Advertisement
Research Article

Transcriptional Regulation of N-Acetylglutamate Synthase

  • Sandra Kirsch Heibel,

    Affiliations: Center for Genetic Medicine Research, Children's National Medical Center, Washington, D. C., United States of America, Molecular and Cellular Biology Program, University of Maryland, College Park, Maryland, United States of America

    X
  • Giselle Yvette Lopez,

    Affiliation: Department of Pathology, Duke University Medical Center, Durham, North Carolina, United States of America

    X
  • Maria Panglao,

    Affiliation: The George Washington University School of Medicine and Health Sciences, Washington, D. C., United States of America

    X
  • Sonal Sodha,

    Affiliation: Johns Hopkins School of Medicine in Baltimore, Maryland, United States of America

    X
  • Leonardo Mariño-Ramírez,

    Affiliation: Computational Biology Branch, National Center for Biotechnology Information, National Library of Medicine, National Institutes of Health, Bethesda, Maryland, United States of America

    X
  • Mendel Tuchman,

    Affiliation: Center for Genetic Medicine Research, Children's National Medical Center, Washington, D. C., United States of America

    X
  • Ljubica Caldovic mail

    ljubica@cnmcresearch.org

    Affiliation: Center for Genetic Medicine Research, Children's National Medical Center, Washington, D. C., United States of America

    X
  • Published: February 27, 2012
  • DOI: 10.1371/journal.pone.0029527

Abstract

The urea cycle converts toxic ammonia to urea within the liver of mammals. At least 6 enzymes are required for ureagenesis, which correlates with dietary protein intake. The transcription of urea cycle genes is, at least in part, regulated by glucocorticoid and glucagon hormone signaling pathways. N-acetylglutamate synthase (NAGS) produces a unique cofactor, N-acetylglutamate (NAG), that is essential for the catalytic function of the first and rate-limiting enzyme of ureagenesis, carbamyl phosphate synthetase 1 (CPS1). However, despite the important role of NAGS in ammonia removal, little is known about the mechanisms of its regulation. We identified two regions of high conservation upstream of the translation start of the NAGS gene. Reporter assays confirmed that these regions represent promoter and enhancer and that the enhancer is tissue specific. Within the promoter, we identified multiple transcription start sites that differed between liver and small intestine. Several transcription factor binding motifs were conserved within the promoter and enhancer regions while a TATA-box motif was absent. DNA-protein pull-down assays and chromatin immunoprecipitation confirmed binding of Sp1 and CREB, but not C/EBP in the promoter and HNF-1 and NF-Y, but not SMAD3 or AP-2 in the enhancer. The functional importance of these motifs was demonstrated by decreased transcription of reporter constructs following mutagenesis of each motif. The presented data strongly suggest that Sp1, CREB, HNF-1, and NF-Y, that are known to be responsive to hormones and diet, regulate NAGS transcription. This provides molecular mechanism of regulation of ureagenesis in response to hormonal and dietary changes.

Introduction

Ammonia, the toxic product of protein catabolism, is converted to urea by the urea cycle in the liver of mammals. Incorporation of two nitrogen atoms into urea is catalyzed by six enzymes: three of them mitochondrial, N-acetylglutamate synthase (NAGS; EC 2.3.1.1), carbamylphosphate synthetase 1 (CPS1; EC 6.4.3.16) and ornithine transcarbamylase (OTC; EC 2.1.3.3), and the other three cytosolic, argininosuccinate synthetase (ASS; EC 6.3.4.5), argininosuccinate lyase (ASL; EC 4.3.2.1) and arginase 1 (Arg1; EC 3.5.3.1).

NAGS catalyzes the formation of N-acetylglutamate (NAG), an essential allosteric activator of CPS1, in the mitochondrial matrix of hepatocytes and small intestine epithelial cells [1], [2]. Within hepatocytes, NAGS activity and NAG abundance are regulated by L-arginine, ammonia, and dietary protein intake [3], [4], [5] and therefore, the NAGS/NAG system may play a critical role in the regulation of ureagenesis in response to these factors [6]. While studies in the 1980s and 1990s identified the cis-acting motifs regulating transcription of the urea cycle enzymes CPS1 [7], [8], [9], [10], OTC [11], [12], [13], [14], ASS [15], [16], [17], ASL [18], [19], [20], and Arg1 [21], [22], the mammalian NAGS gene was not identified until 2002 [2] and we can now report for the first time on its transcriptional regulation.

Many studies have identified regulatory links between the urea cycle genes and glucocorticoids and glucagon [23], [24], [25], however the mechanism of regulation differs for each gene [24], [26], [27], [28], [29]. Transcription of CPS1 is activated by TATA-binding protein (TBP) while its proximal and distal enhancers contain binding sites for glucocorticoids and cAMP responsive factors including CCAAT-enhancer bind protein (C/EBP), activator protein-1 (AP-1), glucocorticoid receptor (GR) and cAMP response element binding (CREB). Sites for binding tissue specific factors including hepatic nuclear factor 3 (HNF-3) are also present [25], [30], [31]. Tissue specific expression of the OTC gene is induced in the intestine and liver by HNF-4, which binds in the promoter [13], [14], [32] while binding of both HNF-4 and C/EBP to the enhancer, induces high expression levels in the liver [12], [13], [14], [25], [33]. ASS transcription is regulated by cooperative binding of multiple specificity protein 1 (Sp1) 16,34,35,36. ASL is regulated through Sp1 and the positive regulator, nuclear factor Y (NF-Y), which binds within the promoter of ASL to activate its transcription [18], [19], [20], [37]. Sp1 and nuclear factor 1 (NF-1)/CCAAT-binding transcription factor (CTF) activate ARG1 transcription while two C/EBP factors and two unidentified proteins bind within an enhancer in intron 7 to confer glucocorticoid responsiveness [22].

Abundance of urea cycle enzymes correlates with dietary protein intake [3], [28]. Transcription of urea cycle genes is in part regulated by the glucocorticoid and glucagon signaling pathways [29], [38]. Therefore, we postulate that there exists a nitrogen sensing mechanism that is both responsive to amino acid(s) and hormone stimulation and that an understanding of the transcriptional regulation of NAGS could contribute to the understanding of such mechanism.

In this study, we identified two regulatory regions upstream of the NAGS translation start site that contain highly conserved protein-binding DNA motifs. We subsequently confirmed that these regions function as promoter and enhancer and that the enhancer is most effective in liver cells. Avidin-agarose protein-DNA pull-down assays have been used to confirm binding of Sp1 and CREB within the NAGS promoter and Hepatic Nuclear Factor 1 (HNF-1) and NF-Y within the enhancer regions. Chromatin immunoprecipitation (ChIP) and quantitative real-time PCR have been used to independently verify that Sp1 and CREB bind to the promoter region, and HNF-1 and NF-Y bind to the enhancer region. We also used 5′RACE analysis to identify multiple transcription start sites for NAGS that may be species and tissue specific. These findings provide new information on the regulation of the NAGS gene, and suggest possible mechanisms for coordinated regulation of the genes involved in ureagenesis.

Materials and Methods

Bioinformatic Analysis of the Upstream Regulatory Regions

Pair-wise Alignment Analysis.

Identification of highly conserved regions was conducted by gathering 15 kilobases of genomic sequence 5′ of the NAGS translational start site and sequence of intron one in 7 mammalian species including: human (NM_153006.2), chimpanzee (XM_001152480.1), dog (XM_548066.2), cow (XM_618194.4), horse (XM_001917302.1), mouse (NM_145829.1) and rat (NM_001107053.1). The highly conserved regulatory regions of CPS1 were identified by gathering 15 kilobases of genomic sequences 5′ of the translational start site from human (NM_001875), chimpanzee (XM_001146604), dog (XM_856862), mouse (NM_001080809), and rat (NM_017072). Genomic sequences were subject to pair-wise comparison using BLAST bl2seq tool [39]. Parameters included expect threshold of 10, match and mismatch scores of 1 and −2, respectively, gap existence and extension scores of 5 and 2 respectively, and maximum expected value E = 0.001. Regions of high conservation were identified as sequences with more than 80% identity that were at least 100 bp long and present in four or more species.

Cis-eLement OVERrepresentation (CLOVER) Analysis.

The Cis-element OVERrepresentation (CLOVER) [40] program was used to predict the over-represented motifs within the highly conserved regulatory regions of NAGS and CPS1. CLOVER analysis of these conserved regions identified known protein binding DNA motifs in the TRANSFAC Pro database by calculating over-representation of these sequences compared to a background of ppr_build_33.fa generated from NCBI build 33 [41]. Matrices recognized by multiple transcription factors in the same family are represented by one family member unless otherwise noted. Genomic sequences of the highly conserved regions were aligned using CLUSTALW version 2.0.10 [42].

Plasmid Constructs

The promoter and enhancer of NAGS, were amplified from human genomic DNA with primer pairs hPromXH and hEnhXH or hPromHXrev and hEnhHXrev (Table S1), respectively, to introduce XhoI and HindIII restriction enzyme sites and allow subcloning in forward and reverse orientation. Platinum Taq PCRx DNA Polymerase (Invitrogen) was used for amplification with the following conditions: initial denaturation at 95°C for 2 min., followed by 35 cycles of denaturation at 95°C for 30 sec., annealing at 57°C for 30 sec. and extension at 68°C for 1 min., and final extension at 68°C for 6 min. Promoter and enhancer PCR products were ligated with TOPO-TA sequencing vector (Invitrogen) according to manufacturer's instructions and referred to as TOPOProm, TOPOEnh, TOPOPromRev, and TOPOEnhRev, respectively. Mouse Nags (mNags) promoter and enhancer were inserted into TOPO-TA vector following the same methods. Correct DNA sequences were confirmed using sequencing primers specified by Invitrogen.

TOPOProm, TOPOEnh, TOPOPromRev, TOPOEnhRev, pGL4.10 (Promega) basic vector containing firefly (Photinus pyralis) luciferase luc2, and pGL4.23 (Promega) vector containing a minimal TATA promoter with luc2 were cut with XhoI (New England Biolabs) and HindIII (New England Biolabs). The vectors were treated with Antarctic Alkaline Phosphatase (AAP) (New England Biolabs) according to manufacturer's instructions, and the NAGS regions were ligated with the vectors to form the plasmids in Table 1. TOPOEnh was also amplified with primer pair hEnhBS (Table S1), to introduce BamHI and SalI restriction enzyme sites at the 5′ and 3′ ends of the enhancer, respectively. The amplified enhancer product and 4.10Prom were cut with BamHI (New England Biolabs) and SalI (New England Biolabs), the vector was treated with AAP, and the enhancer was ligated with the vector (Table 1). Plasmids containing mouse NAGS promoter and enhancer were generated using the same methods with the primer pairs listed in Table S1 and plasmids in Table 1. Correct sequences were confirmed using primers specified by Promega.

thumbnail

Table 1. Plasmids generated for luciferase reporter assays.

doi:10.1371/journal.pone.0029527.t001

Point mutations in the binding sites for transcription factors Sp1, HNF-1 and NF-Y were selected based on functional analysis of Sp1 [43], [44], [45], HNF-1 [46], [47], and NF-Y [48], [49] binding in other genes. Mutations were engineered by Integrated DNA Technologies and provided in pIDTSMART-KAN vectors (IDT) (Table 2). Plasmids with mutant Sp1, HNF-1, and NFY were cut with XhoI and HindIII. Reporter plasmids pGL4.10, and pGL4.23 were cut with XhoI and HindIII and treated with AAP. Mutated inserts were ligated with vectors to form the plasmids 4.10Sp1m, 4.23HNF-1m, and 4.23NFYm (Table 1). Correct sequences were confirmed using primers specified by Promega.

thumbnail

Table 2. Mutations in Sp1 and CREB binding sites in the promoter, and HNF-1 and NF-Y in the enhancer of human NAGS.

doi:10.1371/journal.pone.0029527.t002

Point mutations in the CREB binding site, c.-7T>C and c.-5T>A (Table 2), were selected based on functional analysis of CREB binding [50], [51] in other genes and were engineered into the NAGS gene using QuickChange Lightening Site-Directed Mutagenesis Kit (Agilent) according to manufacturer's instructions. Primers hCREBm Fw and Rv (Table S1) amplified 50 ng of template plasmid 4.10Prom to create 4.10CREBm. The correct sequence was confirmed using primers specified by Promega.

The expression vectors encoding Sp1 or HNF-1 cDNA were under control of the cytomegalovirus promoter (Origene).

Tissue culture

Cell culture and transfection.

Human hepatoma cells (HepG2) (donated by Dr. Marshall Summar, Children's National Medical Center, Washington, DC) were cultured in complete media containing RPMI 1640 medium (Invitrogen) supplemented with 10% fetal bovine serum (FBS) (ATCC) and 5% Penicillin/Streptomycin (Invitrogen) under 5% CO2 at 37°C. Human alveolar basal epithelial cells (A549) (donated by Dr. Mary Rose, Children's National Medical Center, Washington, DC) were cultured in complete media containing Ham's F-12 medium (Invitrogen) supplemented with 10% FBS and 5% Penicillin/Streptomycin. Human colorectal adenocarcinoma cells (Caco-2) (ATCC) were cultured in Eagle's Minimum Essential Medium (Invitrogen) supplemented with 20% FBS. Cells were plated at a density of 5×105 cells/well on 24-well culture plates 24 hours prior to transfection. The cells (90–95% confluent for HepG2 and A549, 80–85% confluent for Caco-2) were then transfected using Lipofectamine 2000 reagent (Invitrogen) and cultured in transfection media containing medium and serum only. A total of 0.25 ug of DNA was transfected with 0.225 ug of vector expressing luc2 and 0.025 ug of pGL4.74 vector containing Renilla reniformis luciferase (hRluc) as an internal control (Promega). For co-transfections 0.225 ug of luc2 vector was combined with either 0.25 ug of expression vector or empty vector pUC19 (Invitrogen), and 0.025 ug of hRluc control vector.

Reporter assays

24 hours following transfection, cells were assayed for both firefly and Renilla luciferase activity using Dual-Luciferase Reporter Assay System (Promega) and Berthold Centro 960 luminometer (Berthold) according to the manufacturer's protocol. All reporter assay measurements were corrected for transfection efficiency by normalizing the firefly luciferase signal to the Renilla luciferase values. Expression level of each construct was determined relative to luciferase expression under control of the NAGS promoter in each cell line. All results are an average of three independent experiments that were each carried out in triplicate. Values were expressed as mean ± SEM and analyzed using Student's t-test.

5′ Rapid Amplification of cDNA Ends (RACE)

5′ RACE (Version 2.0; Invitrogen) was performed using RNA isolated from donated mouse livers by Trizol reagent (Invitrogen). RNA from mouse small intestine (Origene), human duodenum (Ambion), or human liver (Ambion) was commercially available. Products were synthesized with human or mouse NAGS specific primers complementary to sequence within Exon 1 (Table S2). All reactions began with 5 ug of total RNA and the RACE procedure was conducted according to manufacturer's instructions. Second strand synthesis was conducted using Ex Taq Polymerase (TaKaRa Bio Inc.) PCR products were subcloned into pCR 2.1-TOPO vector (Invitrogen) and RACE products were sequenced with primers specified by the manufacturer.

Avidin-Agarose DNA-Protein Pull-Down Assay

Biotinylated DNA probes.

Probes for Avidin-Agarose DNA-Protein Pull-Down Assays were generated by PCR amplification of genomic DNA isolated from donated mouse tails using Pure Gene DNA Purification Kit (Gentra). Probes were generated using biotinylated or non-biotinylated forward primer and non-biotinylated reverse primers with Platinum Taq PCRx DNA Polymerase (Invitrogen) and amplification conditions: initial denaturation at 95°C for 2 min., followed by 35 cycles of denaturation at 95°C for 30 sec., annealing at 60°C for 30 sec. and extension at 68°C for 1 min., and final extension at 68°C for 6 min. The mouse Nags (mNags) promoter regions A and B (Figure 1) were amplified with primer pair mNAGS-Prom Region A, from +97 to −259, relative to the translation initiation codon and with mNAGS-Prom Region B, from −302 to −776, respectively (Table S3). A region of mNags, that is not highly conserved in mammals, −1056 to −1320, was amplified using primer pair mNAGS-Prom-NC to serve as a negative control for the promoter regulatory region. The enhancer region of mNAGS, spanning from −2834 to −3167, was amplified using forward primer pair mNAGS-enh. The negative control for the enhancer region, a non-conserved region located close to enhancer, was the amplification product of primer pair mNAGS-Enh-NC spanning −5569 to −5997 upstream of mNags. Additional negative controls, non-biotinylated probes, were generated using each primer pair.

thumbnail

Figure 1. Regions upstream of the mammalian NAGS genes that are highly conserved.

Conservation of mammalian NAGS DNA by phastCons (green) and phyloP (blue) algorithms is shown with the highly-conserved regions indicated in red boxes (A). Pair-wise blast analysis of mammalian non-coding regions of NAGS identified highly conserved sequences upstream of the translational start site termed the promoter (purple) and enhancer (cyan) (B).

doi:10.1371/journal.pone.0029527.g001
Preparation of nuclear extracts.

Nuclear extract was isolated from donated adult mouse livers of C57BL/6 mice using Nuclear Extraction Kit (Origene) according to manufacturer's instructions. The protein concentration of the nuclear extract was determined using bovine serum albumin as the protein standard with Bradford Assay dye concentrate reagents (Bio-Rad). On average, 10 mg of nuclear protein was obtained from mouse liver.

Binding Protocol and Western Blot.

For the avidin-agarose protein-DNA pull-down assay [52], 1 mg of nuclear extract in PBS buffer containing inhibitors (PBSI; 1× PBS with 0.5 mM PMSF, 25 mM β-glycerophosphate, mM NaF), 15 ug of DNA probe, and avidin-agarose beads (Sigma) were combined and incubated for 16 hrs on a rotating shaker at 4°. The probe and bead concentrations were in excess to ensure complete pull-down of DNA–protein complexes. Following incubation, the supernatant was reserved while the beads were washed 3 times with cold PBSI and then resuspended and boiled in Laemmli protein denaturing buffer (Bio-Rad) with 0.2 M DTT. The supernatant was also combined with denaturing buffer with DTT and boiled; all samples were loaded onto 10% SDS–polyacrylamide gel. The proteins were separated by electrophoresis, transferred to a nitrocellulose membrane, and then identified by immunoblotting using primary antibodies at 1:2000 dilution of antibody to Sp1 (Santa Cruz Biotech; Millipore), 1:1000 dilution of CREB-1α/β (Santa Cruz Biotech), and 1:3000 dilution of C/EBPα/β (Santa Cruz Biotech) for the promoter region and 1:500 dilution of HNF-1α/ß (Santa Cruz Biotech), 1:1000 dilution of NF-Yα (Santa Cruz Biotech) and 1:2000 dilution of SMAD2/3 (Santa Cruz Biotech) for the −3 kb conserved region. The membrane was than incubated with 1:20,000 dilution of donkey anti-rabbit secondary antibody conjugated to horseradish peroxidase (Pierce) and bands were visualized using SuperSignal West Pico Kit (Pierce) according to manufacturer's instructions.

Chromatin Immunoprecipitation

Tissue preparation and DNA immunoprecipitation.

Donated livers from adult C57BL/6 mice were minced and chromatin was precipitated using SimpleChIP Enzyme Chromatin Kit (Origene) with the variation for whole tissue. Briefly, fresh tissue was minced and washed with PBS including Protease Inhibitor Complete tablets (Roche). Proteins and DNA were cross-linked with 1.5% formaldehyde, and tissue was disaggregated with dounce homogenizer. Chromatin was sheared to an approximate size of 100–1000 bp by micrococcal nuclease digestion followed by sonication. Immunoprecipitation was conducted using antibodies to transcription factors Sp1 (Millipore), CREB (Santa Cruz Biotech), C/EBP (Santa Cruz Biotech), HNF-1 (Santa Cruz Biotech), NF-Y (Santa Cruz Biotech), SMAD2 (Santa Cruz Biotech) and AP-2 (Santa Cruz Biotech) and control antibodies to histone H3 and non-specific rabbit IgG (Cell Signaling Technologies). Chromatin was eluted from protein G agarose beads, cross-linking was reversed, and DNA was purified according to manufacturer's instructions.

Real-time PCR quantification.

ChIP enriched DNA samples included 2% input control and dilutions for a standard curve, positive control immunoprecipitate from anti-histone H3 antibody sample, negative control immunoprecipitation from anti-rabbit IgG antibody, no antibody control, water control, and test antibodies. Enriched DNA was subject to quantitative real-time PCR using iTaq SYBR Green Supermix with ROX (Bio-Rad) and gene specific primers (Table S4) including negative locus primers to Chemokine ligand 2 (MIP-2) on a 7900HT Fast Real-Time PCR System (Applied Biosystems). Amplification conditions included initial denaturation at 95°C for 2 min., followed by 50 cycles of denaturation at 95°C for 30 sec., annealing at 60°C for 30 sec. and extension at 72°C for 30 sec., with dissociation steps of 95° for 15 sec. followed by 50° for 15 sec. and finally 95° for 15 sec. Samples were amplified and analyzed using 7900HT Sequence Detection System Software (Applied Biosystems). Values were expressed as mean ± SEM and analyzed using Student's t-test.

Results

Selected regions of non-coding DNA upstream of NAGS are highly conserved

15 kilobase of genomic DNA sequence 5′ of the translational start site of NAGS and sequence of the first intron from human, chimpanzee, dog, horse, cow, mouse and rat were aligned and compared using pair-wise BLAST. Comparisons showed three highly conserved regions upstream of human NAGS at −57 to −284, −498 to −576, and −2978 to −3344 relative to the start ATG, and no significant conservation within the intron or between −5 and −15 kb upstream (Figure 1). The region within −1 kb of the translational start site was designated as the putative promoter while the region 3 kb upstream was designated a putative regulatory element. Figure 1 also shows an alignment of mammalian NAGS genes using phastCons (green) and phyloP (blue), which identified three non-coding regions of conservation located 3 kb upstream, immediately upstream, and within the first intron of NAGS, respectively (Figure 1). The phastCons, phyloP and our analyses of conservation within the NAGS gene differed due to different algorithms that were used to identify regions of conservation [39], [53], [54].

To validate our strategy for identification of conserved regions, the same analyses were conducted for CPS1, a gene in which a proximal promoter and an enhancer element located 6.3 kb upstream of rat Cps1, have been characterized [55], [56], [57]. 15 kb of CPS1 genomic DNA sequence 5′ of the translational start site was collected from human, chimpanzee, dog, mouse and rat and compared using pair-wise BLAST. Five regions of high conservation were identified including the previously reported proximal promoter located immediately upstream of the translation initiation codon and the enhancer at −7392 to −7966 relative to ATG of the human CPS1 gene (Figure S1). In addition, three previously unknown regions, termed A, B and C, were also identified at −5, −10.5 and −12 kb relative to CPS1 translation initiation codon (Figure S1). PhastCons and phyloP alignment of mammalian genomic DNA identified the same 5 conserved regions (Figure S1).

Highly conserved, non-coding regions of NAGS function as promoter and enhancer elements for gene transcription

Reporter assays were used to examine the functionality of each of the following: wild type NAGS promoter (4.10Prom), control reversed promoter (4.10PromRev), enhancer alone (4.10Enh), promoter and enhancer (4.10PromEnh), and enhancer in both orientations with the heterologous TATA-box promoter (4.23Enh and 4.23EnhRev) by measuring the expression of a luciferase reporter gene in cultured HepG2 cells (Figure 2A). Vectors pGL4.13, pGL4.23, and pGL4.10 containing firefly luciferase luc2, with an SV40 promoter, a minimal TATA-promoter, or without a promoter respectively, were used as positive, baseline reference, and negative assay controls. Vector pGL4.74, containing Renilla luciferase hRluc, was co-transfected with each plasmid to control for transfection efficiency.

thumbnail

Figure 2. Highly conserved regulatory regions, upstream of the NAGS gene, function as promoter and enhancer elements.

In liver derived cells the NAGS promoter (4.10Prom), promoter+enhancer (4.10PromEnh), enhancer with TATA promoter (4.23Enh), and positive control promoter vector (pGL4.13) significantly simulate transcription while the enhancer (4.10Enh), basic vector (pGL4.10) does not stimulate transcription above baseline (A). Reverse insertion of the promoter (4.10PromRev) did not stimulate transcription compared to 4.10Prom and pGL4.10 vector (B), but reverse enhancer (4.23EnhRev) significantly stimulated transcription compared to 4.23Enh and pGL4.23 vector (C). Calculated results are an average of three independent experiments that were each carried out in triplicate, normalized to Rluc expression, and expressed relative to the promoter for each experiment with error reported as ±SEM. Lowercase letters indicate statistically significant differences.

doi:10.1371/journal.pone.0029527.g002

The human NAGS promoter alone (plasmid 4.10Prom), stimulated transcription of the luciferase gene while the upstream regulatory region (plasmid 4.10Enh) alone, did not (Figure 2A). When the NAGS promoter and upstream regulatory region were both present (4.10PromEnh plasmid), transcription increased by 50% compared to the promoter alone confirming that the upstream conserved region can function as an enhancer of transcription. When the NAGS enhancer was paired with a heterologous promoter containing a TATA-box, in the 4.23Enh construct, the transcription of luciferase about three times higher compared to construct with minimal TATA-box. The backbone vector 4.10 did not stimulate expression of the luciferase gene. As expected, positive control vector 4.13, containing a strong promoter, activated transcription in this cell culture system (Figure 2A). The promoter in the reverse orientation (4.10PromRev) did not activate luciferase expression indicating that the NAGS promoter acts in a direction dependent manner (Figure 2B). The ability of the NAGS enhancer (4.23EnhRev) to stimulate transcription with the heterologous promoter was orientation independent (Figure 2C). Similar results were obtained for reporter assays using mouse promoter and enhancer (Figure S2).

Transcription of NAGS initiates at multiple sites

Following discovery of the NAGS promoter, the transcriptional start sites (TSS) in human and mouse liver and small intestine were identified using 5′ RACE (Figure 3A and B). Cloned and sequenced amplification products from 5′RACE were aligned along the 5′ non-coding region of NAGS along with TSS identified in the Database of Transcriptional Start Sites (DBTSS) and expressed sequence tags (ESTs) from Genbank. Results suggest that NAGS has multiple TSS and that some may be species and tissue-specific. Combined 5′RACE, DBTSS, and Genbank results indicate that within human liver, the most frequently occurring TSS was at −42 bp upstream of the ATG codon, while in human small intestine it was at −146 bp (Figure 3A). Within mouse tissues, no dominant TSS was evident, but transcription of the NAGS gene initiated most often from −20 bp and −108 bp in liver and −20 bp and −95 bp in small intestine (Figure 3B). Figure 3 also shows several other rare TSS that were identified.

thumbnail

Figure 3. Transcription start sites (TSS) are species and tissue specific.

TSS identified in the promoter of NAGS by 5′RACE analysis (blue circles), the Database of Transcriptional Start Sites (DBTSS) (green circles) and 5′ termination sites of Expressed Sequence Tags (ESTs) from Genbank (orange circles) were aligned on the DNA sequence 5′ of the human (A) and mouse (B) NAGS coding sequence. The arrow indicates the translation start site.

doi:10.1371/journal.pone.0029527.g003

Transcription factors bind highly conserved motifs within the promoter and enhancer of NAGS

When promoters and enhancers from six mammalian NAGS genes were aligned, there were multiple regions of base pair conservation (Figure 4). Cis-eLement OVER-representation (CLOVER) software analysis was employed to identify transcription factor binding motifs in regulatory regions of human, chimpanzee, horse, cow, dog, mouse, and rat NAGS. Analyses of the region +9 to −996 bp (relative to the translational start codon, promoter, Table S6) and −2866 to −3620 bp (enhancer, Table S7) predicted several transcription factor binding motifs that are expressed in the liver, but no TATA-box for transcription initiation. Sp1 binding sites, within the promoter, and the HNF-1 binding motif, within the enhancer, received the highest over-representation scores, but additional motifs with lower scores were also over-represented.

thumbnail

Figure 4. Sequence alignment of NAGS promoters and enhancers from seven mammalian species indicate conserved motifs.

DNA sequence of the promoter (A) and enhancer (B) regions were aligned using CLUSTALW alignment software. CLOVER analysis was used to identify transcription factor binding motifs. Binding sites for C/EBP (green), Sp1 (red), CREB/ATF (pink), AP-2 (purple), HNF-1 (blue), NF-Y (olive), and SMAD 3 (cyan) were highly conserved.

doi:10.1371/journal.pone.0029527.g004

Next, over-represented motifs were mapped on the CLUSTALW alignments (Figure 4A and 4B) and motifs with high conservation, having been identified in at least four out of the seven mammalian species, were examined further. Throughout the promoter, five binding sites for Sp1 were highly conserved, two of which were conserved in all examined species. A binding site recognized by CREB and Activating Transcription Factor-1 (ATF-1) was conserved in four species and overlapped with the translation start codon; a C/EBP binding site was identified farther upstream in region B of the promoter (Figures 4A & 5A). Within the enhancer, a binding site for HNF-1 was conserved in all species. Overlapping binding sites for NF-Y, AP-2 and Mothers Against Decapentaplegic Homolog 3 (SMAD3) were also conserved in all species, while an additional AP-2 binding site, located 5′ of the HNF-1 site, was conserved in four out of seven species (Figure 4B & 5B).

thumbnail

Figure 5. DNA-protein avidin-agarose pull-down assay results confirm transcription factor binding.

Two probes for the promoter (A) and one probe for the enhancer (B) encompass the highly conserved transcription factor binding motifs of NAGS. The motif colors reflect the colors used in figures 4A and B. Assays followed by immunoblot confirmed binding of Sp1 and CREB, but not C/EBP within the promoter (C) and HNF-1 and NF-Y, but not SMAD3 or AP-2 within the enhancer regions (D). Lanes 1–4 represent precipitated proteins from mouse liver nuclear extract bound to biotinylated probes of the regions of interest (Lane 1), non-biotinylated probes of the regions of interest (Lane 2), biotinylated probes of non-specific regions (Lane 3), and no probe (Lane 4). Lanes 5–8 represent supernatant fluid from overnight incubation of biotinylated probes of the region of interest (Lane 5), non-biotinylated probes of the region of interest (Lane 6), biotinylated probes of the non-specific regions (Lane 7), or no probe (Lane 8). Immunoblots are representative of at least three replicate experiments.

doi:10.1371/journal.pone.0029527.g005

To validate computational strategy for identification of transcription factor binding sites, the enhancers of human, chimpanzee, dog, mouse, and rat Cps1 were analyzed using CLOVER, and the experimentally identified binding motifs for C/EBP, CREB, GR, AP-1 and HNF-3 [55], [56], [57] were detected along with additional unreported motifs for HNF-4, AR, C/EBP and HNF-3 (Figure S3, Table S5). The detection of experimentally confirmed binding motifs in CPS1 has made the use of CLOVER for bioinformatic analysis of NAGS credible.

A DNA-protein pull-down assay was devised to test the bioinformatic prediction of specific binding sites. Two biotin-labeled DNA probes for the promoter (Figure 5A) encompassed regions A and B (Lane 1 in Figure 5C) and one probe (Figure 5B) encompassed the enhancer (Lane 1 in Figure 5D). A biotinylated probe to a region upstream of the NAGS gene, lacking any highly conserved motifs (Lane 3 in Figures 5C and 5D), and non-biotinylated probes to region A or B (Lane 2 in Figures 5C and 5D) were used as negative controls. The supernatant fluid from each pull-down was included as a positive control for the presence of the transcription factor (Lanes 5–8). Intensities of bands corresponding to each transcription factor in supernatant fluids were also used as indicators of pull-down efficiency.

Factors Sp1 and CREB bound to the probe of promoter region A (Lane 1 in Figure 5C). Sp1 also bound to the probe of promoter region B (data not shown) while C/EBP did not bind to this probe (Lane 1 in Figure 5C). Within the enhancer region, transcription factors HNF-1 and NF-Y bound to the probe, however SMAD2/3 and AP2 did not (Lane 1 in Figure 5D). Binding of Sp1, CREB, C/EBP, HNF-1, NF-Y, SMAD2/3, and AP-2 was not detected in the negative controls (Lanes 2–4 in Figures 5C and 5D) while each transcription factor was detected in the positive controls of liver nuclear extract supernatants (Lanes 5–8 in Figures 5C and 5D). Each immunoblot result is representative of at least three replicate experiments.

Binding of transcription factors to the predicted motifs was also confirmed using chromatin immunoprecipitation (ChIP) followed by Real-Time PCR. Measurement compared the enrichment of target DNA regions to the negative control locus MIP-2. ChIP with Sp1 and CREB antibodies significantly enriched the NAGS promoter DNA compared to MIP-2 (p<0.005 and p<0.05, respectively; Figure 6A). ChIP with C/EBP antibody did not enrich the NAGS promoter DNA compared to the negative locus (p>0.05; Figure 6A). The NAGS enhancer was enriched in chromatin immunoprecipitated with antibodies against HNF-1 and NF-Y (p<0.005 and p<0.05, respectively; Figure 6B), but not with antibodies against AP-2 and SMAD2/3 (p>0.05; Figure 6B). Thus, Pull-down and ChIP assays confirmed that Sp1 and CREB bind along the NAGS promoter and HNF-1 and NF-Y bind along the enhancer.

thumbnail

Figure 6. Chromatin Immunoprecipitation (ChIP) results confirm transcription factor binding.

ChIP with transcription factor antibodies was compared to negative control IgG antibody. Real-Time PCR using promoter or enhancer specific primers was compared to primers for the negative locus MIP-2. The results confirmed that Sp1 and CREB but not C/EBP bind within the promoter (A) and HNF-1 and NF-Y but not AP-2 or SMAD2/3 bind within the enhancer region (B) of NAGS. Calculated error was from three replicate experiments and reported as ± SEM. One asterisk (*) indicates p<0.05 and two asterisks (**) indicate p<0.005.

doi:10.1371/journal.pone.0029527.g006

Transcription factors and binding motifs are functionally important for transcription

Reporter assays in liver hepatoma cells with mutated transcription factor binding motifs demonstrate the functional importance of each site. Following these sequence substitutions, transcription factor binding motifs were no longer detected by CLOVER (Table 2). Within the promoter, point mutations in the Sp1 binding sites decreased the expression of reporter gene by 75% (p<0.005) and point mutations in the CREB binding site resulted in a 40% decrease (p<0.005; Figure 7A). Point mutations in the HNF-1 or NF-Y binding sites, in the enhancer, decreased expression of luciferase reporter by 50% (p<0.005 for both; Figure 7B).

thumbnail

Figure 7. Transcription factors Sp1, CREB, HNF-1, and NF-Y are functionally important for stimulating expression of reporter gene transcription.

Mutagenesis of the putative transcription factor binding sites significantly decreases transcription by the promoter (A) and the enhancer with TATA promoter (B) in liver derived cells when compared to non-mutated sites. Addition of Sp1 with the promoter (A) and HNF-1 with the enhancer (B) increases transcription driven by non-mutated constructs. Calculated results are an average of three independent experiments that were each carried out in triplicate, normalized to Rluc expression, and expressed relative to the promoter for each experiment with error reported as ±SEM. Lowercase letters indicate statistically significant differences.

doi:10.1371/journal.pone.0029527.g007

While these results confirm that each motif is important for transcription, the functional importance of Sp1 and HNF-1 proteins is demonstrated by co-expression of the proteins with reporter assay constructs. Co-transfection of Sp1 expression plasmid with the NAGS promoter (4.10Prom) increases expression of luciferase more than 50% (P<0.005; Figure 7A) while co-transfection of HNF-1 expression construct with the enhancer and minimal TATA promoter (4.23Enh), increases expression of the reporter gene by 25% (p>0.05; Figure 7B) suggesting that endogenous Sp1 and, less so, HNF-1 do not saturate their binding motifs on the transfected reporter plasmids.

Reporter assays to compare the effect of the enhancer in liver, intestine and lung cells, included data that were normalized to the reporter expression driven by the NAGS promoter. While the NAGS enhancer (4.10PromEnh) increased expression of the reporter gene by 50% in liver derived cells (Figure 2A), expression of the luciferase gene did not increase in the intestine or lung derived cells (Figure 8) suggesting that the enhancer may determine tissue specificity of NAGS expression. When HNF-1 expression plasmid and 4.10PromEnh were co-transfected into intestine and lung derived cells, transcription was stimulated to levels that were not significantly different from 4.10PromEnh in liver cells (p>0.05) (Figure 8). Because intestine and lung derived cells lack HNF-1 (data not shown), this demonstrated the importance of HNF-1 and NAGS enhancer for the tissue specificity of NAGS expression.

thumbnail

Figure 8. The NAGS enhancer shows tissue specificity.

The enhancer with NAGS promoter (4.10PromEnh) increases transcription relative to the promoter in liver derived cells but not in intestine or lung derived cells (cyan bars) without the addition of HNF-1 protein (teal bars). Calculated results are an average of three independent experiments that were carried out in triplicate, normalized to Rluc expression, and expressed relative to the promoter for each experiment with error reported as ±SEM. Lowercase letters indicate statistically significant differences.

doi:10.1371/journal.pone.0029527.g008

Discussion

In this study we used bioinformatic analyses to predict regulatory regions based on the hypothesis that non-coding DNA sequences that are highly conserved between species are important for gene regulation. Multiple pair-wise BLAST alignments and sequence alignment from the UCSC genome browser were used to identify two conserved regions within NAGS, which were determined to be a promoter and an enhancer. The efficacy of this method was confirmed by successful identification of the experimentally identified promoter and −6.3 kb enhancer [25], [31], along with three additional highly conserved regions, in the non-coding region upstream of CPS1. It should be noted that the high stringency of our BLAST analysis (80% identity and at least 100 bp of aligned sequence in four or more species) was selected to identify conserved regions that could support multiple binding sites where complexes of transcription factors may form [25], [58]. This may have caused us to overlook species specific or isolated binding motifs, such as the recently identified FXR binding site [59].

The reporter assay results confirm that the two highly conserved regions within 1 kb and 3 kb upstream of the translational start site function as promoter and enhancer, respectively. The promoter activates expression of the luciferase reporter gene and we therefore infer that it will activate transcription of NAGS in vivo. Similarly, the enhancer in either orientation increases expression of luciferase by approximately 50% relative to the promoter alone, suggesting that it stimulates NAGS transcription as well. The relatively small but significant effect of the enhancer could be due to spacing differences between the genomic NAGS promoter and enhancer and their spacing in the reporter constructs. Alternatively, while HepG2 cells express transcription factors that we identified using bioinformatic tools, the NAGS enhancer may bind additional factors, absent in HepG2 cells, and have larger effect in vivo than in cultured cells. Another explanation for the relatively small effect of the NAGS enhancer is the possible presence of a proximal enhancer within the region we termed the promoter. Additional experiments are necessary to distinguish between these two possibilities.

Our analysis of the NAGS transcriptional start sites identified multiple TSS that may be species and tissue specific. While the function of each TSS is unknown, these results are consistent with transcription initiation by Sp1 [16], [60], [61], and future experiments may find that they are involved in transcriptional control for tissue specific expression, developmental-stage specific expression, quantitatively different levels of mRNA expression, or may even determine the transcript stability [62].

After we confirmed that the promoter and enhancer initiate and increase transcription, we looked for transcription factors that bind and regulate NAGS in these regions. By filtering for the highly over-represented and spatially conserved binding sites, relative to the translational start codon, we identified Sp1, CREB, and C/EBP in the promoter and HNF-1 AP-2, NF-Y, and SMAD-3 in the enhancer as transcription factors that could bind to the NAGS upstream region. This filtering method was confirmed by analysis of the −6.3 kb enhancer of CPS1 in which binding sites for the previously published C/EBP, CREB, GR, and HNF-3 were identified.

The protein-DNA pull down assays, designed to test which transcription factors among a pool of nuclear proteins bind to amplified sequence of conserved upstream DNA, confirmed that Sp1, CREB, HNF-1 and NF-Y bind to NAGS promoter and enhancer, while we could not detect binding of C/EBP, AP-2 and SMAD3 (Figure 5). We initially used 60 bp probes encompassing a specific binding motif for the protein–DNA pull down assays. However, probes encompassing the entire region were better able to bind transcription factors (data not shown), suggesting that binding is facilitated by interactions with DNA sequences outside predicted binding sites and possibly other transcription factors and co-activators. ChIP analysis was used to confirm binding of the predicted transcription factors to the DNA regions of interest under physiological conditions. ChIP and DNA-pull down assays confirmed that Sp1 and CREB bind to the promoter and HNF-1 and NF-Y bind to the enhancer of NAGS (Figures 5 and 6), while reporter assays demonstrated the functional importance of each binding motif by a decrease in transcription following mutagenesis of the binding sites (Figure 7).

Furthermore, we have demonstrated that Sp1 and HNF-1 are important for stimulation of transcription of NAGS and that HNF-1 determines tissue specificity of NAGS expression. In the liver derived cell line, co-transfection of either Sp1 or HNF-1 expression plasmids with reporter constructs containing the NAGS promoter and enhancer led to increased expression of the reporter gene (Figure 7) suggesting that these two transcription factors regulate expression of NAGS in the liver. In the lung and intestine derived cell lines, expression of HNF-1 was sufficient to activate expression of reporter gene in constructs containing NAGS enhancer and promoter (Figure 8). This suggests that HNF-1 binding to the NAGS enhancer determines tissue specificity of NAGS expression. Testing the effect of over-expression of CREB protein was hindered by its capacity to homo- and heterodimerize with multiple partners [63], [64]. The effect of NF-Y was not tested because this transcription factor is a heterotrimer [65] and its co-expression with reporter plasmids would require stable expression of NF-Y subunit proteins by in vitro cell culture before reporter plasmids can be transfected and assayed for NF-Y effect on transcription.

From the data provided herein, we can speculate on the potential role these factors play in regulating NAGS transcription. First, in the absence of a canonical TATA-box, transcription initiated by Sp1 often results in multiple transcriptional start sites [66], [67]. Sp1 is a strong activator of transcription [16], [68], [69], [70], [71] and when multiple Sp1 sites are present, as in NAGS, multiple Sp1 proteins can form complexes with each other and synergistically activate transcription [16], [69]. Because transcription is significantly increased by co-expression with Sp1 protein and decreased following mutation of the Sp1 binding sites, Sp1 may prove to be the activator of NAGS transcription, similar to its role for ASS, ASL and ARG1 [15], [25].

Second, studies have shown that glucagon and second messenger cAMP trigger a cascade that phosphorylates CREB and allows for DNA binding and activation of transcription [72], [73]. In CPS1 and ASS, CREB stimulates transcription upon glucagon signaling [15], [31]. Decrease in transcription following CREB mutation and the close proximity of Sp1 and CREB binding sites among the TSS suggests that the transcription initiation machinery may be recruited by these factors, and future research should examine this postulate.

Our experiments and other studies [74] confirm the role of HNF-1 in NAGS expression. HNF-1 is essential for stimulation of NAGS expression by its enhancer. This factor is in part regulated by HNF-3, HNF-4, and C/EBP, each of which are known to regulate other urea cycle genes [75], [76], [77]. Future research will focus on the mechanism of control between these factors, HNF-1, and NAGS. Our study has also shown that NF-Y is an activator of NAGS expression, and future studies will focus on the exact mechanism of its function in this context.

The human NAGS gene on the forward strand of chromosome 17 partially overlaps with the peptide YY (PYY) gene, which is on the reverse strand. This overlap was identified with a PYY cDNA isolated from a brain astrocytoma cDNA library that has an 80 nucleotide long exon located between regions A and B of the NAGS promoter [78], [79] (Figure 1). Other full-length PYY transcripts initiate about 500 bp upstream of the PYY coding region, which is located 51 kb upstream of the NAGS translation initiation codon. Recent analysis of human transcripts revealed that many protein coding loci are associated with at least one transcript that initiates from a distal site [80], but the significance or function of these transcripts remains to be elucidated. Partial overlap between human NAGS and PYY genes raises the interesting possibility that these two genes share cis-acting regulatory elements and might be co-regulated [79], [81]. The mechanism of co-regulation of human NAGS and PYY is likely to be complex because of their differing tissue expression patterns [1], [82], [83], [84] including different cell types within the intestine. PYY is expressed in the intestinal neuroendocrine cells [85], [86] while epithelial cells in the small intestine express NAGS [87], [88], together with OTC and CPS1 [13], [89]. Inspection of the transcription factor binding track of the UCSC genome browser revealed two binding sites for the CTCF transcription repressor between NAGS and PYY genes; they are located approximately 9.5 and 21 kb upstream of the NAGS coding region. The CTCF binding sites could act as chromatin insulators [90], [91], [92] and either block regulation of PYY by the NAGS enhancer or enable cell type specific regulation of each gene by the NAGS enhancer and promoter. Our results show that the NAGS promoter in the reverse orientation does not activate transcription of the reporter gene in liver derived cells (Figure 2), but this does not preclude transcription activation in other cell types, not tested in this study. It is possible that the NAGS promoter, enhancer, or other NAGS regions, regulates expression of PYY [84], and reporter assays in tissues and cultured cells which express PYY would test this hypothesis.

While regulation of NAGS by Sp1, CREB, HNF-1, NF-Y, and factors that regulate them, requires additional study, identification of regions that regulate human NAGS and OTC have enabled diagnosis of patients with clinical symptoms of urea cycle disorders, but lacking disease causing mutations in the coding regions of the genes [93], [94]. Recently, we identified a patient with a mutation in the enhancer of NAGS and confirmed the diagnosis of NAGS deficiency by showing that the mutation significantly decreases transcription of NAGS [93]. This example suggests that identification of regulatory regions within genes will lead to more and better diagnoses of urea cycle disorders and other genetic diseases and to accurate genetic counseling.

In conclusion, this study identified a promoter and a tissue specific enhancer of NAGS and functionally relevant transcription factor binding motifs within these regions. The results show that Sp1 and CREB bind to the NAGS promoter, suggesting that glucagon and cAMP signaling may regulate the expression of NAGS. Within the enhancer, HNF-1 may be an important factor in the coordinated regulation of this urea cycle gene transcription through its interaction with HNF-3, HNF-4 and C/EBP while the role of NF-Y is less clear considering that NF-Y may function as an activator or repressor. While additional studies will be needed to further define the roles of these factors, these results contain the first thorough analysis of NAGS and suggest networks of control between signaling cascades, NAGS and the coordinated regulation of the other urea cycle genes.

Supporting Information

Figure S1.

Regions Upstream of mammalian CPS1 genes are highly conserved. Three new highly conserved regions were identified within 15 kb 5′ of the CPS1 translational start site. Conservation algorithms phastCons (green) and phyloP (blue) from the UCSC genome browser indicate regions that are highly conserved across all mammals (A). Pair-wise blast analysis of human, chimpanzee, dog, mouse, and rat 5′ non-coding region of CPS1 were used to identify two known and three previously unknown regions of high conservation, referred to enhancer/repressor regions A, B, and C. Highly conserved regions within the CPS1 5′ non-coding sequence include the proximal promoter, region A, the -enhancer, region B, and region C.

doi:10.1371/journal.pone.0029527.s001

(TIF)

Figure S2.

Highly conserved regulatory regions, upstream of the mouse Nags gene, function as promoter and enhancer elements. Mouse promoter (m4.10Prom), promoter and enhancer (m4.10PromEnh), and enhancer with TATA promoter (m4.23Enh) stimulated transcription while enhancer lacking a promoter (m4.10Enh) did not in liver cells. Calculated results are an average of three independent experiments that were carried out in triplicate, normalized to Rluc expression, and expressed relative to the promoter for each experiment with error reported as ±SEM.

doi:10.1371/journal.pone.0029527.s002

(TIF)

Figure S3.

Novel transcription factor binding motifs, in the enhancer region of CPS1, were identified using CLOVER. Several highly conserved transcription factor binding sites were present in the enhancer region. An asterisk denotes an experimentally verified transcription factor binding site. All motifs were spatially conserved between mammalian species.

doi:10.1371/journal.pone.0029527.s003

(TIF)

Table S1.

Sequences of primers that were used to amplify human or mouse DNA by PCR for insertion of the promoter and enhancer regions into sequencing and reporter assay vectors.

doi:10.1371/journal.pone.0029527.s004

(DOCX)

Table S2.

Primer sequences used to determine transcription start sites of NAGS with 5′ RACE. Primers were designed according to manufacturer's instructions and used to determine transcription start sites of human and mouse NAGS in liver and small intestine RNA using 5′ RACE.

doi:10.1371/journal.pone.0029527.s005

(DOCX)

Table S3.

Primer sequences used to generate DNA probes of the specified regions of mNags. Primers were used to generate DNA probes, by PCR, of the promoter, enhancer, or non-specific specified regions of mNags.

doi:10.1371/journal.pone.0029527.s006

(DOCX)

Table S4.

Primer sequences used for quantitative real-time PCR analysis of chromatin immunoprecipitation samples.

doi:10.1371/journal.pone.0029527.s007

(DOCX)

Table S5.

Results of CLOVER analysis of the enhancer region with sequence information for human and mouse CPS1. Results were filtered to exclude motifs for transcription factors that are not expressed in the liver.

doi:10.1371/journal.pone.0029527.s008

(DOCX)

Table S6.

Results of CLOVER analysis of the promoter region with sequence information for human and mouse NAGS. Results were filtered to exclude motifs for transcription factors that are not expressed in liver.

doi:10.1371/journal.pone.0029527.s009

(DOCX)

Table S7.

Results of CLOVER analysis of the enhancer region with sequence information for human and mouse NAGS. Results were filtered to exclude motifs for transcription factors that are not expressed in the liver.

doi:10.1371/journal.pone.0029527.s010

(DOCX)

Acknowledgments

We would like to thank Dr. Marshall Summar for providing HepG2 cells and Dr. Mary Rose for providing A549 cells.

Author Contributions

Analyzed the data: SKH. Contributed reagents/materials/analysis tools: LM-R. Wrote the paper: SKH. Designed the experiments: SKH. Critically reviewed the manuscript: MT. Conceived the study and reviewed the manuscript: LC. Performed reporter assays, 5′-RACE, DNA pill-down assays, ChIP, bioinformatic analysis and wrote the paper: SKH. Carried out and analyzed bioinformatic analysis of the NAGS upstream regulatory region: GYL MP SS LM-R.

References

  1. 1. Caldovic L, Morizono H, Gracia Panglao M, Gallegos R, Yu X, et al. (2002) Cloning and expression of the human N-acetylglutamate synthase gene. Biochem Biophys Res Commun 299: 581–586.
  2. 2. Caldovic L, Morizono H, Yu X, Thompson M, Shi D, et al. (2002) Identification, cloning and expression of the mouse N-acetylglutamate synthase gene. Biochem J 364: 825–831.
  3. 3. Schimke RT (1962) Differential effects of fasting and protein-free diets on levels of urea cycle enzymes in rat liver. J Biol Chem 237: 1921–1924.
  4. 4. Kawamoto S, Ishida H, Mori M, Tatibana M (1982) Regulation of N-acetylglutamate synthetase in mouse liver. Postprandial changes in sensitivity to activation by arginine. Eur J Biochem 123: 637–641.
  5. 5. Tatibana M, Kawamoto S, Sonoda T, Mori M (1982) Enzyme regulation of n-acetylglutamate synthesis in mouse and rat liver. Adv Exp Med Biol 153: 207–216.
  6. 6. Shigesada K, Tatibana M (1978) N-Acetylglutamate synthetase from rat-liver mitochondria. Partial purification and catalytic properties. Eur J Biochem 84: 285–291.
  7. 7. Goping IS, Shore GC (1994) Interactions between repressor and anti-repressor elements in the carbamyl phosphate synthetase I promoter. J Biol Chem 269: 3891–3896.
  8. 8. Howell BW, Lagace M, Shore GC (1989) Activity of the carbamyl phosphate synthetase I promoter in liver nuclear extracts is dependent on a cis-acting C/EBP recognition element. Mol Cell Biol 9: 2928–2933.
  9. 9. Lagace M, Goping IS, Mueller CR, Lazzaro M, Shore GC (1992) The carbamyl phosphate synthetase promoter contains multiple binding sites for C/EBP-related proteins. Gene 118: 231–238.
  10. 10. Schoneveld OJ, Gaemers IC, Hoogenkamp M, Lamers WH (2005) The role of proximal-enhancer elements in the glucocorticoid regulation of carbamoylphosphate synthetase gene transcription from the upstream response unit. Biochimie 87: 1033–1040.
  11. 11. Kimura A, Nishiyori A, Murakami T, Tsukamoto T, Hata S, et al. (1993) Chicken ovalbumin upstream promoter-transcription factor (COUP-TF) represses transcription from the promoter of the gene for ornithine transcarbamylase in a manner antagonistic to hepatocyte nuclear factor-4 (HNF-4). J Biol Chem 268: 11125–11133.
  12. 12. Kimura T, Chowdhury S, Tanaka T, Shimizu A, Iwase K, et al. (2001) CCAAT/enhancer-binding protein beta is required for activation of genes for ornithine cycle enzymes by glucocorticoids and glucagon in primary-cultured hepatocytes. FEBS Lett 494: 105–111.
  13. 13. Murakami T, Nishiyori A, Takiguchi M, Mori M (1990) Promoter and 11-kilobase upstream enhancer elements responsible for hepatoma cell-specific expression of the rat ornithine transcarbamylase gene. Mol Cell Biol 10: 1180–1191.
  14. 14. Nishiyori A, Tashiro H, Kimura A, Akagi K, Yamamura K, et al. (1994) Determination of tissue specificity of the enhancer by combinatorial operation of tissue-enriched transcription factors. Both HNF-4 and C/EBP beta are required for liver-specific activity of the ornithine transcarbamylase enhancer. J Biol Chem 269: 1323–1331.
  15. 15. Guei TR, Liu MC, Yang CP, Su TS (2008) Identification of a liver-specific cAMP response element in the human argininosuccinate synthetase gene. Biochem Biophys Res Commun 377: 257–261.
  16. 16. Anderson GM, Freytag SO (1991) Synergistic activation of a human promoter in vivo by transcription factor Sp1. Mol Cell Biol 11: 1935–1943.
  17. 17. Boyce FM 3rd, Pogulis RJ, Freytag SO (1989) Paradoxical regulation of human argininosuccinate synthetase cDNA minigene in opposition to endogenous gene: evidence for intragenic control sequences. Somat Cell Mol Genet 15: 123–129.
  18. 18. Matsubasa T, Takiguchi M, Matsuda I, Mori M (1994) Rat argininosuccinate lyase promoter: the dyad-symmetric CCAAT box sequence CCAATTGG in the promoter is recognized by NF-Y. J Biochem 116: 1044–1055.
  19. 19. Dorn A, Bollekens J, Staub A, Benoist C, Mathis D (1987) A multiplicity of CCAAT box-binding proteins. Cell 50: 863–872.
  20. 20. Hooft van Huijsduijnen R, Li XY, Black D, Matthes H, Benoist C, et al. (1990) Co-evolution from yeast to mouse: cDNA cloning of the two NF-Y (CP-1/CBF) subunits. EMBO J 9: 3119–3127.
  21. 21. Santoro C, Mermod N, Andrews PC, Tjian R (1988) A family of human CCAAT-box-binding proteins active in transcription and DNA replication: cloning and expression of multiple cDNAs. Nature 334: 218–224.
  22. 22. Takiguchi M, Mori M (1991) In vitro analysis of the rat liver-type arginase promoter. J Biol Chem 266: 9186–9193.
  23. 23. Morris SM Jr (2002) Regulation of enzymes of the urea cycle and arginine metabolism. Annu Rev Nutr 22: 87–105.
  24. 24. Morris SM Jr, Moncman CL, Rand KD, Dizikes GJ, Cederbaum SD, et al. (1987) Regulation of mRNA levels for five urea cycle enzymes in rat liver by diet, cyclic AMP, and glucocorticoids. Arch Biochem Biophys 256: 343–353.
  25. 25. Takiguchi M, Mori M (1995) Transcriptional regulation of genes for ornithine cycle enzymes. Biochem J 312(Pt 3): 649–659.
  26. 26. Nebes VL, Morris SM Jr (1988) Regulation of messenger ribonucleic acid levels for five urea cycle enzymes in cultured rat hepatocytes. Requirements for cyclic adenosine monophosphate, glucocorticoids, and ongoing protein synthesis. Mol Endocrinol 2: 444–451.
  27. 27. Ryall JC, Quantz MA, Shore GC (1986) Rat liver and intestinal mucosa differ in the developmental pattern and hormonal regulation of carbamoyl-phosphate synthetase I and ornithine carbamoyl transferase gene expression. Eur J Biochem 156: 453–458.
  28. 28. Schimke RT (1963) Studies on factors affecting the levels of urea cycle enzymes in rat liver. J Biol Chem 238: 1012–1018.
  29. 29. Hazra A, DuBois DC, Almon RR, Snyder GH, Jusko WJ (2008) Pharmacodynamic modeling of acute and chronic effects of methylprednisolone on hepatic urea cycle genes in rats. Gene Regul Syst Bio 2: 1–19.
  30. 30. Abdullah Abu Musa DM, Kobayashi K, Yasuda I, Iijima M, Christoffels VM, et al. (1999) Involvement of a cis-acting element in the suppression of carbamoyl phosphate synthetase I gene expression in the liver of carnitine-deficient mice. Mol Genet Metab 68: 346–356.
  31. 31. Schoneveld OJ, Hoogenkamp M, Stallen JM, Gaemers IC, Lamers WH (2007) cyclicAMP and glucocorticoid responsiveness of the rat carbamoylphosphate synthetase gene requires the interplay of upstream regulatory units. Biochimie 89: 574–580.
  32. 32. Murakami T, Takiguchi M, Inomoto T, Yamamura K, Mori M (1989) Tissue- and developmental stage-specific expression of the rat ornithine carbamoyltransferase gene in transgenic mice. Dev Genet 10: 393–401.
  33. 33. Sladek FM, Zhong WM, Lai E, Darnell JE Jr (1990) Liver-enriched transcription factor HNF-4 is a novel member of the steroid hormone receptor superfamily. Genes Dev 4: 2353–2365.
  34. 34. Boyce FM, Anderson GM, Rusk CD, Freytag SO (1986) Human argininosuccinate synthetase minigenes are subject to arginine-mediated repression but not to trans induction. Mol Cell Biol 6: 1244–1252.
  35. 35. Boyce FM 3rd, Freytag SO (1989) Regulation of human argininosuccinate synthetase gene: induction by positive-acting nuclear mechanism in canavanine-resistant cell variants. Somat Cell Mol Genet 15: 113–121.
  36. 36. Jackson MJ, Allen SJ, Beaudet AL, O'Brien WE (1988) Metabolite regulation of argininosuccinate synthetase in cultured human cells. J Biol Chem 263: 16388–16394.
  37. 37. Sunyakumthorn P, Boonsaen T, Boonsaeng V, Wallace JC, Jitrapakdee S (2005) Involvement of specific proteins (Sp1/Sp3) and nuclear factor Y in basal transcription of the distal promoter of the rat pyruvate carboxylase gene in beta-cells. Biochem Biophys Res Commun 329: 188–196.
  38. 38. Snodgrass PJ (1991) Dexamethasone and glucagon cause synergistic increases of urea cycle enzyme activities in livers of normal but not adrenalectomized rats. Enzyme 45: 30–38.
  39. 39. Altschul SF, Madden TL, Schaffer AA, Zhang J, Zhang Z, et al. (1997) Gapped BLAST and PSI-BLAST: a new generation of protein database search programs. Nucleic Acids Res 25: 3389–3402.
  40. 40. Frith MC, Fu Y, Yu L, Chen JF, Hansen U, et al. (2004) Detection of functional DNA motifs via statistical over-representation. Nucleic Acids Res 32: 1372–1381.
  41. 41. Marino-Ramirez L, Spouge JL, Kanga GC, Landsman D (2004) Statistical analysis of over-represented words in human promoter sequences. Nucleic Acids Res 32: 949–958.
  42. 42. Chenna R, Sugawara H, Koike T, Lopez R, Gibson TJ, et al. (2003) Multiple sequence alignment with the Clustal series of programs. Nucleic Acids Res 31: 3497–3500.
  43. 43. Kim HJ, Ko MS, Kim HK, Cho WJ, Lee SH, et al. Transcription factor Sp1 regulates basal transcription of the human DRG2 gene. Biochim Biophys Acta 1809: 184–190.
  44. 44. Zhang W, Tian Z, Sha S, Cheng LY, Philipsen S, et al. Functional and sequence analysis of human neuroglobin gene promoter region. Biochim Biophys Acta 1809: 236–244.
  45. 45. Convertini P, Infantino V, Bisaccia F, Palmieri F, Iacobazzi VRole of FOXA and Sp1 in mitochondrial acylcarnitine carrier gene expression in different cell lines. Biochem Biophys Res Commun 404: 376–381.
  46. 46. Michels AJ, Hagen TM (2009) Hepatocyte nuclear factor 1 is essential for transcription of sodium-dependent vitamin C transporter protein 1. Am J Physiol Cell Physiol 297: C1220–1227.
  47. 47. Wang Z, Burke PAHepatocyte nuclear factor-4alpha interacts with other hepatocyte nuclear factors in regulating transthyretin gene expression. FEBS J 277: 4066–4075.
  48. 48. Tue NT, Yoshioka Y, Yamaguchi MNF-Y transcriptionally regulates the Drosophila p53 gene. Gene 473: 1–7.
  49. 49. Pallai R, Simpkins H, Chen J, Parekh HKThe CCAAT box binding transcription factor, nuclear factor-Y (NF-Y) regulates transcription of human aldo-keto reductase 1C1 (AKR1C1) gene. Gene 459: 11–23.
  50. 50. Xiang H, Wang J, Boxer LM (2006) Role of the cyclic AMP response element in the bcl-2 promoter in the regulation of endogenous Bcl-2 expression and apoptosis in murine B cells. Mol Cell Biol 26: 8599–8606.
  51. 51. Callens N, Baert JL, Monte D, Sunesen M, Van Lint C, et al. (2003) Transcriptional regulation of the murine brca2 gene by CREB/ATF transcription factors. Biochem Biophys Res Commun 312: 702–707.
  52. 52. Deng WG, Zhu Y, Montero A, Wu KK (2003) Quantitative analysis of binding of transcription factor complex to biotinylated DNA probe by a streptavidin-agarose pulldown assay. Anal Biochem 323: 12–18.
  53. 53. Pollard KS, Hubisz MJ, Rosenbloom KR, Siepel ADetection of nonneutral substitution rates on mammalian phylogenies. Genome Res 20: 110–121.
  54. 54. Siepel A, Bejerano G, Pedersen JS, Hinrichs AS, Hou M, et al. (2005) Evolutionarily conserved elements in vertebrate, insect, worm, and yeast genomes. Genome Res 15: 1034–1050.
  55. 55. Christoffels VM, van den Hoff MJ, Lamers MC, van Roon MA, de Boer PA, et al. (1996) The upstream regulatory region of the carbamoyl-phosphate synthetase I gene controls its tissue-specific, developmental, and hormonal regulation in vivo. J Biol Chem 271: 31243–31250.
  56. 56. Christoffels VM, Grange T, Kaestner KH, Cole TJ, Darlington GJ, et al. (1998) Glucocorticoid receptor, C/EBP, HNF3, and protein kinase A coordinately activate the glucocorticoid response unit of the carbamoylphosphate synthetase I gene. Mol Cell Biol 18: 6305–6315.
  57. 57. Christoffels VM, van den Hoff MJ, Moorman AF, Lamers WH (1995) The far-upstream enhancer of the carbamoyl-phosphate synthetase I gene is responsible for the tissue specificity and hormone inducibility of its expression. J Biol Chem 270: 24932–24940.
  58. 58. Klein H, Vingron M (2007) Using transcription factor binding site co-occurrence to predict regulatory regions. Genome Inform 18: 109–118.
  59. 59. Renga B, Mencarelli A, Cipriani S, D'Amore C, Zampella A, et al. The nuclear receptor FXR regulates hepatic transport and metabolism of glutamine and glutamate. Biochim Biophys Acta 1812: 1522–1531.
  60. 60. Emami KH, Burke TW, Smale ST (1998) Sp1 activation of a TATA-less promoter requires a species-specific interaction involving transcription factor IID. Nucleic Acids Res 26: 839–846.
  61. 61. Muckenfuss H, Kaiser JK, Krebil E, Battenberg M, Schwer C, et al. (2007) Sp1 and Sp3 regulate basal transcription of the human APOBEC3G gene. Nucleic Acids Res 35: 3784–3796.
  62. 62. Schibler U, Sierra F (1987) Alternative promoters in developmental gene expression. Annu Rev Genet 21: 237–257.
  63. 63. Hai T, Hartman MG (2001) The molecular biology and nomenclature of the activating transcription factor/cAMP responsive element binding family of transcription factors: activating transcription factor proteins and homeostasis. Gene 273: 1–11.
  64. 64. De Cesare D, Sassone-Corsi P (2000) Transcriptional regulation by cyclic AMP-responsive factors. Prog Nucleic Acid Res Mol Biol 64: 343–369.
  65. 65. Matuoka K, Yu Chen K (1999) Nuclear factor Y (NF-Y) and cellular senescence. Exp Cell Res 253: 365–371.
  66. 66. Juang HH, Costello LC, Franklin RB (1995) Androgen modulation of multiple transcription start sites of the mitochondrial aspartate aminotransferase gene in rat prostate. J Biol Chem 270: 12629–12634.
  67. 67. Pave-Preux M, Aggerbeck M, Veyssier C, Bousquet-Lemercier B, Hanoune J, et al. (1990) Hormonal discrimination among transcription start sites of aspartate aminotransferase. J Biol Chem 265: 4444–4448.
  68. 68. Kadonaga JT, Courey AJ, Ladika J, Tjian R (1988) Distinct regions of Sp1 modulate DNA binding and transcriptional activation. Science 242: 1566–1570.
  69. 69. Li L, He S, Sun JM, Davie JR (2004) Gene regulation by Sp1 and Sp3. Biochem Cell Biol 82: 460–471.
  70. 70. Solomon SS, Majumdar G, Martinez-Hernandez A, Raghow R (2008) A critical role of Sp1 transcription factor in regulating gene expression in response to insulin and other hormones. Life Sci 83: 305–312.
  71. 71. Wierstra I (2008) Sp1: emerging roles–beyond constitutive activation of TATA-less housekeeping genes. Biochem Biophys Res Commun 372: 1–13.
  72. 72. Montminy M, Koo SH, Zhang X (2004) The CREB family: key regulators of hepatic metabolism. Ann Endocrinol (Paris) 65: 73–75.
  73. 73. Mayr B, Montminy M (2001) Transcriptional regulation by the phosphorylation-dependent factor CREB. Nat Rev Mol Cell Biol 2: 599–609.
  74. 74. Heibel SK, Ah Mew N, Caldovic L, Daikhin Y, Yudkoff M, et al. N-carbamylglutamate enhancement of ureagenesis leads to discovery of a novel deleterious mutation in a newly defined enhancer of the NAGS gene and to effective therapy. Hum Mutat 32: 1153–1160.
  75. 75. Kuo CJ, Conley PB, Chen L, Sladek FM, Darnell JE Jr, et al. (1992) A transcriptional hierarchy involved in mammalian cell-type specification. Nature 355: 457–461.
  76. 76. Sladek FM (1993) Orphan receptor HNF-4 and liver-specific gene expression. Receptor 3: 223–232.
  77. 77. Ktistaki E, Talianidis I (1997) Modulation of hepatic gene expression by hepatocyte nuclear factor 1. Science 277: 109–112.
  78. 78. Strausberg RL, Feingold EA, Grouse LH, Derge JG, Klausner RD, et al. (2002) Generation and initial analysis of more than 15,000 full-length human and mouse cDNA sequences. Proc Natl Acad Sci U S A 99: 16899–16903.
  79. 79. Lomenick JP, Melguizo MS, Mitchell SL, Summar ML, Anderson JW (2009) Effects of meals high in carbohydrate, protein, and fat on ghrelin and peptide YY secretion in prepubertal children. J Clin Endocrinol Metab 94: 4463–4471.
  80. 80. Birney E, Stamatoyannopoulos JA, Dutta A, Guigo R, Gingeras TR, et al. (2007) Identification and analysis of functional elements in 1% of the human genome by the ENCODE pilot project. Nature 447: 799–816.
  81. 81. Mitchell S, Murdock D, Summar M (2008) Plasma peptide tyrosine tyrosine (PYY) levels are increased in urea cycle disorder patients. Mol Gen Metab 93: 258 (abstract).
  82. 82. Neill MA, Aschner J, Barr F, Summar ML (2009) Quantitative RT-PCR comparison of the urea and nitric oxide cycle gene transcripts in adult human tissues. Mol Genet Metab 97: 121–127.
  83. 83. Myrsen-Axcrona U, Ekblad E, Sundler F (1997) Developmental expression of NPY, PYY and PP in the rat pancreas and their coexistence with islet hormones. Regul Pept 68: 165–175.
  84. 84. Ekblad E, Sundler F (2002) Distribution of pancreatic polypeptide and peptide YY. Peptides 23: 251–261.
  85. 85. Lundberg JM, Tatemoto K, Terenius L, Hellstrom PM, Mutt V, et al. (1982) Localization of peptide YY (PYY) in gastrointestinal endocrine cells and effects on intestinal blood flow and motility. Proc Natl Acad Sci U S A 79: 4471–4475.
  86. 86. Lukinius AI, Ericsson JL, Lundqvist MK, Wilander EM (1986) Ultrastructural localization of serotonin and polypeptide YY (PYY) in endocrine cells of the human rectum. J Histochem Cytochem 34: 719–726.
  87. 87. Geng M, Li T, Kong X, Song X, Chu W, et al. Reduced expression of intestinal N-acetylglutamate synthase in suckling piglets: a novel molecular mechanism for arginine as a nutritionally essential amino acid for neonates. Amino Acids 40: 1513–1522.
  88. 88. Uchiyama C, Mori M, Tatibana M (1981) Subcellular localization and properties of N-acetylglutamate synthase in rat small intestinal mucosa. J Biochem 89: 1777–1786.
  89. 89. Dubois N, Cavard C, Chasse JF, Kamoun P, Briand P (1988) Compared expression levels of ornithine transcarbamylase and carbamylphosphate synthetase in liver and small intestine of normal and mutant mice. Biochim Biophys Acta 950: 321–328.
  90. 90. Ishihara K, Oshimura M, Nakao M (2006) CTCF-dependent chromatin insulator is linked to epigenetic remodeling. Mol Cell 23: 733–742.
  91. 91. Renda M, Baglivo I, Burgess-Beusse B, Esposito S, Fattorusso R, et al. (2007) Critical DNA binding interactions of the insulator protein CTCF: a small number of zinc fingers mediate strong binding, and a single finger-DNA interaction controls binding at imprinted loci. J Biol Chem 282: 33336–33345.
  92. 92. Majumder P, Gomez JA, Chadwick BP, Boss JM (2008) The insulator factor CTCF controls MHC class II gene expression and is required for the formation of long-distance chromatin interactions. J Exp Med 205: 785–798.
  93. 93. Heibel SK, Ah Mew N, Caldovic L, Daikhin Y, Yudkoff M, et al. (2011) N-carbamylglutamate enhancement of ureagenesis leads to discovery of a novel deleterious mutation in a newly defined enhancer of the NAGS gene and to effective therapy. Hum Mutat 32: 1153–1160.
  94. 94. Luksan O, Jirsa M, Eberova J, Minks J, Treslova H, et al. (2010) Disruption of OTC promoter-enhancer interaction in a patient with symptoms of ornithine carbamoyltransferase deficiency. Hum Mutat 31: E1294–1303.