Role of upstream stimulatory factor 2 in glutamate dehydrogenase gene transcription

in Journal of Molecular Endocrinology
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  • 1 Secció de Bioquímica i Biologia Molecular, Departament de Bioquímica i Fisiologia, Facultat de Farmàcia i Ciències de l’Alimentació, Universitat de Barcelona, Barcelona, Spain
  • 2 Servei de Bioquímica Clínica, Hospital Sant Joan de Déu, Barcelona, Spain

Glutamate dehydrogenase (Gdh) plays a central role in ammonia detoxification by catalysing reversible oxidative deamination of l-glutamate into α-ketoglutarate using NAD+ or NADP+ as cofactor. To gain insight into transcriptional regulation of glud, the gene that codes for Gdh, we isolated and characterised the 5′ flanking region of glud from gilthead sea bream (Sparus aurata). In addition, tissue distribution, the effect of starvation as well as short- and long-term refeeding on Gdh mRNA levels in the liver of S. aurata were also addressed. 5′-Deletion analysis of glud promoter in transiently transfected HepG2 cells, electrophoretic mobility shift assays, chromatin immunoprecipitation (ChIP) and site-directed mutagenesis allowed us to identify upstream stimulatory factor 2 (Usf2) as a novel factor involved in the transcriptional regulation of glud. Analysis of tissue distribution of Gdh and Usf2 mRNA levels by reverse transcriptase-coupled quantitative real-time PCR (RT-qPCR) showed that Gdh is mainly expressed in the liver of S. aurata, while Usf2 displayed ubiquitous distribution. RT-qPCR and ChIP assays revealed that long-term starvation down-regulated the hepatic expression of Gdh and Usf2 to similar levels and reduced Usf2 binding to glud promoter, while refeeding resulted in a slow but gradual restoration of both Gdh and Usf2 mRNA abundance. Herein, we demonstrate that Usf2 transactivates S. aurata glud by binding to an E-box located in the proximal region of glud promoter. In addition, our findings provide evidence for a new regulatory mechanism involving Usf2 as a key factor in the nutritional regulation of glud transcription in the fish liver.

Abstract

Glutamate dehydrogenase (Gdh) plays a central role in ammonia detoxification by catalysing reversible oxidative deamination of l-glutamate into α-ketoglutarate using NAD+ or NADP+ as cofactor. To gain insight into transcriptional regulation of glud, the gene that codes for Gdh, we isolated and characterised the 5′ flanking region of glud from gilthead sea bream (Sparus aurata). In addition, tissue distribution, the effect of starvation as well as short- and long-term refeeding on Gdh mRNA levels in the liver of S. aurata were also addressed. 5′-Deletion analysis of glud promoter in transiently transfected HepG2 cells, electrophoretic mobility shift assays, chromatin immunoprecipitation (ChIP) and site-directed mutagenesis allowed us to identify upstream stimulatory factor 2 (Usf2) as a novel factor involved in the transcriptional regulation of glud. Analysis of tissue distribution of Gdh and Usf2 mRNA levels by reverse transcriptase-coupled quantitative real-time PCR (RT-qPCR) showed that Gdh is mainly expressed in the liver of S. aurata, while Usf2 displayed ubiquitous distribution. RT-qPCR and ChIP assays revealed that long-term starvation down-regulated the hepatic expression of Gdh and Usf2 to similar levels and reduced Usf2 binding to glud promoter, while refeeding resulted in a slow but gradual restoration of both Gdh and Usf2 mRNA abundance. Herein, we demonstrate that Usf2 transactivates S. aurata glud by binding to an E-box located in the proximal region of glud promoter. In addition, our findings provide evidence for a new regulatory mechanism involving Usf2 as a key factor in the nutritional regulation of glud transcription in the fish liver.

Introduction

Glutamate dehydrogenase (Gdh) catalyses reversible oxidative deamination of glutamate to form α-ketoglutarate and ammonia while reducing NAD(P)+ to NAD(P)H. Encoded by the glud gene, Gdh plays a major role in ammonia detoxification in the liver, acid excretion by providing urinary ammonia in the kidneys, amplification of glucose-stimulated insulin secretion in pancreatic β-cells, cycling of the neurotransmitter glutamate between neurons and astrocytes and gluthatione synthesis, among others (Newsholme et al. 2003, Karaca et al. 2011, Göhring & Mulder 2012, Treberg et al. 2014, Bunik et al. 2016). Located in the mitocondrial matrix, Gdh activity is subjected to a complex regulation. Gdh is strongly inhibited by GTP and activated by ADP. GTP binding is antagonised by phosphate and ADP, but is synergistic with NADH bound to a second, non-catalytic site. Gdh is also activated by leucine and other monocarboxylic acids, while it is inhibited by palmitoyl-CoA and diethylstilbestrol (Li et al. 2014, Plaitakis et al. 2017). Reversible cystein-specific ADP-ribosylation inactivates Gdh (Haigis et al. 2006). Gdh activity comes, at least in part, from association to a multienzyme complex in the mitochondrion, as it was deduced by the fact that short-chain 3-hydroxyacyl-CoA dehydrogenase (SCHAD) inhibits Gdh via protein–protein interaction in the pancreas, where SCHAD is expressed at high levels (Li et al. 2010).

Glutamate metabolism in fish differs from that of mammals. Glutamate is primarily deaminated in the fish liver with ammonia production, while in mammals most glutamate is transaminated to aspartate (Peres & Oliva-Teles 2006). The fact that Gdh plays a major role in amino acid oxidation in the liver, led to consider Gdh a marker for protein utilisation and ammonia excretion in fish (Liu et al. 2012). Gdh is mainly expressed in the piscine liver, and high-protein diets usually stimulate growth and hepatic Gdh activity in fish (Bibiano Melo et al. 2006, Borges et al. 2013, Viegas et al. 2015, Coutinho et al. 2016). Dietary protein increases plasma free amino acids, which in turn enhances Gdh-dependent deamination and leads to higher rates of ammonia excretion. Dietary supplementation of glutamate downregulates Gdh mRNA levels and decreases reductive Gdh activity in the liver of Sparus aurata (Gómez-Requeni et al. 2003, Caballero-Solares et al. 2015) and reduces Gdh activity in Pagellus bogaraveo (Figueiredo-Silva et al. 2010), while it increases in Oncorhynchus mykiss (Moyano et al. 1991).

Little is known about transcriptional regulation of Gdh expression in vertebrates. In silico analysis allowed detection of potential binding sites for a number of transcription factors, such as Sp1, AP-1, and AP-2 in humans, and Sp1 and Zif268 in rats (Das et al. 1993, Michaelidis et al. 1993). The functionality of these sites remains unclear. A glucocorticoid-responsive region was located in the gene promoter of the C8S mouse astrocyte-derived cell line (Hardin-Pouzet et al. 1996). Deletion of the gene coding for the transcriptional coactivator p300 in the human colon carcinoma cell line HCT116 down-regulates Gdh expression (Bundy et al. 2006). Despite the important role exerted by Gdh in various tissues, to our knowledge there are no reports that have adressed isolation and molecular characterisation of glud gene promoter from fish.

With the aim of increasing current knowledge about the transcriptional regulation of glud, in the present study we characterised for the first time a piscine glud promoter and addressed the role of upstream stimulatory factor 2 (Usf2) on glud transcription in gilthead sea bream (Sparus aurata). In addition to report for the first time transactivation of glud gene promoter by Usf2, we explored changes in Gdh and Usf2 expression associated with starvation and refeeding in the liver of S. aurata.

Materials and methods

Experimental animals

S. aurata juveniles obtained from Piscimar (Burriana, Castellón, Spain) were maintained at 20°C in 260-L aquaria supplied with running seawater as described (Fernández et al. 2007). Nutritional regulation of Gdh and Usf2 expression was analysed in the liver of 18-day fed fish, 19-day starved fish and fish refed for 6 h, 24 h, 5 days and 14 days. The diet was supplied at 25 g/kg body weight (BW) once a day (10 h) and contained 46% protein, 9.3% carbohydrates, 22% lipids, 10.6% ash, 12.1% moisture and 21.1 kJ/g gross energy. To prevent stress, fish were anesthetised with MS-222 (1:12,500) before handling. Twenty-four hours after the last meal, fish were sacrificed by cervical section and tissues were dissected out, frozen in liquid N2 and kept at −80°C until use. The University of Barcelona’s Animal Welfare Committee approved the experimental procedures (proceeding #461/16) in compliance with local and EU legislation.

Characterization of the transcription start site

The 5′ end of S. aurata Gdh mRNA was determined using the First Choice RLM-RACE Kit (Thermo Fisher Scientific). RLM-RACE allows amplification of cDNA only from full-length, capped mRNA (Schaefer 1995). Briefly, 10 µg of total RNA from S. aurata liver were treated with calf intestine alkaline phosphatase and submitted to phenol–chloroform extraction and isopropanol precipitation. RNA was resuspended and treated with tobacco acid pyrophosphatase to remove the cap structure from full-length mRNAs, while leaving a 5′-monophosphate required for further ligation of the 5′ RACE adapter oligonucleotide provided with the kit. Following random-primed reverse transcription, a nested PCR amplified the 5′ end of S. aurata Gdh mRNA using gene-specific primers CG1307 and CG1308, designed from the S. aurata partial cDNA sequence with GenBank no. JX073708 (Table 1). The single 942-bp amplicon generated was ligated into pGEM-T easy (Promega). Identical nucleotide sequence was obtained by sequence analysis of three independent clones.

Table 1

Oligonucleotides used in the present study.

PrimerSequence (5′–3′)GenBank accession no.
CG1307GTCTTGTCCTGGAAGCCTGGTGTCAJX073708
CG1308GGCTGAGATACGACCGTGGATACCTCCCJX073708
CG1315GACAGGAGAAGGGGGGTAGAATGAACGACMF459045
CG1316AACAACAAGGACAATGGGGGTGACGACAGMF459045
CG1342CCCAGCTGTCAGTTGGACAGCACGGMF459046
CG1344CCCCCGGGACACGGTGAGGAGCTGCMF459046
CG1345CCCCGCTCTTCCGCGTGAGTCCCGMF459046
CG1543GGTATTTCGGGGAGCTGCTGAGMF459045
CG1544CGCATCAGGGACGAGGACAMF459045
CG1552CTCTCCGCGGCTCGTGCTGCCTTTTAAAGCAAACTGACACAGTTTTTCATTCCCCACTCGGCCAGAGGACMF459046
CG1557AGAGCTGAGGCAAAGCAACC*
CG1558GGGGAGGACGCATTCACTAA*
CG1561CACCCGGTCATGTGACCTACACMF459046
CG1562TGTAGGTCACATGACCGGGTGGMF459046
CG1563AAACTGACACAGCATGTCATTCCCCACTCGGCMF459046
CG1564CCGAGTGGGGAATGACATGCTGTGTCAGTTTGMF459046
CG1565AAACTGACACAGTTTTTCATTCCCCACTCGGCMF459046
CG1566CCGAGTGGGGAATGAAAAACTGTGTCAGTTTGMF459046
CG1701CCAGCACAATGACATTTCTATTGMF459046
CG1702GTTAAAAAACTTGTATGGTTGMF459046
CG1703CGCGCGCTGTCAGTTGGACAGCACMF459046
CG1705ACAGCAGCTCCTCACCGTGTCCMF459046
AS-EF1FwCCCGCCTCTGTTGCCTTCGAF184170
AS-EF1RvCAGCAGTGTGGTTCCGTTAGCAF184170
JDRT18STTACGCCCATGTTGTCCTGAGAM490061
JDRT18ASAGGATTCTGCATGATGGTCACCAM490061
QBACTINFCTGGCATCACACCTTCTACAACGAGX89920
QBACTINRGCGGGGGTGTTGAAGGTCTCX89920
JS1711GTACCTCGAGGCCAGTTCTACGTCATGATCAAGAGTCATGACGTAGAACTGGCCTCTTTTTGGAAA*
JS1712AGCTTTTCCAAAAAGAGGCCAGTTCTACGTCATGACTCTTGATCATGACGTAGAACTGGCCTCGAG*

The following primers contain restriction sites (underlined): CG1344 (SmaI), CG1345 (BsrBI) and CG1342 (PvuII). Bold and double-underlined letters indicate site-directed mutations in primers CG1552, CG1565 and CG1566.

*CG1557, CG1558, JS1711 and JS1712 were designed from recent transcriptome sequencing data (Silva-Marrero et al. 2017).

Isolation of the 5′-flanking region of S. aurata glud by chromosome walking

The 5′-flanking region of glud was isolated using the Universal GenomeWalker Kit (Clontech) and gene-specific primers CG1315 and CG1316 (Table 1), which were designed from the 5′ end of S. aurata Gdh cDNA. Blunt-end digestion of S. aurata genomic DNA with DraI, EcoRV, PvuII, and StuI generated four libraries that were ligated to the GenomeWalker adaptor as described (Metón et al. 2006). Two PCR rounds were performed on each library with gene-specific primers CG1316 and CG1315 for the primary and nested PCR, respectively. The longer amplicon (~2.1-kb) was isolated from the PvuII library and ligated into pGEM-T Easy (Promega) to generate pGEM-GDH2057. Two independent clones were fully sequenced on both strands following the ABI Prism BigDye Terminator Cycle Sequencing Ready Reaction kit instructions (Applied Biosystems).

Reporter and shRNA expression constructs

To generate pGDH1286, the S. aurata glud promoter fragment spanning positions −1286 to +70 relative to the transcriptional start was obtained by digestion of pGEM-GDH2057 with NotI, followed by fill-in and NheI digestion. The product was subcloned into the SmaI/NheI site of the promoterless luciferase reporter plasmid pGL3-Basic (Promega). To obtain pGDH982, the promoter fragment resulting from digestion of pGDH1286 with PstI followed by chew back, fill-in and HindIII digestion, was subcloned into pGL3-Basic, previously digested with MluI, filled-in and HindIII digested. pGDH+19, pGDH85 and pGDH128 were generated by PCR using pGDH1286 as template and primer pairs CG1344 (with a 5′-anchor sequence containing a SmaI site; Table 1)/RVprimer3 (Promega), CG1345 (with a 5′-anchor sequence containing a BsrBI site; Table 1)/RVprimer3, and CG1342 (with a 5′-anchor sequence containing a PvuII site; Table 1)/RVprimer3, respectively. PCR products were digested with SmaI, BsrBI and PvuII, respectively, isolated and subcloned into pGL3-Basic, previously digested with MluI, filled-in and HindIII digested. pGDH413 was produced by NdeI/MluI digestion, filling-in and self-ligation of pGDH1286. pGDH982∆−44/+70 was obtained by Cfr42I digestion of pGDH982, isolation of the longest product and self-ligation. pGDH982mutUsf2 was generated by PCR using pGDH1286 as template and primer pair CG1552 (harbouring a mutated E-box and a 5′-anchor sequence with a Cfr42I site; Table 1)/GLprimer2 (Promega), and subcloning of the resulting amplicon into pGDH982, previous Cfr42I digestion of amplicon and plasmid. To generate pCpG-sh1Usf2, the double-stranded product obtained by hybridisation of oligonucleotides JS1711 and JS1712 (Table 1; designed using siRNA Wizard software, InvivoGen, San Diego, CA, USA), was ligated into the HindIII/Acc65I site of pCpG-siRNA (InvivoGen). All constructs were verified by cycle sequencing.

Cell transfection and luciferase assay

Human hepatoma-derived HepG2 cells (ATCC HB 8065) were cultured at 37°C and 5% CO2 in DMEM supplemented with 2 mM glutamine, 110 mg/L sodium pyruvate, 10% foetal bovine serum, 100 IU/mL penicillin and 100 μg/mL streptomycin. HepG2 cells at 45–50% confluence were transiently transfected in six-well plates using the calcium phosphate coprecipitation method. Transfection mixture included 4 μg of reporter construct and 500 ng of CMV-β plasmid (lacZ) to correct for variations in transfection efficiency, and was prepared with or without 400 ng of the Usf2 expression vector. Up to 400 ng of pCpGsh1Usf2 were added to perform shRNA-mediated silencing assays. Empty plasmids were added to ensure equal DNA amounts. Cells were harvested 16 h later, washed in PBS and incubated for 15 min in 300 μL of Cell Culture Lysis Reagent (Promega). Cell debris was pelleted, and luciferase activity in the supernatant was assayed in a TD-20/20 Luminometer (Turner Designs, Sunnyvale, CA, USA) after addition of Luciferase Assay Reagent (Promega). β-Galactosidase activity of the clear lysate was assayed as described (Metón et al. 2006). The Usf2 expression plasmid was kindly provided by Dr B Viollet (Viollet et al. 1996).

Electrophoretic mobility shift assay

Double-stranded oligonucleotides GDH−22/+9, GDH−22/+9mutUSF (carrying a mutated E-box) and USF2-cons (with a consensus Usf2 binding box) were obtained by hybridisation of oligonucleotide pairs CG1563/CG1564, CG1565/CG1566 and CG1561/CG1562 (Table 1), respectively. Two hundred picomole of double-stranded oligonucleotides were 3′-end labelled with digoxigenin-11-ddUTP in a 20-µL reaction for 30 min at 37°C using terminal transferase (Hoffman-La Roche, Basel, Switzerland). EDTA (18 mM) was added to stop the reaction. Binding reactions contained 100 mM HEPES, pH 7.6, 5 mM EDTA, 50 mM (NH4)2SO4, 5 mM dithiothreitol, 1% Tween 20, 150 mM KCl, 0.05 µg/µL non-specific competitor poly (d(I–C)), nuclear extracts of HepG2 cells overexpressing Usf2, and labelled probe. DNA–protein complexes were electrophoresed at 4°C on a 5% polyacrylamide gel using 0.5× Tris-borate-EDTA buffer. DNA was transferred by contact blotting to Nytran membranes and cross-linked by UV irradiation for 3 min. Labelled probes were immunodetected with antidigoxigenin conjugated to alkaline phosphatase using CDP-Star (Hoffman-La Roche) as chemiluminescent substrate. Digital imaging of membranes was performed using ImageQuant LAS 4000 mini (GE Healthcare). For competition experiments, HepG2 extracts were preincubated for 30 min with 200- and 1000-fold molar excess of unlabelled double-stranded USF2-cons.

Nuclear extracts

Nuclear extracts were prepared from HepG2 cells as described (Andrews & Faller 1991) with minor modifications. Cells grown at ~80% confluency were washed, scraped into 1.5 mL of cold PBS, pelleted by centrifugation for 10 s at 1000 g and resuspended in 0.4 mL of buffer A (10 mM HEPES, pH 7.9, 1.5 mM MgCl2, 10 mM KCl, 0.5 mM dithiothreitol, 0.2 mM phenylmethylsulfonyl fluoride). Following 10 min of incubation, cells were vortexed for 10 s and centrifuged. The pellet was resuspended in 20 µL of ice-cold buffer C (20 mM HEPES, pH 7.9, 25% glycerol, 420 mM NaCl, 1.5 mM MgCl2, 0.2 mM EDTA, 0.5 mM dithiothreitol, 0.2 mM phenylmethylsulfonyl fluoride) and incubated 20 min for high-salt extraction. After pelleting cell debris, the supernatant was kept at −80°C until use.

Western blot

Proteins in cell extracts were subjected to 10% PAGE-SDS electrophoresis, transferred to a polyvinylidene fluoride membrane and immunoblotted with mouse anti-Usf2 (Santa Cruz Biotechnology) and rabbit anti-actin (Sigma-Aldrich) antibodies. Chemiluminescent detection proceeded using alkaline phosphatase-conjugated secondary antibodies and the Clarity Western ECL Substrate Kit (Bio-Rad).

Quantitative real-time RT-PCR analysis

Total RNA (1 μg) isolated from tissue samples was retrotranscribed 1 h at 37°C with M-MLV RT (Promega). Gdh and Usf2 mRNA levels were determined in a Step One Plus Real Time PCR System (Applied Biosystems) using 0.4 µM of each primer (CG1543/CG1544 and CG1557/CG1558 for Gdh and Usf2, respectively; Table 1), 10 μL of SYBR Green (Applied Biosystems), and 1.6 μL of 1:10 diluted cDNA in a 20 µL-reaction. The temperature cycle protocol for amplification was 95°C for 10 min, followed by 40 cycles at 95°C for 15 s and 62°C for 1 min. A dissociation curve was run after each experiment to ensure amplification of single products. Gdh and Usf2 mRNA levels were normalised with the CT geometric mean value of ribosomal subunit 18s, beta-actin and elongation factor 1-alpha (EEF1A1), which were amplified using primer pairs JDRT18S/JDRT18AS, QBACTINF/QBACTINR and AS-EF1Fw/AS-EF1Rv, respectively (Table 1). Variations in gene expression were calculated by the standard ΔΔCT method (Pfaffl 2001).

Chromatin immunoprecipitation

Chromatin from S. aurata liver was isolated, cross-linked and sonicated to an average size of 100–600 bp as described (Metón et al. 2006). Following preclearing with 15 μL of protein A/G-agarose beads, immunoprecipitation of chromatin (100 μg) proceeded overnight at 4°C with or without 2 μg of antibody (anti-Usf2 or anti-Srebp1; Santa Cruz Biotechnology). Immune complexes were incubated with protein A/G-agarose beads, washed, eluted and reverse cross-linked with 0.4 mg/mL proteinase K for 2 h at 37°C and overnight at 65°C. Purified DNA was subjected to PCR using primer pairs CG1701/CG1702 and CG1703/CG1705 (Table 1), which amplify glud sequences −1766 to −1535 and −133 to +41, respectively. Nutritional regulation of Usf2 binding to glud promoter was determined by qPCR using primer pair CG1703/CG1705 (Table 1), Usf2-immunoprecipitated chromatin or input chromatin, and the protocol described in the previous section. Twenty-five microgram of non-immunoprecipitated chromatin was reverse cross-linked and retained as a positive control and to normalise qPCR results (input).

Statistics

SPSS Version 22 (IBM) was used to analyse data with Student’s two-tailed unpaired t-test or one-way ANOVA when comparing more than two groups. For ANOVA, significant differences were determined with the Bonferroni post hoc test.

Results

Cloning of the 5′-flanking region of S. aurata glud

A 5′ RLM RACE was performed on total RNA isolated from S. aurata liver to determine the nucleotide sequence at the 5′ end and the transcription start site of S. aurata Gdh mRNA. Analysis of the single fragment obtained indicated that S. aurata Gdh mRNA initiates 140 nucleotides upstream from the translation start codon. Availability of the 5′ end of Gdh mRNA enabled to design gene-specific primers to isolate a 2057 bp fragment upstream from the translation start codon of S. aurata glud by chromosome walking. Sequence analysis of the 1286-bp 5′-flanking region with JASPAR (Sandelin et al. 2004) revealed the presence of a TATA box at positions −32 to −18 relative to the transcription start site. Potential transcription factor sites included binding boxes for Usf2, Cebp and Hnf1, among others (Fig. 1). The nucleotide sequence of S. aurata Gdh mRNA and glud promoter were submitted to the GenBank database under accession numbers MF459045 and MF459046, respectively.

Figure 1
Figure 1

Sequence analysis of the 5′-flanking region comprised between positions −1316 to +143 relative to the transcriptional start of S. aurata glud. Chromosome walking allowed isolation of the genomic sequence upstream from the transcription start site of S. aurata Gdh, which is shown in capital letters. An arrow indicates the transcription start site. The translation start codon is in boldface and underlined. Putative binding sites for relevant transcription factors are boxed.

Citation: Journal of Molecular Endocrinology 60, 3; 10.1530/JME-17-0142

Functionality of S. aurata glud promoter

To assess whether the 5′-flanking region of S. aurata glud encompasses a functional promoter, nucleotide positions −1286 to +70 relative to the transcriptional start were subcloned into the promoterless pGL3-Basic plasmid upstream from the luciferase reporter gene. Consistent with the presence of a functional promoter, transient transfection of HepG2 cells with the resulting construct (pGDH1286; −1286 to +70) resulted in a 45-fold increase of luciferase activity relative to pGL3-Basic (Fig. 2). To identify functional regions involved in modulation of basal Gdh expression in S. aurata, sequential 5′-deletion of the isolated promoter was performed. HepG2 cells were transfected with pGL3-Basic constructs harbouring deletion fragments of glud promoter (5′ ends ranging from −1286 to +19 and 3′ ends at −70) fused to the luciferase reporter gene. The longer 5′ constructs (pGDH1286 and pGDH982; −982 to +70) yielded a 45-fold increase in luciferase activity relative to the empty vector. The reporter constructs pGDH413 (−413 to +70), pGDH128 (−128 to +70) and pGDH85 (−85 to +70) exhibited a 25- to 30-fold increase of promoter activity compared to pGL3-Basic. A significant drop of activity was observed using the smallest construct (pGDH+19; +19 to +70) or pGDH982∆−44/+70, which is a deleted construct that encompasses promoter sequences located at positions −982 to +70, but lacks the region comprised between positions −44 and +70 (Fig. 2). Therefore, the core promoter of S. aurata glud localises within 85 bp upstream from the transcriptional start, suggesting the presence of cis-acting elements in this region.

Figure 2
Figure 2

Functional analysis of the 5′-flanking region of S. aurata glud in HepG2 cells. The top left part represents the genomic organization of the 5′-flanking region of S. aurata glud. Relative position of relevant restriction sites and exon 1 are indicated. Nucleotide numbering starts with +1, which corresponds to the transcriptional start. Reporter constructs having varying 5′ ends and identical 3′ ends (+70), except for pGDH982∆−44/+70, were transfected in HepG2 cells along with pCMVβ to normalise for transfection efficiency. Luciferase activity is expressed as fold increase over promoterless reporter plasmid pGL3-Basic. Results shown are the mean ± s.d. from three independent experiments performed in duplicate. Different letters indicate significant differences among conditions (P < 0.05).

Citation: Journal of Molecular Endocrinology 60, 3; 10.1530/JME-17-0142

Transactivation of S. aurata glud promoter by Usf2

Analysis with JASPAR indicated the presence of an E-box that could function as a putative Usf2 binding site in the proximal promoter region of S. aurata glud (Fig. 1). Usf proteins regulate the transcription of essential gene networks. The fact that USF2 null-mutant mice are small and exhibit decreased fertility and reduced lifespan, while USF1 null mice present a rather normal phenotype, highlights Usf2 as the more important Usf variant (Sirito et al. 1998, Horbach et al. 2014). To study the role of Usf2 in Gdh expression we performed transfection experiments on HepG2 cells in the presence and absence of an expression plasmid encoding Usf2. Cotransfection with the Usf2 expression plasmid together with pGDH85 or longer 5′ constructs increased ~4-fold glud promoter activity compared to the basal activity of the corresponding promoter constructs. No Usf2-dependent enhancement of glud transcription was observed when using the shortest construct (pGDH+19). Altogether, these results suggest that a functional Usf2 binding site may be located within 85 pb upstream from the transcriptional start. Consistently, cotransfection of HepG2 cells with the Usf2 expression plasmid and pGDH982∆−44/+70, which lacks the region between positions −44 to +70, did not transactivate glud (Fig. 3).

Figure 3
Figure 3

Effect of Usf2 on the promoter activity of S. aurata glud gene in HepG2 cells. HepG2 cells were transfected with pGL3-Basic and promoter constructs pGDH+19, pGDH85, pGDH128, pGDH413, pGDH982 or pGDH982∆44/+70 along with pCMVβ and with or without an expression plasmid encoding Usf2. The promoter activity of the constructs alone was set at 1. Results are presented as mean ± s.d. values of three independent duplicate experiments. Statistical significance related to promoter activity of reporter constructs in absence of the Usf2 expression plasmid is indicated as follows: ***P < 0.001.

Citation: Journal of Molecular Endocrinology 60, 3; 10.1530/JME-17-0142

Usf2 binds to glud promoter

The shorter reporter construct that exhibited Usf2-mediated transactivation (pGDH85) contains an E-box at positions −10 to −5 relative to the transcriptional start. Bearing in mind that Usf2 transactivates numerous genes by binding to E-boxes (Corre & Galibert 2005), electrophoretic mobility shift assays (EMSA) were performed with nuclear extracts obtained from HepG2 cells overexpressing Usf2. Probes GDH−22/+9 (with the putative E-box at positions −10 to −5) and USF2-cons (containing a consensus Usf2 binding site) generated one major shifted band with the same mobility. The shifted DNA-protein complex disappeared by competition with 200- to 1000-fold molar excess of unlabelled USF2-cons. These results confirmed binding of Usf2 to a response element at positions −22 to +9 of S. aurata glud. Bandshift experiments performed using nuclear extracts of HepG2 cells overexpressing Usf2 and a labelled probe harbouring positions −22 to +9 of glud but with a mutated E-box element (GDH−22/+9mutUSF) completely prevented the formation of DNA–protein complexes (Fig. 4A).

Figure 4
Figure 4

(A) Analysis of USF binding to glud promoter by electrophoretic mobility shift assay. To perform a competition analysis, nuclear extracts of HepG2 cells overexpressing Usf2 were incubated with labelled oligonucleotides USF2-cons (lanes 1–4), GDH−22/+9 (lanes 5–8) or GDH−22/+9mutUSF (lanes 9–10). Lanes 1, 5 and 9 contained no extract. Lanes 2 and 6 show binding of nuclear extracts to labelled probes without competitor. Lanes 3 and 7 show competition with 200-fold molar excess of unlabelled double-stranded competitor (USF2-cons). Lanes 4, 8 and 10 show competition with 1000-fold molar excess of unlabelled double-stranded competitor (USF2-cons). DNA–protein complexes are indicated by an arrow. NE, nuclear extracts. (B) In vivo association of Usf2 with S. aurata glud promoter. A ChIP assay was performed on S. aurata liver. The upper part of the figure shows a schematic drawing of S. aurata glud promoter, location of the PCR primers (arrows) and sequence of E-box at position −10 to −5 relative to the transcriptional start (underlined). After cross-linking with 1% formaldehyde, chromatin was sheared by sonication, and immunoprecipitated in the presence of anti-Usf2 and anti-Srebp1 antibodies, or incubated without antibodies. Immune complexes were collected with protein A/G-agarose beads, and following intensive washing, bound DNA-complexes were eluted and reverse cross-linked. Analysis of purified DNA was performed by PCR with primer pairs to amplify glud promoter positions −1766/−1535 or −133/+41 relative to the transcription start site. The PCR products were electrophoresed on an agarose gel and visualised by means of RedSafe nucleic acids staining.

Citation: Journal of Molecular Endocrinology 60, 3; 10.1530/JME-17-0142

Chromatin immunoprecipitation (ChIP) was performed to study association of Usf2 with S. aurata glud promoter in vivo. Following ChIP with anti-Usf2, PCR analysis on purified DNA using primer pairs to amplify glud promoter positions −133 to +41 confirmed that the E-box at positions −10 to −5 contains a functional Usf2 binding site in vivo. No binding was observed with a different antibody (anti-Srebp1), no antibody or using oligonucleotides to amplify an upstream region (−1766 to −1535) (Fig. 4B).

Mutating the E-box abolishes transactivation by Usf2

To analyse the effect of mutating the E-box located at positions −10 to −5 on Usf2-dependent transactivation of glud, we generated a reporter construct containing the same mutations introduced in the double-stranded oligonucleotide GDH−22/+9mutUSF used for bandshift assays. Usf2 failed to enhance transcriptional activity of the resulting construct (pGDH982mutUSF2; −982 to +70 with a mutated E-box). Indeed, Western blot analysis revealed immunodetection of endogenous Usf2 in HepG2 cells, while confirmed overexpression of Usf2 after cotransfection with the Usf2 expression plasmid (Fig. 5A). Furthermore, cotransfection of HepG2 cells with pGDH982 and a construct expressing an shRNA to knock-down Usf2 (pCpGsh1Usf2) abolished Usf2-dependent transactivation of glud promoter (Fig. 5B). Therefore, the E-box located at positions −10 to −5 relative to the major transcriptional start of the S. aurata glud is responsible for transactivation by Usf2.

Figure 5
Figure 5

(A) Effect of Usf2 on the promoter activity of glud containing a mutated E-box. The upper part of the figure shows a representative Western blot analysis of immunodetectable Usf2 and actin proteins in extracts of HepG2 cells transfected with the promoter constructs pGDH+19, pGDH85, pGDH982 or pGDH982mutUsf2, along with pCMVβ and with or without an expression plasmid encoding Usf2. The lower part of the figure shows induction of promoter activity in HepG2 cells transfected with the promoter constructs pGDH+19, pGDH85, pGDH982 or pGDH982mutUsf2, along with pCMVβ and with or without an expression plasmid encoding Usf2. The luciferase activity of the reporter constructs alone was set at 1. Results are presented as mean ± s.d. values of three independent duplicate experiments. Statistical significance related to promoter activity of reporter constructs in absence of the Usf2 expression plasmid is indicated as follows: **P < 0.01; ***P < 0.001. (B) Effect of Usf2 silencing on Usf2-dependent transactivation of glud promoter. HepG2 cells were transfected with the promoter construct pGDH982 along with pCMVβ, an expression plasmid encoding Usf2 and increasing amounts of pCpGsh1Usf2. The luciferase activity of pGDH982 in the absence of pCpGsh1Usf2 was set at 1. Results are presented as mean ± s.d. values of triplicate experiments. Different letters indicate significant differences among conditions (P < 0.05).

Citation: Journal of Molecular Endocrinology 60, 3; 10.1530/JME-17-0142

Tissue distribution of Gdh and Usf2 expression in S. aurata

To study tissue specificity of Gdh and Usf2 expression in S. aurata, reverse transcriptase-coupled quantitative real-time PCR (RT-qPCR) was performed in tissue samples of fed S. aurata. The highest mRNA levels of Gdh were found in the liver, followed by the intestine, heart and kidney. Gdh expression was barely detectable in other tissues. Usf2 was ubiquitously expressed, albeit higher Usf2 mRNA abundance was exhibited by the brain and spleen, followed by the heart, gill, kidney and liver (Fig. 6).

Figure 6
Figure 6

Tissue distribution of Usf2 and Gdh expression in S. aurata. RT-qPCR assays of Usf2 and Gdh mRNA levels were performed on total RNA isolated from the spleen, gill, brain, heart, fat, liver, intestine, skeletal muscle and kidney of 18-day fed fish. Expression levels for each gene were normalised using 18S, beta-actin and EEF1A1 as housekeeping genes. Results are presented as mean ± s.d. (n = 4).

Citation: Journal of Molecular Endocrinology 60, 3; 10.1530/JME-17-0142

Effect of starvation and refeeding on the hepatic expression of Gdh and Usf2

Having concluded that Usf2 can bind and transactivate glud promoter in the liver, we addressed the role that Usf2 may exert in the nutritional regulation of hepatic glud transcription in S. aurata. Gdh and Usf2 mRNA levels were determined by RT-qPCR in liver samples of 18-day fed fish, 19-day starved fish and fish refed up to 14 days. Nutritional changes affected similarly Gdh and Usf2 expression. Starvation significantly decreased 1.7-fold mRNA abundance of both Gdh and Usf2. Remarkably, a trend to present lower expression levels of Gdh and Usf2 than starved fish was observed 6 h after refeeding. Thereafter, Gdh and Usf2 gradually recovered their mRNA levels until reaching total restoration after 14 days of refeeding (Fig. 7A). ChIP assays showed that starvation decreased Usf2 binding to glud promoter, while a trend to recover the values observed in fed fish was observed after 14 days of refeeding (Fig. 7B).

Figure 7
Figure 7

Effect of starvation and refeeding on Usf2 and Gdh mRNA levels, and Usf2 binding to glud promoter in the liver of S. aurata. (A) RT-qPCR assays of Usf2 and Gdh mRNA levels were performed on total RNA isolated from the liver of 18-day fed, 19-day starved, and refed fish for 6 h, 24 h, 5 days and 14 days. Expression levels for each gene were normalised using 18S, beta-actin and EEF1A1 as housekeeping genes. Results are presented as mean ± s.d. (n = 6). (B) ChIP analysis of Usf2 association with glud promoter in the liver of 18-day fed, 19-day starved, and refed fish for 6 h and 14 days. Results are presented as mean ± s.d. (n = 3). Different letters indicate significant differences among conditions (P < 0.05).

Citation: Journal of Molecular Endocrinology 60, 3; 10.1530/JME-17-0142

Discussion

In the liver, Gdh is essential for ammonia detoxification, nitrogen metabolism and urea synthesis (Karaca et al. 2011, Treberg et al. 2014). However, knowledge of the transcription factors involved in the regulation of glud gene expression is scarce. To study the transcriptional regulation of glud, we addressed cloning and characterisation of S. aurata glud promoter by chromosome walking. Functionality of S. aurata glud promoter was confirmed by transient transfection of HepG2 cells with fusion constructs of sequential 5′-deletions of the isolated genomic fragment to the luciferase gene. We found that the core functional promoter of S. aurata glud is comprised within 85 bp upstream from the transcription start site. The presence of a putative Usf2 binding box in the proximal region of glud gene promoter prompted us to study involvement of Usf2 in the transcriptional regulation of glud.

Usf proteins belong to the basic helix-loop-helix-leucine zipper (bHLHzip) transcription factor family and are encoded by two different genes: usf1 and usf2. Usf proteins regulate the transcription of a wide number of genes involved in stress and immune responses, cell cycle and proliferation, and carbohydrate and lipid metabolism, among other functions, by binding as homodimers and heterodimers to the E-box binding motif CANNTG (being NN nucleotides in most cases either GC or CG), non-canonical E-boxes and pyridine-rich initiator sites (Viollet et al. 1996, Corre & Galibert 2005, Pawlus et al. 2012). Transient transfection studies in HepG2 cells together with EMSA and ChIP assays allowed us to demonstrate that Usf2 transactivates the promoter activity of S. aurata glud through binding to the E-box located at positions −10 to −5 upstream from the transcriptional start. Transactivation of glud by Usf2 was confirmed by introducing mutations in the E-box that abolished binding of Usf2 and prevented Usf2-dependent enhancement of glud transcription. Indeed, transfection of HepG2 cells with an shRNA expression plasmid to knock-down Usf2 abolished Usf2-dependent transactivation of glud promoter.

Optimal growth of teleostean fish requires higher levels of dietary protein than other vertebrates. Fish metabolism, and more remarkably that of carnivorous fish, enables efficient use of amino acids for growth and to obtain energy (Li et al. 2009, Kaushik & Seiliez 2010, Liu et al. 2012). The fish liver is the main site for amino acid catabolism, where Gdh exerts a major role in amino acid transdeamination by catalysing oxidative deamination of glutamate and giving rise to the end product of protein catabolism, ammonia (Lushchak et al. 2008). As for other fish species (Liu et al. 2012) and similar to mammals (Plaitakis et al. 2017), we found that Gdh is mainly expressed in the liver of S. aurata, while high mRNA levels were also observed in the kidney, heart and intestine. In contrast to Gdh, Usf2 displayed ubiquitous expression in S. aurata tissues, albeit higher mRNA levels were found in the brain, spleen, heart, gill, kidney and liver. These results are consistent with the pattern of tissue distribution of Usf2 in other vertebrates (Sirito et al. 1994, Fujimi & Aruga 2008). Our findings suggest that in addition to Usf2-dependent transactivation of glud promoter, other yet unknown transcription factors may contribute to upregulation of Gdh mRNA levels in the piscine liver. Posttranslational modifications as phosphorylation or interaction with other transcription factors and cofactors may also explain tissue-specific differences in Usf2 action (Spohrer et al. 2016).

Since Gdh expression can be considered a significant marker for protein utilisation and ammonia excretion in fish (Liu et al. 2012), we also adressed the effect of nutritional status on hepatic mRNA levels of Gdh, and the role that Usf2 may have on glud transcription under starvation and during the starved-to-fed transition in the liver of S. aurata. Long-term starvation similarly affected the hepatic expression of Usf2 and Gdh, which significantly decreased to about 60% of the values observed in fed fish, and reduced Usf2 binding to glud promoter. Downregulation of Gdh expression in starved S. aurata may be related with a mechanism preventing insulin secretion in β-cells. In favour of this hypothesis, overexpression of Gdh in mice increases insulin secretion (Carobbio et al. 2004), whereas Gdh inhibition in pancreatic β-cells decrease impairs insulin secretion (Carobbio et al. 2009). Furthermore, activating mutations in Gdh causes hyperinsulinemia and hyperammonemia in humans (Li et al. 2014, Barrosse-Antle et al. 2017). However, the effect of nutritional status on Gdh expression seems species-specific in fish. In contrast to S. aurata, starvation did not affect Gdh activity in Salmo gairdneri (Tranulis et al. 1991), while it increased Gdh activity in the liver of Oncorhynchus mykiss, Protopterus dolloi and Dentex dentex, and Gdh mRNA levels in Danio rerio (Sánchez-Muros et al. 1998, Frick et al. 2008, Pérez-Jiménez et al. 2012, Tian et al. 2015). We cannot discard that in addition to species-specificity, differences in the effect of starvation on the expression of Gdh among experiments may result also from diet composition, ration size and feeding regime. In this regard, it is well known that dietary protein greatly influences the hepatic activity of Gdh in fish (Liu et al. 2012, Borges et al. 2013, Caballero-Solares et al. 2015, Viegas et al. 2015, Coutinho et al. 2016). Indeed, it was reported that starvation decreased or unaffected Gdh activity in the liver of starved Dicentrarchus labrax depending on dietary protein levels (Pérez-Jiménez et al. 2007). As for Gdh, starvation decreased hepatic mRNA levels of Usf2 in the liver of S. aurata. In this regard, it was previously reported that high glucose levels upregulate Usf2 expression in human-derived HK-2 cells and primary rat mesangial cells (Shi et al. 2008, Visavadiya et al. 2011, Wang 2015). Therefore, low levels of glycemya associated to long-term starvation may be critical to downregulate Usf2 expression, which in turn may lead to decreased Gdh mRNA levels in the liver of S. aurata.

As for starvation, Usf2 and Gdh mRNA levels followed the same expression pattern after short- and long-term refeeding in the liver of S. aurata: a slow but gradual recovery of the values observed in fed fish. Five days of refeeding did not promote significantly higher expression levels than in starved fish for both Usf2 and Gdh. However, 14 days of refeeding allowed restoration of pre-starvation values. Furthermore, Usf2 and Gdh mRNA values in the liver of 14-day refed fish showed a trend to present slightly higher levels than fed fish. Similarly, long-term refeeding after starvation increased Gdh activity in the liver of Dicentrarchus labrax and Dentex dentex to values higher than in control fed fish (Pérez-Jiménez et al. 2007, 2012). Conceivably, refeeding after long-term starvation may require a long period of adaptation involving enhanced nutrient catabolism to restore metabolic parameters, as pointed out for other key enzymes involved in the intermediary metabolism (Soengas et al. 2006, Polakof et al. 2007). Therefore, an increased hepatic expression of Gdh may be essential for glutamate deamination and transdeamination of dietary amino acids in long-term refed fish to provide α-ketoglutarate for the Krebs cycle and supply ATP for energetic demands and biosynthesis.

Although Gdh and Usf2 may be subjected to similar regulatory cascades, the fact that Usf2 mRNA levels showed a complete correlation with Gdh expression during starvation and refeeding suggests that Usf2 may have a major role in the nutritional regulation of glud transcription in the liver of S. aurata. Involvement of Usf2 in the expression of genes encoding key enzymes in amino acid metabolism, such as Gdh, is consistent with previous observations showing transcriptional control of genes related to lipid, carbohydrate and energy metabolism by USF family members in mammals (Shih & Towle 1994, Lefrançois-Martinez et al. 1995, Iynedjian 1998, Martin et al. 2003, Corre & Galibert 2005, Pawlus et al. 2012).

In conclusion, in the present study we report for the first time characterisation of a piscine glud gene promoter and provide evidence for a novel regulatory mechanism that links Usf2 to the nutritional regulation of glud transcription in the fish liver.

Declaration of interest

The authors declare that there is no conflict of interest that could be perceived as prejudicing the impartiality of the research reported.

Funding

This work was supported by the AGL2016-78124-R grant (MEC, Spain; cofunded by the European Regional Development Fund, EC).

Author contribution statement

I V B and I M conceived and designed the study. C G, J I S-M and M C S performed the experiments. C G, I V B and I M analysed the data and edited the manuscript.

Acknowledgements

The authors thank Piscimar (Burriana, Castellón, Spain) for providing S. aurata juveniles, the Aquarium of Barcelona (Barcelona, Spain) for supplying filtered seawater, and Dr B Viollet (Institut Cochin, France) for providing the Usf2 expression vector.

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    Sequence analysis of the 5′-flanking region comprised between positions −1316 to +143 relative to the transcriptional start of S. aurata glud. Chromosome walking allowed isolation of the genomic sequence upstream from the transcription start site of S. aurata Gdh, which is shown in capital letters. An arrow indicates the transcription start site. The translation start codon is in boldface and underlined. Putative binding sites for relevant transcription factors are boxed.

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    Functional analysis of the 5′-flanking region of S. aurata glud in HepG2 cells. The top left part represents the genomic organization of the 5′-flanking region of S. aurata glud. Relative position of relevant restriction sites and exon 1 are indicated. Nucleotide numbering starts with +1, which corresponds to the transcriptional start. Reporter constructs having varying 5′ ends and identical 3′ ends (+70), except for pGDH982∆−44/+70, were transfected in HepG2 cells along with pCMVβ to normalise for transfection efficiency. Luciferase activity is expressed as fold increase over promoterless reporter plasmid pGL3-Basic. Results shown are the mean ± s.d. from three independent experiments performed in duplicate. Different letters indicate significant differences among conditions (P < 0.05).

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    Effect of Usf2 on the promoter activity of S. aurata glud gene in HepG2 cells. HepG2 cells were transfected with pGL3-Basic and promoter constructs pGDH+19, pGDH85, pGDH128, pGDH413, pGDH982 or pGDH982∆44/+70 along with pCMVβ and with or without an expression plasmid encoding Usf2. The promoter activity of the constructs alone was set at 1. Results are presented as mean ± s.d. values of three independent duplicate experiments. Statistical significance related to promoter activity of reporter constructs in absence of the Usf2 expression plasmid is indicated as follows: ***P < 0.001.

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    (A) Analysis of USF binding to glud promoter by electrophoretic mobility shift assay. To perform a competition analysis, nuclear extracts of HepG2 cells overexpressing Usf2 were incubated with labelled oligonucleotides USF2-cons (lanes 1–4), GDH−22/+9 (lanes 5–8) or GDH−22/+9mutUSF (lanes 9–10). Lanes 1, 5 and 9 contained no extract. Lanes 2 and 6 show binding of nuclear extracts to labelled probes without competitor. Lanes 3 and 7 show competition with 200-fold molar excess of unlabelled double-stranded competitor (USF2-cons). Lanes 4, 8 and 10 show competition with 1000-fold molar excess of unlabelled double-stranded competitor (USF2-cons). DNA–protein complexes are indicated by an arrow. NE, nuclear extracts. (B) In vivo association of Usf2 with S. aurata glud promoter. A ChIP assay was performed on S. aurata liver. The upper part of the figure shows a schematic drawing of S. aurata glud promoter, location of the PCR primers (arrows) and sequence of E-box at position −10 to −5 relative to the transcriptional start (underlined). After cross-linking with 1% formaldehyde, chromatin was sheared by sonication, and immunoprecipitated in the presence of anti-Usf2 and anti-Srebp1 antibodies, or incubated without antibodies. Immune complexes were collected with protein A/G-agarose beads, and following intensive washing, bound DNA-complexes were eluted and reverse cross-linked. Analysis of purified DNA was performed by PCR with primer pairs to amplify glud promoter positions −1766/−1535 or −133/+41 relative to the transcription start site. The PCR products were electrophoresed on an agarose gel and visualised by means of RedSafe nucleic acids staining.

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    (A) Effect of Usf2 on the promoter activity of glud containing a mutated E-box. The upper part of the figure shows a representative Western blot analysis of immunodetectable Usf2 and actin proteins in extracts of HepG2 cells transfected with the promoter constructs pGDH+19, pGDH85, pGDH982 or pGDH982mutUsf2, along with pCMVβ and with or without an expression plasmid encoding Usf2. The lower part of the figure shows induction of promoter activity in HepG2 cells transfected with the promoter constructs pGDH+19, pGDH85, pGDH982 or pGDH982mutUsf2, along with pCMVβ and with or without an expression plasmid encoding Usf2. The luciferase activity of the reporter constructs alone was set at 1. Results are presented as mean ± s.d. values of three independent duplicate experiments. Statistical significance related to promoter activity of reporter constructs in absence of the Usf2 expression plasmid is indicated as follows: **P < 0.01; ***P < 0.001. (B) Effect of Usf2 silencing on Usf2-dependent transactivation of glud promoter. HepG2 cells were transfected with the promoter construct pGDH982 along with pCMVβ, an expression plasmid encoding Usf2 and increasing amounts of pCpGsh1Usf2. The luciferase activity of pGDH982 in the absence of pCpGsh1Usf2 was set at 1. Results are presented as mean ± s.d. values of triplicate experiments. Different letters indicate significant differences among conditions (P < 0.05).

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    Tissue distribution of Usf2 and Gdh expression in S. aurata. RT-qPCR assays of Usf2 and Gdh mRNA levels were performed on total RNA isolated from the spleen, gill, brain, heart, fat, liver, intestine, skeletal muscle and kidney of 18-day fed fish. Expression levels for each gene were normalised using 18S, beta-actin and EEF1A1 as housekeeping genes. Results are presented as mean ± s.d. (n = 4).

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    Effect of starvation and refeeding on Usf2 and Gdh mRNA levels, and Usf2 binding to glud promoter in the liver of S. aurata. (A) RT-qPCR assays of Usf2 and Gdh mRNA levels were performed on total RNA isolated from the liver of 18-day fed, 19-day starved, and refed fish for 6 h, 24 h, 5 days and 14 days. Expression levels for each gene were normalised using 18S, beta-actin and EEF1A1 as housekeeping genes. Results are presented as mean ± s.d. (n = 6). (B) ChIP analysis of Usf2 association with glud promoter in the liver of 18-day fed, 19-day starved, and refed fish for 6 h and 14 days. Results are presented as mean ± s.d. (n = 3). Different letters indicate significant differences among conditions (P < 0.05).

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