Activation of the Human Asparagine Synthetase Gene by the Amino ...

THE JOURNAL OF BIOLOGICAL CHEMISTRY © 2000 by The American Society for Biochemistry and Molecular Biology, Inc.

Vol. 275, No. 35, Issue of September 1, pp. 26976 –26985, 2000 Printed in U.S.A.

Activation of the Human Asparagine Synthetase Gene by the Amino Acid Response and the Endoplasmic Reticulum Stress Response Pathways Occurs by Common Genomic Elements* Received for publication, January 4, 2000, and in revised form, June 14, 2000 Published, JBC Papers in Press, June 15, 2000, DOI 10.1074/jbc.M000004200

Ione P. Barbosa-Tessmann‡, Chin Chen‡, Can Zhong‡, Fai Siu‡, Sheldon M. Schuster‡, Harry S. Nick§, and Michael S. Kilberg‡¶ From the ‡Department of Biochemistry and Molecular Biology and the §Department of Neuroscience, University of Florida College of Medicine, Gainesville, Florida 32610

The human asparagine synthetase (AS) gene is transcriptionally regulated by amino acid deprivation (amino acid response, AAR) and the endoplasmic reticulum stress response (ERSR), also known as the unfolded protein response pathway. The results reported here document the novel observation that induction of the AS gene by the AAR and ERSR pathways occurs via the same set of genomic elements. Data supporting this conclusion include transient transfection of AS promoter/ reporter gene constructs that illustrate that the transcriptional control elements used by both pathways are contained with nucleotides ⴚ111 to ⴚ34 of the AS promoter. In vivo footprinting analysis of this region identified six specific protein-binding sites. Within two of these sites, altered footprinting was observed following amino acid or glucose deprivation, but the patterns were identical for both the AAR and the ERSR pathway. Site-directed mutation of individual nucleotides within these two binding sites confirmed their importance for regulated transcription, and none of the mutations resulted in loss of response of only one pathway. Neither of these two sites corresponds to a recently identified ERSR cis-element, nor do they contain consensus sequences for known transcription factors. Collectively, the data document that there are at least two independent transcriptional mechanisms for gene activation by the ERSR pathway, one of which terminates at the same genomic elements used by the AAR pathway.

The control of transcription by cellular metabolites is a mechanism that allows cells to respond to changes in their nutritional environment. For example, depriving cells of glucose results in enhanced transcription for the glucose-regulated proteins (GRP)1 GRP78 and GRP94 (1) and the transcription * This work was supported by NIDDK National Institutes of Health Grant DK-52064 (to M. S. K.). The costs of publication of this article were defrayed in part by the payment of page charges. This article must therefore be hereby marked “advertisement” in accordance with 18 U.S.C. Section 1734 solely to indicate this fact. ¶ To whom correspondence should be addressed: Dept. of Biochemistry and Molecular Biology, University of Florida College of Medicine, Box 100245, JHMHC, Gainesville, FL 32610-0245. Tel.: 352-392-2711; Fax: 352-392-6511; E-mail: [email protected]. 1 The abbreviations used are: GRP, glucose-regulated protein; ER, endoplasmic reticulum; ERSR(E), endoplasmic reticulum stress response (element); L7a, ribosomal protein L7a; CHOP, C/EBP homology protein; gadd153, growth arrest and DNA damage protein 153; nt, nucleotide(s); AS, asparagine synthetase; AAR, amino acid response; bp, base pair(s); MEM, minimal essential medium; GH, growth hormone; MTT, metallothionein; TK, thymidine kinase; EMSA, electrophoretic mobility shift assay; GDH, glutamate dehydrogenase.

factor CHOP/GADD153 (2). Transcription of these genes also is up-regulated by a variety of factors that have in common disruption of endoplasmic reticulum (ER) function and accumulation of misfolded proteins (3, 4). This activation of transcription is mediated by a signal transduction pathway referred to as the ER stress response (ERSR) in mammalian cells, also called the unfolded protein response in yeast. Examples of ERSR pathway activators include depletion of calcium stores, inhibition of N-linked glycosylation by glucose deprivation or tunicamycin treatment, or amino acid analogs that incorporate into proteins and cause misfolding (3, 4). It is important to note that amino acid deprivation, in contrast to the presence of incorporated amino acid analogs, does not induce ERSR-activated genes such as GRP78 (5), presumably because slowing of protein synthesis does not result in a significant accumulation of misfolded proteins within the ER. However, there are a number of genes for which transcription is enhanced following amino acid limitation (6). For example, there is considerable evidence for transcriptional induction of asparagine synthetase (AS) by amino acid deprivation (7–10), a process that will be referred to as the amino acid response (AAR) pathway. In mammalian cells, most of the data regarding ERSR-mediating cis-elements arose from sequence analysis of the 5⬘ upstream genomic regions for GRP78, GRP94, calreticulin, and protein-disulfide isomerase. This sequence comparison revealed that all of these genes contain multiple copies of the consensus sequence 5⬘-CCAAT-N9-CCAG-3⬘, with a GC-rich 9-bp spacer region (11, 12). The transcription factors CBF/ NF-Y and ATF6 are believed to bind to the CCAAT and CCACG sequences, respectively, within the ERSR element (12–14). In contrast, much less information is available regarding the genomic elements responsible for the transcriptional activation by the mammalian AAR pathway. Guerrini et al. (8) used deletion analysis and block substitution mutagenesis to analyze the proximal promoter of the human AS gene. Through those experiments they identified the palindromic sequence 5⬘-CATGATG-3⬘ at nucleotides ⫺70 to ⫺64 as a potential AAR element. However, no corresponding transcription factors have been identified for this regulatory sequence. Recently, Barbosa-Tessmann et al. (5) demonstrated that human AS gene expression was increased in response to glucose deprivation as well as amino acid deprivation. Transient transfection of a reporter gene driven by AS genomic 5⬘-upstream fragments documented that the glucose-dependent activation was transcriptional, and induction by tunicamycin treatment or by amino acid analogs illustrated that the activation was triggered by the ERSR pathway (15). The metabolic significance of the ERSR-dependent activation of the AS gene is potentially linked to the observation that for cells lacking

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Activation of the Asparagine Synthetase Gene sufficient AS activity, depletion of asparagine results in cell cycle arrest (7) and induction of apoptosis (16, 17). Interestingly, activation of the ERSR pathway also is associated with apoptosis (18, 19). The objective of the research presented here was to distinguish between the genomic elements responsible for transcriptional activation of the human AS gene in response to the AAR and ERSR pathways. Although previous deletion analysis suggested that the cis-elements for the AAR and ERSR pathways may exist in the same region of the human AS promoter (15), the initial hypothesis for the present set of experiments was that each of these pathways induced AS expression through independent cis-elements. Deletion and mutagenesis analysis of AS promoter fragment-reporter gene constructs and in vivo footprinting were used to identify important regulatory elements within the AS gene promoter. Surprisingly, these studies document the novel observation that both the AAR and ERSR pathways utilize the same set of genomic elements to activate transcription of the human AS gene. Given that these sequences do not correspond to the previously reported mammalian ERSR element consensus sequence (11, 12), the data also demonstrate that there are at least two independent transcriptional mechanisms for induction of ERSR genes and that one of those mechanisms overlaps with that used by the AAR pathway. MATERIALS AND METHODS

Cell Culture—Human hepatoma HepG2 cells were cultured in minimal essential medium (MEM), pH 7.4, supplemented with 25 mM NaHCO3, 2 mM glutamine, 10 ␮g/ml streptomycin sulfate, 100 ␮g/ml penicillin G, 28.4 ␮g/ml gentamycin, 0.023 ␮g/ml N-butyl-p-hydroxybenzoate, 0.2% (w/v) bovine serum albumin, and 10% (v/v) fetal bovine serum. Cells were maintained at 37 °C in a 5% CO2, 95% air incubator in T-75 flasks. To induce expression of the AS gene, cells were cultured to near 70 – 80% confluence, and then incubated for 12 h in either complete MEM, glucose-free-MEM, or histidine free-MEM, each supplemented with 2 mM glutamine and 10% dialyzed fetal bovine serum. Human Asparagine Synthetase Genomic Clone—A human AS genomic clone was obtained by PCR screening of a PAC genomic library (Genome Systems Inc., St. Louis, MO) using a set of primers (5⬘CAAACCAAGTTCAGAAGCCTCCC-3⬘ and 5⬘-AAGCAGGTCAGGGTGATGTGGC-3⬘) to yield a 328-bp sequence covering the proximal promoter region/exon 1 junction. Positive clones were rescreened with a second set of primers (5⬘-TGCAATGATGGCAAATGCAGCC-3⬘ and 5⬘ACTTGTAGTGGGTCAGCGTGCGG-3⬘) covering a 185-bp sequence within exon 12. Primers were designed based on published sequences of the human AS gene and cDNA (20, 21). Three independent PAC clones were obtained, each containing all 13 exons and a large portion of the 5⬘-upstream region. One of the clones was digested with XbaI, yielding a large fragment that contained a portion of exon 1 and 10.9 kbp of 5⬘-upstream sequence (⫺10, 895/⫹1, 377). The primary transcription start site (⫹1) was taken as that defined by Guerrini et al. (8). Gene Deletions—The p0GH vector contains the entire human growth hormone (GH) gene, but lacks a functional promoter (22). To define AS regulatory sequences by transient transfection, a collection of p0GH plasmids containing specific regions of the AS 5⬘-flanking region were generated. Based on the human AS genomic sequence, specific primers were designed to PCR amplify the following fragments ⫺3121/⫹51, ⫺615/⫹51, ⫺475/⫹51, ⫺376,⫹51, ⫺274/⫹51, ⫺173/⫹51, and ⫺72/⫹51 of the proximal 5⬘-upstream region of the gene. In subsequent experiments, sequential deletion constructs of the ⫺173/⫹51 fragment at both the 5⬘ and 3⬘ ends were also prepared. Primers always contained BamHI restriction sites at the 5⬘ end, to permit subsequent cloning. The PCR products were cloned using the TA cloning system from Invitrogen (Carlsbad, CA) and then subcloned into the BamHI site of the p0GH plasmid. The directionality and sequence of the inserts in the final constructs was confirmed by restriction digestion and sequencing. Site-directed Mutagenesis and Block Substitutions—Mutagenesis of specific nucleotides was accomplished by PCR as described by Ho et al. (23). All substitutions were made by replacing pyrimidine for pyrimidine and purine for purine. The PCR products were cloned into the BamHI site of the p0GH expression vector, as described above, and the orientation and integrity of each insert was checked by DNA sequencing. The AS/GH reporter plasmids containing these mutated sequences

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were transiently transfected into HepG2 cells, and the ability of each mutant sequence in promoting basal or nutrient-dependent transcription was analyzed by GH Northern blotting, as described below. Transient Transfection and Northern Analysis—HepG2 cells were transfected with the p0GH reporter vector containing no promoter (22), a 1.8-kilobase pair EcoRI-BglII fragment of the mouse metallothionein-I (MTT) gene as a control promoter that should not respond to either amino acid or glucose deprivation (22, 24), or the indicated fragments of the human AS gene. The cells were transfected at 70 – 80% confluence in 100-mm dishes using the batch transfection technique described by Barbosa-Tessmann et al. (15). After transfection and a subsequent 24-h incubation in MEM, cells were divided into four or five 60-mm dishes and cultured for another 24 h prior to incubation for 12 h in complete MEM, histidine-free MEM, or glucose-free MEM, as described in each figure legend. Therefore, within each experiment, cells incubated in the different treatment media arose from the same transfected cell population, thus eliminating differences in transfection efficiency between treatments. The ability of each promoter fragment to drive transcription of the GH reporter gene was evaluated by measuring GH mRNA content. The transfection efficiency between the initial 100-mm dishes was monitored by co-transfection with an expression vector (pcDNA3.1) containing lacZ driven by the cytomegalovirus promoter and subsequent Northern analysis for lacZ mRNA. Total cellular RNA was isolated according to the procedure described by Chomczynski and Sacchi (25). 32P-Radiolabeled cDNA probe synthesis and Northern analysis was performed as described in BarbosaTessmann et al. (5). Quantification was done by scanning the exposed film and analyzing the data using Un-Scan-It computer software (Silk Scientific Inc., Orem, UT). Relative intensity of each band obtained for the regulated genes of GH, AS, or GRP78 was normalized to the signal intensity of an mRNA for which expression did not change under the experimental conditions, either ribosomal protein L7a (L7a) for the ERSR or glutamate dehydrogenase (GDH) for the AAR. Qualitatively similar results were reproducibly obtained when the transfection experiments were repeated. Given the quantitative variation for transfection between batches of cells, the data from a single experiment is shown. The radiolabeled probe for human AS was the entire cDNA. The cDNA probe for GRP78 was a 1498-bp sequence between the PstI and EcoRI sites (26). The GH cDNA probe was a 651-bp sequence containing the entire open reading frame (22). The cDNA probe for the L7a was a 600-bp sequence obtained from Dr. Tatsuo Tanaka (University of Ryukyus, Okinawa, Japan). The cDNA probe for the GDH was the entire cDNA (obtained from Dr. W. H. Lamers, University of Amsterdam, Netherlands). In Vivo Footprinting—For the in vivo treatment with dimethyl sulfate, five 100-mm dishes of HepG2 cells (6.6 ⫻ 106 cell/dish) for each treatment were incubated for 12 h in complete MEM, histidine-free MEM, or glucose-free MEM. The cells were rinsed once with phosphatebuffered saline, and then incubated at room temperature for 30 s in phosphate-buffered saline containing 0.5% dimethyl sulfate. Cells were quickly rinsed twice with 10 –15 ml of ice-cold phosphate-buffered saline and then immediately lysed with 3 ml of lysis solution (50 mM Tris-HCl, pH 8.5, 50 mM NaCl, 25 mM EDTA, pH 8.0, 0.5% SDS, and 300 ␮g/ml proteinase K) and genomic DNA was isolated (27). As a control, genomic DNA was extracted from cells incubated in complete MEM for 12 h, but not treated with dimethyl sulfate. This “naked” DNA was restriction digested with BamHI, phenol-chloroform extracted, and then treated with 0.25% dimethyl sulfate in vitro for 30 s, as described by Maxam and Gilbert (28). A 70-␮g aliquot of the dimethyl sulfate-modified DNA (both in vivo and in vitro treated samples) was mixed with 10 ␮l of piperidine (10 M) and incubated at 90 °C for 30 min to cleave the DNA at the methylated guanine residues (28). After precipitation and three cycles of resuspension in water and lyophilization to remove piperidine, the DNA was resuspended in 20 ␮l of TE buffer. Ligation-mediated PCR was performed (27) by annealing 6 pmol of primer-1 (5⬘-CTCACCTGGGCGTAAGCAGG-3⬘ top strand, 5⬘-CAAACCAAGTTCAGAAGCCTCCC-3⬘ bottom strand) to 3 ␮g of dimethyl sulfate-modified and piperidine-cleaved DNA in 1 ⫻ Vent polymerase buffer (10 mM KCl, 10 mM (NH4)2SO4, 20 mM Tris-HCl, pH 8.8, 2 mM MgSO4, 0.1% Triton X-100), in a total volume of 15 ␮l, with denaturation at 95 °C for 10 min followed by primer annealing at 45 °C for 30 min. The first strand synthesis was performed in 1 ⫻ Vent polymerase buffer with 4 mM MgSO4, 0.25 mM of each dNTP, and 2 units of Vent polymerase using incubations of 1 min each at 53, 55, 57, 60, 62, 64, and 68 °C, followed by incubations for 3 min each at 72 and 76 °C. Following first strand synthesis, a doublestranded linker sequence was ligated to the piperidine-generated DNA

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fragments. The double-stranded linker was composed of two complementary oligonucleotides, a 25-mer (5⬘-GCGGTGACCCGGGAGATCTGAATTC-3⬘) annealed to an 11-mer (5⬘-GAATTCAGATC-3⬘), described by Mueller and Wold (29). After ligation of the linker, the DNA was phenol-chloroform extracted and the nested set of linked genomic DNA fragments from the region of interest were amplified by PCR using a second gene-specific oligonucleotide primer, termed primer-2 (5⬘GCAGGTCAGGGTGATGTGGCGGG-3⬘ top strand, 5⬘-GCAGGTAGCCTGGGCGGAGCTCTGAG-3⬘ bottom strand), and the linker-specific 25-mer (5⬘-GCGGTGACCCGGGAGATCTGAATTC-3⬘). PCR conditions, described by Hornstra and Yang (27), were the same for both DNA strands and performed with Taq DNA polymerase. The annealing temperature (68 °C) was determined experimentally to establish the conditions that yielded the best results. Samples were denatured at 95 °C for 1 min, annealed at 68 °C for 2 min, and extended at 76 °C for 3 min for 25 cycles, except that for each cycle of amplification, the extension time was increased by 5 s. After 25 cycles, samples were incubated at 76 °C for 30 min in 5 ␮l of a fresh PCR incubation solution containing 2.5 mM each dNTP, 1.75 mM MgCl2, 1 ⫻ Taq DNA polymerase buffer (20 mM Tris-HCl, pH 8.4, 50 mM KCl), and 1 unit of Taq polymerase to ensure full extension of each template. The PCR reaction was stopped by adding 1 ␮l of 0.5 EDTA and the amplified fragments were recovered by phenol-chloroform extraction, ethanol precipitation, and resuspended in 20 ␮l of water. The amplified genomic DNA fragments were separated on a 8.3 M urea, 5% Long Ranger (FMC Bioproducts, Rockland, ME) polyacrylamide DNA sequencing gel (60 cm), electrotransferred and UV cross-linked (30) to a Zetabind positively charged nylon membrane (Cuno, Meriden, CT), and then hybridized with a 32P-radiolabeled oligonucleotide (5⬘-AAACAGGCGCACTGAGACGC-3⬘ top strand, 5⬘-AAACAGGCGCACTGAGACGC-3⬘ bottom strand) positioned internally to primer-2. Hybridization conditions (at 40 °C) and washing procedures (at 45 °C) were the same as those described for Northern analysis. After densitometry of each lane of a preliminary gel, the amount of each sample loaded was adjusted to obtain equal signal intensity in all lanes. Electromobility Mobility Shift Assay—Total nuclear extracts were prepared from HepG2 cells incubated for 18 h in either complete MEM (control) or in MEM lacking histidine (⫺His) (31). Double-stranded oligonucleotide probes were 32P-radiolabeled by extension of overlapping ends with Klenow fragment in the presence of [␣-32P]dCTP (31). An aliquot of nuclear extract (5 ␮g of protein) was preincubated for 10 min at 4 °C in a total volume of 20 ␮l containing 40 mM Tris, pH 7.5, 200 mM NaCl, 2 mM dithiothreitol, 10% glycerol, 0.05% Nonidet P-40, 2 ␮g of poly(dI-dC), 0.05 mM EDTA, and with (“WT” lanes, Fig. 7) or without (“⫹c” lanes, Fig. 7) 6 ng (⫻ 100) unlabeled competitor oligonucleotide. Then 0.06 ng of 32P-radiolabeled probe (10,000 dpm) was added and the incubation continued for 20 min at room temperature. The mixture was subjected to electrophoresis on an 8% polyacrylamide gel and the results visualized by autoradiography. All experiments were repeated with at least two independently prepared nuclear extracts. RESULTS

Genomic Map—Previous genomic clones of human AS were reported by Greco et al. (20) and Zhang et al. (21), but both groups pieced together several independent phage clones to assemble the complete genomic sequence and a comparison of the two clones revealed a number of discrepancies regarding gene organization, intron-exon boundaries, and transcription start sites. Given these differences, we decided to obtain a human AS genomic clone as single piece of DNA by screening a genomic PAC library (32). Restriction digestion and Southern hybridization established that three BamHI fragments encompassed 26.4-kilobase pairs containing all 13 exons and a large region of the 5⬘-upstream region of the human AS gene. One of the BamHI fragments, containing nucleotides ⫺3121 to ⫹6618, was used to sequence the proximal 3121-bp segment of the 5⬘-upstream region (GenBank accession number AF239815). The restriction map and the 3121 bp of sequence from our genomic clone was in agreement with the corresponding region obtained from sequencing chromosome 7 as part of the Human Genome Project (GenBank accession number AC005326) and most closely agreed with that previously published by Greco et al. (20). Deletion Analysis—The following experiments were designed

to identify those genomic elements responsible for activation of the AS gene by the AAR and ERSR pathways. The initial characterization involved deletion analysis beginning with a GH reporter construct preceded by the AS fragments ⫺10,895/ ⫹1,377, ⫺3,121/⫹51, and ⫺615/⫹51. Each construct was transiently transfected into HepG2 hepatoma cells by the batch transfection protocol described above. After a 12-h incubation in MEM, MEM lacking histidine, or MEM lacking glucose, RNA was isolated and Northern analysis of GH mRNA content was taken as a measure of transcriptional activity. An increase in endogenous AS and GRP78 mRNA were used to document the activation of both the AAR and ERSR pathways, whereas GDH or ribosomal protein L7a mRNA content served as negative controls. There was little or no GH mRNA produced after transfection of the promoter-less construct p0GH, and the presence of the MTT promoter yielded constitutive expression, as expected (Control, Fig. 1, A and B). In contrast, all three AS gene fragments promoted basal transcription of the GH gene that was significantly enhanced by deprivation for either histidine (⫺His, Fig. 1, A and B) or glucose (⫺Glc, Fig. 1, C and D). Those same AS fragments, when subcloned in the reverse orientation, were unable to produce a similar enhancement of GH transcription. The 5⬘ end of the ⫺615/⫹51 fragment was further deleted to find the smallest region of the AS gene promoter that still retained AAR or ERSR activity. Surprisingly, for both the AAR (Fig. 2, A and B) and ERSR pathways (Fig. 2, C and D), all fragments tested were effective in enhancing transcription until the 5⬘ end was reduced from nt ⫺173 to ⫺72. To further investigate the possible separation of the sequences necessary for the two nutrient-dependent pathways, the ⫺173/⫹51 bp AS sequence was deleted from both the 5⬘ and 3⬘ directions. Previous experiments from our laboratory had documented that the genomic elements required for activation of the AS gene by the ERSR pathway resided between nt ⫺111 and ⫺34 (15), so only the AAR pathway was monitored. Sequential deletion of the AS 5⬘-upstream sequence from nt ⫺173 to ⫺149 actually caused an enhancement of both basal and AAR-dependent transcription (Fig. 3, A and B). Whether or not this 24-bp sequence contains a repressor element will be the subject of future experiments. Further deletion from ⫺149 to ⫺111 resulted in a moderate decline in transcriptional activity, but the value following histidine deprivation was still greater than the ⫺173/⫹51 construct (Fig. 3, A and B). However, reduction from ⫺111 to nt ⫺90 or less caused a decrease in transcription, and although the increased transcription by the AAR pathway was reduced, it was still detectable (Fig. 3, A and B). The results parallel those obtained when similar constructs were tested for activation by the ERSR pathway (15). The reduction of transcriptional activity from nt ⫺111 to ⫺90 coincides with the deletion of the last of three GC boxes, located at nt ⫺106 to ⫺97, for which the sequence is identical to the Sp1 consensus (33). The sequential 3⬘ deletions of the ⫹173/⫹51 construct from nt ⫹51 to ⫺34 resulted in a graded decline in basal transcription, but induction by amino acid deprivation still occurred (Fig. 3, C and D). Further 3⬘ deletion to nt ⫺52 resulted in little or no detectable transcription, even after nutrient deprivation. Once again, the results for activation of the AAR pathway shown here are similar to those obtained previously following induction of the ERSR pathway (15). For constructs that were deleted in the 3⬘ direction beyond nt ⫹1 of the AS sequence, transcription presumably started within the GH gene. The p0GH vector includes the entire GH gene, and the region around the GH transcription start site contains multiple Inr-like initiator sequences which, in conjunction with the multiple GC boxes of the AS promoter, probably promote transcription initiation (34).


FIG. 1. Deletion analysis of the human AS gene 5ⴕ-upstream region. HepG2 human hepatoma cells were transfected with a GH reporter construct lacking a promoter (p0GH), containing the constitutive metallothionein promoter (MTT), or the indicated sequence of the human AS gene 5⬘-upstream region. After employing the batch transfection technique, described in the text, cells were incubated for 12 h in complete MEM (Control), histidine-free MEM (⫺His), or glucose-free MEM (⫺Glc). Expression of the GH reporter mRNA was monitored by probing a Northern blot (20 ␮g of RNA per lane) with a 32P-radiolabeled GH cDNA. To confirm the expression of endogenous AS under histidine or glucose deprivation, the blots were stripped and rehybridized with an AS cDNA. To illustrate consistent lane loading, the blots were rehybridized with either GDH cDNA (panels A and B) or L7a (panels C and

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Interestingly, the deletion studies indicated that the genomic element(s) required for activation of the AS gene by either the AAR or the ERSR pathways were contained within nt ⫺111 to ⫺34 of the proximal promoter. To confirm this interpretation, the ⫺111/⫺34 AS fragment was placed in front of the GH reporter gene, in both the forward and reverse orientations. The presence of the ⫺111/⫺34 AS sequence did not result in a level of basal transcription as great as the ⫺173/⫹51 sequence (Fig. 4, A and B, Control), but the fragment was effective in promoting enhanced transcription following tunicamycin treatment (Fig. 4, C and D) or depletion for either histidine (Fig. 4, A and B) or glucose (Fig. 4, C and D) in the medium. To test whether or not the ⫺111/⫺34 AS sequence could transfer regulated expression to the thymidine kinase (TK) promoter, the fragment was cloned into the HindIII site of a TK promoter/GH reporter construct (pTKGH), which contains 200 bp of the promoter sequence from herpes simplex virus thymidine kinase in front of the human GH reporter gene (22). The TK promoter alone exhibited low activity in promoting basal transcription of the GH gene and the addition of the AS ⫺111/⫺34 sequence had little effect (Fig. 4, A and B). However, in response to activation of either the AAR (second panel of Fig. 4, A and B) or the ERSR (Fig. 4, C and D) pathway, the ⫺111/⫺34 AS gene fragment significantly increased TK-driven transcription. Interesting, in contrast to the longer fragments of the promoter (Fig. 1), this “minimal AS control unit” conferred nutrient-dependent regulation in the reverse orientation, with or without the TK promoter (Fig. 4). In Vivo Footprinting—Using dimethyl sulfate as a molecular probe, in vivo footprinting was performed to define, at single nucleotide resolution, the protein-binding sites within the first 173 nt of the AS 5⬘-flanking region. HepG2 cells, incubated for 12 h in MEM (Control) or medium lacking either histidine (Fig. 5, A and C) or glucose (Fig. 5, B and D) were then subjected to footprinting analysis. Relative to the in vitro dimethyl sulfatetreated DNA (“Naked”), numerous guanine nucleotides exhibited altered dimethyl sulfate reactivity in control cells that appeared as either a diminished (“protected”) or an enhanced hybridization signal. In addition, there were specific guanine nucleotides that exhibited additional protection (“nutrient-dependent protection”) when cells were deprived of either histidine or glucose relative to the control DNA samples from cells maintained in complete MEM. Collectively, the results have lead us to propose, as a working model, the existence of six independent protein-binding sites (numbered I-VI) that are each constitutively occupied. However, for two of these sites (sites V and VI) there are detectable increases in DNA-protein interaction in response to activation of either the AAR or the ERSR pathway. The observation that identical in vivo footprinting results were obtained in response to activation of either nutrientsensing pathway was unexpected. Binding site I had protected guanine nucleotides at positions ⫺144, ⫺142, ⫺141, ⫺140, ⫺139, and ⫺137, on the bottom strand (Fig. 5, C and D), and an enhanced guanine at position ⫺146, on the top strand (Fig. 5, A and B). Binding site II had protected guanines at nt ⫺131, ⫺130, ⫺127, ⫺124, ⫺122, ⫺121, ⫺120, and ⫺119 and an enhanced guanine at position ⫺125 on the bottom strand, and a

D). As a positive control for the ERSR pathway, the glucose deprivation blot was also reprobed to measure endogenous GRP78 mRNA. For panels B and D, the GH reporter gene mRNA expression was quantified, corrected for loading, normalized as percentage of the longest AS fragment for the histidine- or glucose-deprived data (indicated with an asterisk), and plotted in a bar graph relative to the loading control. The schemes in the x axis designate the fragments present in the transfected plasmids for each lane.

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FIG. 2. Deletion analysis of the ⴚ615/ⴙ51 AS promoter fragment. HepG2 hepatoma cells were transfected with a GH reporter construct lacking a promoter (p0GH), containing the constitutive metallothionein promoter (MTT), or containing the indicated sequence of the AS gene. After incubating cells for 12 h in complete MEM (Control), histidine-free MEM (-His), or glucose-free MEM (⫺Glc), expression of the GH reporter mRNA was monitored by Northern analysis (20 ␮g of RNA per lane). As positive controls, the blots were stripped and rehybridized with 32P-radiolabeled cDNA probes to measure the endogenous AS (AAR pathway) and GRP78 (ERSR pathway) mRNA content. Equal loading between the lanes was verified by reprobing the blots with cDNAs for either GDH or L7a. For panels B and D, the GH reporter gene mRNA expression was quantified, corrected for loading, normalized as percentage of the longest AS fragment (indicated with an asterisk), and plotted in a bar graph. The schemes in the x axis designate the fragments present in the transfected plasmids for each lane.

FIG. 3. Determination of the minimal nutrient-responsive AS sequence. Northern analysis was performed using RNA (20 ␮g/lane) extracted from HepG2 human hepatoma cells that were transiently transfected with a human GH reporter construct lacking a promoter (p0GH), containing the constitutive metallothionein promoter (MTT), or containing the AS ⫺173/⫹51 sequence sequentially deleted from either the 5⬘ or 3⬘ direction, as indicated. After employing the batch transfection technique described in the text, cells were incubated for 12 h in MEM (Control) or histidine-free MEM (⫺His). GH reporter mRNA was monitored as a measure of transcriptional activity. Expression of endogenous AS mRNA, a positive control for the AAR pathway, and GDH mRNA, a control for lane loading, was monitored by stripping and rehybridizing the blot with the respective radiolabeled probes. For panels B and D, the GH reporter gene mRNA expression was quantified, corrected for loading, normalized as percentage of the longest AS fragment for the histidine-starved data (indicated with an asterisk), and plotted in a bar graph. The diagrams in the x axis designate the fragments present in the transfected plasmids, for each lane.

protected guanine at position ⫺128 on the top strand. Binding site III was delineated by protected guanines at positions ⫺105, ⫺104, ⫺102, ⫺100, ⫺99, ⫺98, and ⫺97, and an en-

hanced guanine at nt ⫺103 on the bottom strand. In addition, binding site III also had an enhanced guanine at nt ⫺107, and protected guanines at positions ⫺106 and ⫺101 on the top


FIG. 4. The AS genomic fragment ⴚ111/ⴚ34 can confer AAR and ERSR-dependent regulation to the thymidine kinase promoter. The ⫺111/⫺34 sequence from the AS promoter was subcloned either in front of the promoterless GH gene (p0GH plasmid) or in front of a thymidine kinase promoter/GH gene construct (pTKGH), as described in the text. These constructs, with the AS sequence in the forward (F) or the reverse (R) orientation, were transfected into HepG2 human hepatoma cells and tested for the ability to promote GH transcription after incubation for 12 h in complete MEM (Control), histidine-free MEM (⫺His), glucose-free MEM (⫺Glc), or 5 mg/ml tunicamycin-containing MEM (⫹TM). The transcriptional activity was evaluated by Northern analysis (20 ␮g of RNA/lane) of GH mRNA and then the blots were reprobed with AS and GRP78 cDNAs to document activation of AAR and ERSR pathways, respectively. GDH and L7a mRNA content was measured to show equal loading among lanes. The GH reporter gene mRNA expression was quantified, corrected for loading, normalized as percentage of the GH expression under regulation of the ⫺173/⫹51 control AS fragment (indicated with an asterisk), and plotted in a bar graph. The schemes in the x axis designate the fragments present in the transfected plasmids for each lane.

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strand. Binding site IV was not well defined, but did exhibit a protected guanine at nt ⫺79 on the bottom strand and a protected guanine residues at nt ⫺85 on the top strand. Guanines at nt ⫺81 and ⫺82 on the top strand may be slightly protected, but the degree of protection obtained from several gels was never strong. Binding site V had two guanine nucleotides on the top strand, at positions ⫺67 and ⫺64, that were both moderately protected in control cells, but further protected in a nutrient-dependent manner, that is, in response to deprivation for either histidine or glucose (Fig. 5, A and B). Binding site V also had protected guanine nucleotides at ⫺56, ⫺57, and ⫺60 on the bottom strand (Fig. 5, C and D). On the bottom strand, binding site VI exhibited a guanine, nt ⫺44, that was slightly protected in control cells, but to a greater extent following nutrient deprivation. A second bottom strand guanine within site VI, nt ⫺49, was enhanced in its reactivity relative to naked DNA (Fig. 5, C and D). Near several of the guanine contact sites, specific adenine nucleotides consistently showed enhanced dimethyl sulfate reactivity, relative to the in vitro treated DNA. Dimethyl sulfate can methylate adenines at the N-3 position (28), and the enhancement of adenines ⫺61, ⫺62, ⫺63, and ⫺66 on the top strand are particularly obvious examples. Mutagenesis of Specific Nucleotides—Based on the in vivo footprinting analyses, several nucleotides were selected for site-directed mutagenesis (purine for purine, pyrimidine for pyrimidine) to determine if the transcriptional activation by the AAR and the ERSR pathways could be separated by mutation of specific protein contact sites. Each of these mutations was created within the context of the ⫺173/⫹51 AS fragment, which was then cloned in front of the promoterless GH gene and transiently transfected into HepG2 cells. Transfected cells were incubated for 12 h in complete MEM, MEM lacking histidine to activate the AAR pathway, or MEM containing TM or lacking glucose to activate the ERSR pathway (Fig. 6). Interestingly, for each mutation, the nutrient-dependent activation of transcription was the same, regardless of whether the AAR and ERSR pathway was triggered. Mutation of guanine ⫺85 within binding site IV, which was protected in the in vivo footprinting experiments, had no effect on induced transcription rates (Fig. 6A/B). Mutations at nt ⫺67, ⫺64, and ⫺62, all within binding site V, reduced both basal and nutrient-dependent GH transcription (Fig. 6, A, B, and D). In contrast, mutation of the 4 nt from ⫺74 to ⫺71 and guanine ⫺60, nucleotides that appear to flank the two sides of binding site V, had no adverse effect on transcription (Fig. 6, A-C). These results suggest that the core sequence of binding site V is localized from about nt ⫺67 to ⫺62. For binding site VI, mutation of nt ⫺47 and ⫺44 resulted in a loss of nutrient-regulated expression, whereas alteration of nt ⫺40, ⫺42, ⫺48, and ⫺50 had little or no effect (Fig. 6, A, B, and E). These mutations help to further identify binding sites V and VI, and provide additional evidence of their importance for nutrient-dependent regulation of the AS gene. However, none of the mutations resulted in a differential response to activation of the two nutrient-sensing pathways, thus, providing additional support for the proposal that the same set of genomic sequences are required for both nutrient signaling pathways. As shown above, deletion of the AS 5⬘-upstream sequence to nt ⫺90 or less caused a decrease in transcription (Fig. 3, A and B) and there was a strong footprint covering the GC-rich sequence from ⫺107 to ⫺97 (Fig. 5, C and D). To further test the importance of this particular GC box for activation of the AS gene by the AAR and ERSR pathways, block substitution was performed so that the sequence 5⬘-CCCCGCCCC-3⬘ (nt ⫺105/ ⫺97) was mutated to 5⬘-ACTTATTCT-3⬘ in the context of the

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FIG. 5. In vivo dimethyl sulfate footprinting analysis of the AS gene 5ⴕ-flanking sequence. HepG2 cells were incubated for 12 h in complete MEM (Control), histidine-free MEM (⫺His), or glucose-free MEM (⫺Glc). Cells were then treated with dimethyl sulfate and subjected to footprinting analysis. To obtain a complete guanine ladder, dimethyl sulfate treatment was also performed in vitro on isolated DNA (Naked). Primers used for the ligation mediated PCR and other details for the footprinting analysis are described in the text. Panels A (⫺His) and B (⫺Glc) represent analysis of the top strand of DNA, whereas panels C (⫺His) and D (⫺Glc) represent the bottom strand. Each lane represents an individual plate of cells, and the results for each condition were repeated at least twice. Open circles represent protected guanine nucleotides, closed circles represent enhanced guanine nucleotides, open triangles represent guanine nucleotides for which protection was greater in nutrient-deprived cells relative to complete MEM, and the arrowheads represent enhanced adenine nucleotides. A duplicate set of symbols are used, one immediately next to the blot and another next to the corresponding sequence. The roman numerals denote the six regions proposed to contain protein binding sites.


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FIG. 6. Mutagenesis of individual nucleotides within the ⴚ173/ⴙ51 AS sequence. Either wild-type ⫺173/⫹51 AS sequence or the same region containing the indicated mutated nucleotide was subcloned in front of the GH reporter gene. Panels A–E represent independent experiments. The lanes marked Sp1* represent the ⫺173/⫹51 sequence with a block substitution for nt ⫺105/⫺97 (5⬘-CCCCGCCCC-3⬘ to 5⬘-ACTTATTCT-3⬘) within binding site III. The constructs were transfected into HepG2 human hepatoma cells, using as controls the promoterless GH plasmid (p0GH) or the metallothionein promoter/GH construct (MTT). After incubating cells for 12 h in control MEM (Control), histidine-free MEM (⫺His), or glucose-free MEM (⫺Glc), RNA was extracted and GH reporter mRNA was monitored by Northern analysis. Reprobing the blot with cDNAs for AS and GRP78 served as positive controls for AAR and ERSR induction, whereas equal lane loading was evaluated by reprobing the blot with cDNAs for either L7a or GDH.

⫺173/⫹51 construct and then transfected into HepG2 cells (lanes marked “Sp1*” in Fig. 6, A and B). The mutated GC box construct showed decreased basal transcription (compare the lanes marked ⫺173/⫹51 and Sp1* in the Control panel of Fig. 6A), but the AAR or ERSR-induced activity was comparable to the wild-type promoter sequence. Note that this site III construct retains GC-rich sites I and II, defined by in vivo footprinting. Electrophoresis Mobility Shift Analysis—The in vivo footprinting and mutagenesis data suggested that sites V and VI are important for nutrient signaling. To determine if mutagenesis of site V and VI sequences, including some of the same mutations tested for transcriptional activation by transient expression (Fig. 6), resulted in loss of protein-DNA complex formation in vitro, EMSA were performed. A 32P-labeled oligonucleotide containing site V (nt in ⫺79/⫺53) formed three complexes (Fig. 7A). The most abundant complex was increased in amount when a nuclear extract from histidine-deprived cells was tested, but each of the complexes could be blocked by excess competitor DNA (compare WT versus ⫹c lanes, Fig. 7). When individual 32P-labeled oligonucleotides, each containing a single mutation at nt ⫺67, ⫺64, or ⫺62 were used, a significant reduction in complex formation was observed. In contrast, mutation of nt ⫺60 did not prevent starvation-enhanced formation of the most abundant complex (Fig. 7A), nor did it block regulated transcriptional activity (Fig. 6A). An oligonucleotide

containing binding site VI (nt ⫺56/⫺26) formed two specific complexes and the slower migrating complex was increased in amount when nuclear extract from histidine-deprived cells was used (Fig. 7B). Although the absolute amounts of this complex were less than that observed for the wild-type sequence, oligonucleotides containing a mutation at either nt ⫺50 or ⫺48 formed protein-DNA complexes that were increased in amount by amino acid starvation. In contrast, mutation of nt ⫺44 did not result in formation of a detectable amount of protein-DNA complex (Fig. 7B). These results are consistent with the functional expression studies for each of these mutations (Fig. 6). To document that not all protein-DNA complexes are enhanced when extracts from histidine-starved cells are tested, a consensus sequence for the Sp family of transcription factors was used in conjunction with antibody specific for Sp3 (Fig. 7C). Two primary complexes were formed, and the presence of the Sp3 antibody during the incubation resulted in a supershift of one of the complexes. When extracts from histidine-starved cells were tested, there was no increase in formation of either complex, rather a decrease was observed that was also evident for the complex that was shifted by the anti-Sp3 (Fig. 7C). DISCUSSION

The data document the novel observation that both the AAR and unfolded protein response (ERSR) nutrient sensing pathways activate the human AS gene through the same set of

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FIG. 7. EMSA with oligonucleotides containing AS promoter site V or VI detect protein binding in vitro, but mutagenesis of specific nucleotides within these sites does not separate activation by the AAR or ERSR pathway. Nuclear extracts from HepG2 cells maintained for 12 h in MEM (Control) or histidine-free MEM (⫺His) were incubated with 32P-radiolabeled oligonucleotides to monitor formation of DNA-protein complexes by EMSA. A 32P-labeled oligonucleotide containing the wild-type sequence for either binding site V (panel A, nt ⫺79/⫺53) or site VI (panel B, nt ⫺56/⫺26) was incubated with nuclear extracts in the absence (WT) or presence (“⫹c”) of a 100-fold excess of wild-type unlabeled oligonucleotide as competitor. In parallel incubations, 32P-labeled oligonucleotides were tested that contained the indicated specific single nucleotide mutations, which correspond to several of those mutations monitored for transcriptional activity, as described in the legend to Fig. 6. The data for panel C was collected using a consensus oligonucleotide (5⬘-ATTCGATCGGGGCGGGGCGAGC-3⬘) for the Sp1 family of transcription factors and anti-Sp3 antibody (Santa Cruz Biotechnology). Lane 1 represents the consensus oligonucleotide plus a ⫻ 100 excess of an unrelated sequence as a nonspecific competitor, lanes 2 and 4 represent the consensus oligonucleotide alone, lanes 3 and 5 represent incubations containing the consensus oligonucleotide plus anti-Sp3 antibody, per the supplier’s directions.

genomic elements. Deletion analysis of the AS 5⬘-flanking region indicated that nt ⫺111 to ⫺34 represents a “minimal control unit” that yields basal and regulated transcription at levels equal to a genomic fragment containing nt ⫺10,895/ ⫹1,377. The in vivo footprinting analysis, along with the mutagenesis, revealed the surprising result that there are two discrete protein-DNA contact sites (labeled binding sites V and VI) that are absolutely required for regulated transcription and that both are required for activation of the gene by either metabolite sensing pathway. A third Sp1-like sequence that enhances both the basal and the regulated transcriptional response was also observed, consistent with a previous report by Guerrini et al. (8). The footprinting data document that sites V and VI are occupied in the basal state, but binding is greater in response to activation of either the AAR or ERSR pathway. These results suggest that the mechanism by which these signal transduction pathways induce the transcriptional rate of the AS gene may be directly dependent on, or a modulation of, the basal transcription machinery. Sequential deletion of the

FIG. 8. Summary of the identification of the AS promoter-binding sites responsive to the AAR and ERSR pathways. The roman numerals denote the working model for assignment of the six proteinbinding sites identified by the in vivo footprinting and mutagenesis experiments. The bars show the general area of protein binding, but are not intended to identify specific boundaries. The in vivo footprinting studies are summarized as follows: open circles represent protected guanine nucleotides, closed circles are enhanced guanine nucleotides, open triangles designate guanine nucleotides for which protection was greater in nutrient-deprived cells relative to complete MEM, and the arrowheads represent enhanced adenine nucleotides. For mutagenesis of single nucleotides within those binding sites, a purine was replaced by purine and a pyrimidine for a pyrimidine. Substitutions that resulted in a loss of both AAR- and ERSR-induced transcription activation are boxed, whereas those nucleotides marked with an asterisk were mutated with no loss of regulated transcriptional activation. No mutants were shown to block transcriptional activation by only one of the nutrient-sensing pathways.

three GC-rich boxes in the AS promoter, sites I-III, demonstrated that retention of site III only (nt ⫺107/⫺97) was sufficient to maintain basal transcription and the highest level of activated response to either the AAR or the ERSR pathway. On the other hand, mutagenesis of site III in the context of the ⫺173/⫹51 construct, which retains sites I and II, inhibited basal transcription, but did not prevent the starvation-induced transcription by either pathway. Therefore, regulated transcriptional activity can be maintained by one or both of the first two GC boxes. By 8-nucleotide block substitutions, the palindromic AS sequence from ⫺70 to ⫺64 (5⬘-CATGATG-3⬘), which is within binding site V, had been shown by Guerrini et al. (8) to be important for the response to amino acid deprivation and those authors proposed that this sequence functions as an amino acid response element. Noting a partial homology within this sequence (nt ⫺66 to ⫺63, 5⬘-ATGA-3⬘) with the consensus sequence for AP1 (5⬘-ATGA(C/G)TCAT-3⬘), Guerrini et al. (8) performed EMSA with an oligonucleotide covering binding site V and purified Fos-Jun proteins, but concluded from those experiments that AP1 was unlikely to be involved in amino acid-dependent regulation of the AS gene. The present data, both functional assays and EMSA, document that mutations at nt ⫺67, ⫺64, and ⫺62 within binding site V are fundamentally important for basal transcription of the AS gene as well as activation by both nutrient-sensing pathways. Thus, the human AS promoter sequence from nt ⫺70/⫺62 functions more broadly than just as an amino acid response element. Previously published EMSA data (15) has documented the formation of two specific protein-DNA complexes in vitro using oligonucleotide probes covering binding sites V (nt ⫺79/⫺53) and VI (nt ⫺55/⫺26). For both sites, the amount of one of the complexes was enhanced when nuclear extracts from glucosestarved cells were tested, and the present data demonstrate that the same is true for nuclear extracts derived from histidine-starved cells. From the experiments involving transient

Activation of the Asparagine Synthetase Gene transfection of mutated AS promoter sequences, randomly selected mutations were investigated by EMSA. In every instance where the sequence exhibited decreased transcriptional activity, the corresponding oligonucleotide also was less effective in forming protein-DNA complexes. Together these two independent approaches indicate that the responsive elements for the AAR and ERSR pathways are not separable by mutagenesis of selected nucleotides within binding sites V and VI. For the AS sequence from nt ⫺49 to ⫺30, including binding site VI, the top strand contains the sequence 5⬘-CGTTACAGGAGCCAGGTCG-3⬘ which yields 3⬘-GCAAT-N9-CCAGC-5⬘ on the bottom strand (see summary of Fig. 8). This AS region has some similarity to the recently reported mammalian ERSR element (ERSE), for which the consensus sequence is 5⬘CCAAT-N9-CCACG-3⬘ (11, 12). Those previous studies showed that mammalian ERSR target genes, such as the GRP family, are activated by multiple copies of this cis-element. However, the AS sequence is not a perfect match and its 3⬘ to 5⬘ orientation is unusual relative to all previously described occurrences of the ERSE, which can occur on the bottom strand, but always in the 5⬘ to 3⬘ direction (11, 12). Obviously, the 3⬘ to 5⬘ orientation of the AS sequence would present a completely different DNA structure for protein binding than does the ERSE sequence. Furthermore, several additional lines of evidence argue against the functionality of this AS sequence as an ERSE (15). Conversely, computer analysis reveals that the previously recognized ERSR-induced mammalian genes do not contain sequence that corresponds to that within the AS-binding sites V and VI (nt ⫺70 to ⫺34) and, with the exception of calreticulin (35) and CHOP/GADD153 (discussed below), these ERSR responsive genes are not induced by amino acid deprivation (15). Therefore, we conclude that the human AS gene contains unique genomic regulatory sequences, within the region bordered by protein-binding sites V and VI, that can mediate transcriptional activation by the ERSR and AAR pathways. Fafournoux and colleagues have shown that, like AS, transcription of the CHOP/GADD153 gene is induced by both the AAR and the ERSR pathways (38, 39). CHOP/GADD153 is a transcription factor in the C/EBP family that has been implicated to be a component or modulator of the ERSR pathway (19, 36, 37). However, in contrast to the present results, deletion analyses by Jousse et al. (38) suggested that the activation of the CHOP/GADD153 gene by the two pathways may occur through distinct cis-elements. Based on computer analysis, neither site V nor VI AS regulatory sequences are present as completely conserved elements in the CHOP/GADD153 proximal promoters, although it does contain a sequence with some similarity to AS promoter site V, as noted by Bruhat et al. (39). Given what is known about the initial steps in the signal transduction pathways used by yeast and mammalian cells in response to amino acid deprivation (6, 40) or to abnormal protein accumulation (19, 41, 42), the presumption is that the AAR and ERSR pathways are not likely to share common steps early in the process. Furthermore, that the ERSE-containing genes are not activated by amino acid starvation is evidence that the two pathways can function independently. However, the present results provide strong evidence that, at least for the human AS gene, these pathways converge on a common set of genomic elements comprised of sequences that do not have absolute identity to any known transcription factors (43, 44). This common set of AS promoter elements may be bound by the same collection of transcription factors, regardless of which

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pathway is activated, or they may bind two different sets of protein factors, each associated with only one of the pathways. Acknowledgments—We thank Dawn Beachy for secretarial support and other members of our laboratories for technical advice and helpful discussion. We also thank Dr. Pierre Fafournoux for helpful discussion and encouraging us to further investigate the mechansim of glucose deprivation on asparagine synthetase gene expression. REFERENCES 1. Lee, A. S. (1992) Curr. Opin. Cell Biol. 4, 267–273 2. Carlson, S. G., Fawcett, T. W., Bartlett, J. D., Bernier, M., and Holbrook, N. L. (1993) Mol. Cell. Biol. 13, 4736 – 4744 3. Tirasophon, W., Welihinda, A. A., and Kaufman, R. J. (1998) Gene Dev. 12, 1812–1824 4. Pahl, H. L. (1999) Physiol. Rev. 79, 683–701 5. Barbosa-Tessmann, I. P., Pineda, V. L., Nick, H. S., Schuster, S. M., and Kilberg, M. S. (1999) Biochem. J. 339, 151–158 6. Laine, R. O., Hutson, R. G., and Kilberg, M. S. (1996) Prog. Nucleic Acid Res. Mol. Biol. 53, 219 –248 7. Gong, S. S., Guerrini, L., and Basilico, C. (1991) Mol. Cell. Biol. 11, 6059 – 6066 8. Guerrini, L., Gong, S. S., Mangasarian, K., and Basilico, C. (1993) Mol. Cell. Biol. 13, 3202–3212 9. Hutson, R. G., and Kilberg, M. S. (1994) Biochem. J. 303, 745–750 10. Hutson, R. G., Kitoh, T., Amador, D. A. M., Cosic, S., Schuster, S. M., and Kilberg, M. S. (1997) Am. J. Physiol. 272, C1691–C1699 11. Yoshida, H., Haze, K., Yanagi, H., Yura, T., and Mori, K. (1998) J. Biol. Chem. 273, 33741–33749 12. Roy, B., and Lee, A. S. (1999) Nucleic Acids Res. 27, 1437–1443 13. Roy, B., Li, W. W., and Lee, A. S. (1996) J. Biol. Chem. 271, 28995–29002 14. Haze, K., Yoshida, H., Yanagi, H., Yura, T., and Mori, K. (1999) Mol. Biol. Cell 10, 3787–3799 15. Barbosa-Tessmann, I. P., Chen, C., Zhong, C., Schuster, S. M., Nick, H. S., and Kilberg, M. S. (1999) J. Biol. Chem. 274, 31139 –31144 16. Story, M. D., Voehringer, D. W., Stephens, L. C., and Meyn, R. E. (1993) Cancer Chem. Pharm. 32, 129 –133 17. Bussolati, O., Belletti, S., Uggeri, J., Gatti, R., Orlandini, G., Dall’Asta, V., and Gazzola, G. C. (1995) Exp. Cell Res. 220, 283–291 18. Zinszner, H., Kuroda, M., Wang, X. Z., Batchvarova, N., Lightfoot, R. T., Remotti, H., Stevens, J. L., and Ron, D. (1998) Gene Dev. 12, 982–995 19. Kaufman, R. J. (1999) Gene Dev. 13, 1211–1233 20. Greco, A., Gong, S. S., Ittmann, M., and Basilico, C. (1989) Mol. Cell. Biol. 9, 2350 –2359 21. Zhang, Y. P., Lambert, M. A., Cairney, A. E. L., Wills, D., Ray, P. N., and Andrulis, I. L. (1989) Genomics 4, 259 –265 22. Selden, R. F., Howie, K. B., Rowe, M. E., Goodman, H. M., and Moore, D. D. (1986) Mol. Cell. Biol. 6, 3173–3179 23. Ho, S. N., Hunt, H. D., Horton, R. M., Pullen, J. K., and Pease, L. R. (1989) Gene (Amst.) 77, 51–59 24. Hamer, D. H., and Walling, M. (1982) J. Mol. Appl. Genet. 1, 273–288 25. Chomczynski, P., and Sacchi, N. (1987) Anal. Biochem. 162, 156 –159 26. Lee, A. S., Delegeane, A., and Scharff, D. (1981) Proc. Natl. Acad. Sci. U. S. A. 78, 4922– 4925 27. Hornstra, I. K., and Yang, T. P. (1993) Anal. Biochem. 213, 179 –193 28. Maxam, A. M., and Gilbert, W. (1980) Methods Enzymol. 65, 499 –560 29. Mueller, P. R., and Wold, B. (1989) Science 246, 780 –786 30. Church, G. M., and Gilbert, W. (1984) Proc. Natl. Acad. Sci. U. S. A. 81, 1991–1995 31. Kerrigan, L. A., and Kadonaga, J. T. (1994) in Current Protocols in Molecular Biology (Ausubel, F. M., eds) John Wiley & Sons, Inc., New York 32. Ioannou, P. A., Amemiya, C. T., Garnes, J., Kroisel, P. M., Shizuya, H., Chen, C., Batzer, M. A., and De Jong, P. J. (1994) Nature Genet. 6, 84 – 89 33. Courey, A. J., and Tjian, R. (1992) in Transcriptional Regulation (McKnight, S. L., and Yamamoto, K. R., eds) pp. 743–769, Cold Spring Harbor Laboratory, Cold Spring Harbor, NY 34. Smale, S. T. (1997) Biochim. Biophys. Acta 1351, 73– 88 35. Heal, R., and McGivan, J. D. (1998) Biochem. J. 329, 389 –394 36. Wang, X.-Z., Harding, H. P., Zhang, Y., Jolicoeur, E. M., Kuroda, M., and Ron, D. (1998) EMBO J. 17, 5708 –5717 37. Wang, X.-Z., Kuroda, M., Sok, J., Batchvarova, N., Kimmel, R., Chung, P., Zinszner, H., and Ron, D. (1998) EMBO J. 17, 3619 –3630 38. Jousse, C., Bruhat, A., Harding, H. P., Ferrara, M., Ron, D., and Fafournoux, P. (1999) FEBS Lett. 448, 211–216 39. Bruhat, A., Jousse, C., Wang, X.-Z., Ron, D., Ferrara, M., and Fafournoux, P. (1997) J. Biol. Chem. 272, 17588 –17593 40. Hinnebusch, A. G. (1997) J. Biol. Chem. 272, 21661–21664 41. Shamu, C. E. (1998) Curr. Biol. 8, R121-R123 42. Sidrauski, C., Chapman, R., and Walter, P. (1998) Cell Biol. 8, 245–249 43. Heinemeyer, T., Wingender, E., Reuter, I., Hermjakob, H., Kel, A. E., Kel, O. V., Ignatieva, E. V., Ananko, E. A., Podkolodnaya, O. A., Kolpakov, F. A., Podkolodny, N. L., and Kolchanov, N. A. (1998) Nucleic Acids Res. 26, 362–367 44. Quandt, K., Frech, K., Karas, H., Wingender, E., and Werner, T. (1995) Nucleic Acids Res. 23, 4878 – 4884