50-, 100-, and 200-fold double-stranded HFE (lanes 8-10). Lanes 11-20, C-rich probe alone (lane 11), after incubation with NIH/3T3 extracts (lane 12), or in the presence of 50- and 100-fold excess of homologous strand (lanes 13 and 14), 50-, 100-, and 200-fold excess of G-rich strand (lanes 15-17) and 50-, 100-, and 200-fold double-stranded HFE (lanes 18-20). C, the C-rich and double-stranded HFE probes were mixed (lane 1) and incubated with NIH/3T3 extracts (lane 2), in the presence of 50-, 100-, and 200-fold double-stranded (lanes 3-5), C-rich (lanes 6-8), or G-rich (lanes 9-11) DNA.

View larger version (28K): [in this window] [in a new window]   Fig. 7.   Specificity of single-stranded binding activities. A, gel shifts of the C-rich probe with NIH/3T3 extracts (lanes 1 and 13), in the presence of a 50- and 100-fold excess of unlabeled probe (lanes 2 and 3) or 100-fold excess of the indicated competitors (lanes 4-12). B, gel shifts of the G-rich probe with NIH/3T3 extracts (lanes 1 and 13), in the presence of 50- and 100-fold excess of unlabeled probe (lanes 5 and 6), or with a 100-fold excess of the indicated competitors (lanes 2-4 and 7-12). Competitor sequences are shown in Table I.

Different Forms of the HFE Show Differential Interactions with Protein Factors in Different Cell Types-- We next asked whether the distribution of single-stranded DNA-binding factors varied in different cells, as observed for RXR and RAR binding to double-stranded HFE. Gel shift assays were carried out with probes corresponding to C-rich, G-rich, and double-stranded HFE using nuclear extracts from liver, or from cycling and growth-arrested NIH/3T3 cells. Consistent with the cell-type distribution of RXR factors seen in Western experiments, the double-stranded HFE complex was more abundant in growth-arrested, compared with cycling, NIH/3T3 cells (Fig. 8, lanes 2 and 3), and highest in liver (Fig. 8, lane 1). G-rich DNA-binding factors showed the reverse pattern, being either more abundant or more active, in cycling compared with quiescent NIH/3T3 cells, and undetectable in liver (Fig. 8, lanes 7-9). The ssC complex was comparatively more widespread, although somewhat less abundant in liver compared with both arrested and proliferating NIH/3T3 cells (Fig. 8, lanes 4-6). Together, the results suggest that different cell types are equipped with distinct sets of factors capable of binding either double-stranded HFE or single DNA strands. Single-stranded complexes most efficiently form with protein extracts from proliferating, i.e. actively transcribing, NIH/3T3 fibroblasts, whereas factors interacting with double-stranded HFE were most abundantly expressed in liver.

View larger version (59K): [in this window] [in a new window]   Fig. 8.   Distribution of factors binding different HFE forms. Double-stranded HFE (lanes 1-3), C-rich (lanes 4-6), and G-rich (lanes 7-9) oligonucleotides were incubated with protein extracts (4 µg) from liver (lanes 1, 4, and 7), growth-arrested (lanes 2, 5, and 8) and cycling (lanes 3, 6, and 9) NIH/3T3 cells.

Plasmids Containing the HFE Sequence Are Sensitive to S1-- Since single-stranded DNA-binding activities were depicted in cell cultures in which the Htf9 locus is transcriptionally active (9), it was important to assess whether single-stranded structures actually formed in the Htf9 promoter. Computer analysis revealed that HFE fell within one potential stem-and-loop structure; theoretical estimates of the free energy associated with the formation of the single-stranded loop give G = 21.7 kcal/mole. We wondered whether these theoretical sequence features gave rise to the formation of unusual DNA structures in the Htf9 initiation region. Supercoiled plasmids carrying either the full-length (pTS-A, 273 bp), or the minimal (pEA-A, 74 bp), Htf9 promoter were digested with S1 nuclease and subsequently restricted with PstI to separate the Htf9 sequences from the plasmid replication origin (Fig. 9A), which might contain unwound DNA. Results in Fig. 9B show that both the pTS-A and pEA-A constructs contained S1-sensitive sequences that were not present in the vector. Hybridization with labeled HFE oligonucleotide confirmed that the region containing HFE was sensitive to S1 cleavage (Fig. 9C). To directly ascertain whether HFE in particular represented a DNA target for S1 cleavage, pA10-derived clones carrying one or three HFE copies (Fig. 9D) were subjected to digestion using increasing S1 nuclease doses. While the pA10 vector was inefficiently cleaved by S1 (Fig. 9E, lanes 1-4), plasmids containing one or more HFE copies were sensitive to S1 in a dose-dependent manner. Microdensitometry quantification of the ratio of supercoiled to linear DNA revealed that the extent of cleavage by S1 increased with the number of HFE copies. Thus, the HFE sequence confers S1 sensitivity to supercoiled plasmids.

View larger version (35K): [in this window] [in a new window]   Fig. 9.   S1 sensitivity in HFE-containing plasmids. A, maps of the pSV0-CAT vector and of Htf9-derived promoter constructs, respectively, carrying the basal (pEA-CAT, 74 bp) and the full-length (pTSA-CAT, 273 bp) promoter region. The pBR322 origin of replication (ORI) is marked. P, PstI; H, HindIII. In the series of constructs, the promoter insert (cloned in the HindIII site) is oriented in the direction of RanBP1 transcription relative to the CAT gene. B, ethidium-bromide staining of constructs subjected to digestion with S1 nuclease (lanes 2, 4, and 6) or to mock digestion (indicated by a dash (-) in lanes 1, 3, and 5), and subsequently restricted with PstI to separate the replication origin from the Htf9 promoter region. Lane 7,  DNA digested with EcoRI and HindIII. C, autoradiograph of the blot from the gel in A after hybridization with labeled HFE oligonucleotide. D, chimeric promoter constructs carrying no (pA10), one (pA10G), or three (pA10G9) HFE copies upstream of the pA10 minimal promoter. The arrow marks the promoter insertion site. E, ethidium bromide-stained gel showing 1 µg of DNA from each construct after digestion with 0, 0.5, 1, and 2.5 units of S1.

The Genomic Region Surrounding HFE Is Sensitive to S1 during Transcription in Vivo-- We finally assessed whether S1-sensitive structures identified in HFE-containing clones were maintained in the higher order organization of genomic DNA. NIH/3T3 cell cultures were either brought to proliferation arrest by serum withdrawal for 48 h, or stimulated to reenter the cycle by adding fresh serum. Flow cytometric analysis confirmed that cells collected prior to serum stimulation were arrested in the G₀/G₁ state, while cells collected 15 h after serum refeeding were traversing S phase (Fig. 10A). Neither Htf9-associated gene was transcribed in growth-arrested cells, while transcription was active in S phase cells (Fig. 10B), confirming our previous data on cell cycle control of the Htf9 promoter (7). Nuclei were isolated from both growth-arrested and S phase NIH/3T3 cultures to assess the Htf9 promoter sensitivity to S1 in transcribing and nontranscribing cells. Nuclei were digested with increasing amounts of S1 nuclease; DNA was extracted, restricted with EcoRI and hybridized with a Htf9-derived fragment that was flush with one EcoRI restriction end (map in Fig. 10C). These experiments depicted a restricted region (approximately 100 bp) that was sensitive to S1 cleavage in S phase NIH/3T3 cells (Fig. 10C, lanes 5-7), but not in NIH/3T3 cultures brought to quiescence (Fig. 10C, lanes 1-3). Indirect end-labeling revealed that the S1-sensitive region in S phase cells encompassed the 74-bp region sufficient for basal transcription as functionally defined in promoter assays (see Fig. 1).

View larger version (26K): [in this window] [in a new window]   Fig. 10.   S1 sensitivity of the HFE-containing region in S phase and in growth-arrested cells. A, flow cytometry analysis of cell cultures maintained in low serum for 48 h (top) and after 15 h of serum restimulation (below). The DNA content of cells is indicated on the x axis, cell numbers are indicated on the y axis. B, total RNA (15 µg) was extracted from growth-arrested (G0, lane 1) or S phase (lane 2) NIH/3T3 cultures, stained with ethidium bromide to assess the integrity of the 28S and 18S ribosomal bands (top panels), blotted, and hybridized with the indicated probes. C, nuclei from growth-arrested (G0) and S phase NIH/3T3 cells were digested with the following S1 units/µg DNA: 1, lanes 1 and 5; 5, lanes 2 and 6; 10, lanes 3 and 7. DNA was extracted, restricted with EcoRI, and hybridized with a fragment flush with one EcoRI end (shown on the map). Lane 5,  EcoRI-HindIII marker fragments (M). The full-length restriction fragment is indicated by a thick arrowhead; fragments derived from internal S1 cleavage (indicated by thin arrowheads) were detected in S phase, but not in G0, NIH/3T3 nuclei. The region of S1 cleavage (arrowheads) encompasses the minimal promoter region (box).

	DISCUSSION

Top Abstract Introduction Procedures Results Discussion References

The HFE Near TS-1 Is Required for Promoter Activity-- A 74-bp promoter region carries sufficient information to direct transcription of both the RanBP1 and Htf9-c genes. We have presently began to characterize the HFE element, originally identified by genomic footprinting, which flanks the TS-1 start site used for initiation of both genes. Promoter sites showing interactions with factors in vivo often identify regulatory elements. HFE is indeed a functional element in both orientations, as mutational inactivation reduces promoter strength in each direction. However, HFE inactivation is sensed differently depending on the orientation and arrangement of neighboring regulatory sequences in the promoter context; HFE inactivation does mildly impair the pES-C promoter, while severely affecting activity of both the pES-A and pEA-C constructs. The effectiveness of regulatory sequences depends, at least in part, on their position in the promoter context. It is possible that regulatory elements included between the pEA-C and pES-C promoter boundaries participate in the assembly of transcriptional complexes and relieve in part the effect of HFE inactivation. Thus, the functional promoter mapping results suggest that the HFE site is involved in mediating interactions among various regulatory elements in each promoter orientation.

RXR Factors Bind the Double-stranded HFE-- In the search for factor(s) protecting HFE, we have identified distinct activities which bind different HFE forms. The double-stranded site is recognized by complexes containing retinoid X receptors. These complexes showed cell type-specific differences in both their relative abundance and interacting partners. NIH/3T3 extracts yielded a complex that was similarly reacted by both anti-RXR and anti-RAR antibodies, while liver extracts formed a complex that was specifically reacted by anti-RXR but not anti-RAR antibodies. Thus, RXRs interacting with HFE associate with specific partners in different cells. These findings were paralleled by the distribution of retinoid receptors: in liver nuclei, all three RXR subtypes were abundant and were recovered in chromatographic fractions positive for HFE binding. In NIH/3T3 cells, RAR factors were also depicted, while only one RXR subtype was above the level of detection in Western experiments.

Specific Factors Bind Each DNA Strand of the HFE-- One unexpected HFE feature is that specific activities present in NIH/3T3 cells bind to each DNA strand. That finding prompted us to investigate whether single-stranded structures actually formed in the Htf9 promoter. Theoretical energy estimates support the possibility that the Htf9 promoter forms a stem-and-loop structure, and the identification of S1-sensitive sites in HFE-containing plasmids suggests that the formation of single-stranded structures represent an inherent feature of the Htf9 initiation region. Many CG-rich, TATA-less promoters potentially form single-stranded loops in the region surrounding the transcription start sites (32). Such looping structures have been suggested to serve as structural landmarks and facilitate the recognition of start sites by basal factors in complex genomes, where TATA box recognition by linear sequence scanning would be inefficient compared with smaller genomes (32).