atRA Regulation of NEDD9, a gene involved in neurite outgrowth and cell adhesion

Archives of Biochemistry and Biophysics 477 (2008) 163–174

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atRA Regulation of NEDD9, a gene involved in neurite outgrowth and cell adhesion D.C. Knutson a, M. Clagett-Dame a,b,c,* a b c

Interdepartmental Graduate Program in Nutritional Sciences, University of Wisconsin-Madison, 433 Babcock Drive, Madison, WI 53706-1544, USA Department of Biochemistry, University of Wisconsin-Madison, 433 Babcock Drive, Madison, WI 53706-1544, USA Pharmaceutical Sciences Division, University of Wisconsin-Madison, 433 Babcock Drive, Madison, WI 53706-1544, USA

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Article history: Received 28 March 2008 and in revised form 10 June 2008 Available online 15 June 2008 Keywords: NEDD9 RAINB2 HEF1 Cas-L Gene regulation Hindbrain Retinoic acid response element Vitamin A

a b s t r a c t We previously identified NEDD9 (RAINB2/HEF1/Cas-L) as a new downstream target of all-trans retinoic acid (atRA) and its receptors in the human neuroblastoma cell line, SH-SY5Y [R.A. Merrill, A.W.-M. See, M.L. Wertheim, M. Clagett-Dame, Dev. Dyn. 231 (2004) 564–575; R.A. Merrill, J.M. Ahrens, M.E. Kaiser, K.S. Federhart, V.Y. Poon, M. Clagett-Dame, Biol. Chem. 385 (2004) 605–614]. We now provide functional evidence that NEDD9 is directly regulated by atRA through a complex retinoic acid response element (RARE) located in the NEDD9 proximal promoter and consisting of four conserved half-sites separated by 1, 5, and 1 intervening base pairs. We show that a region of the human NEDD9 promoter from 1670 to +15 is sufficient to confer atRA-responsiveness and that a complex RARE located from 475 to 445 is necessary for this effect. While mutation of any one half-site does not eliminate complex formation in electrophoretic mobility shift assays (EMSA); these same mutations, when tested in transient transfection assays, markedly decrease atRA-responsiveness. Finally, chromatin immunoprecipitation (ChIP) assays demonstrate that RAR and RXR are bound to the RARE in cells. Ó 2008 Elsevier Inc. All rights reserved.

The vitamin A metabolite, all-trans retinoic acid (atRA),1 plays an essential role in development by regulating many cellular processes such as proliferation, differentiation, and migration [3]. atRA acts by binding to members of the nuclear receptor superfamily, the retinoic acid receptors (RAR). These receptors are ligand-activated transcription factors that modulate the expression of genes under the control of the ligand–receptor complex. There are three subtypes of RAR (a, b, and c) encoded by separate genes, and multiple isoforms for each that arise from differential promoter usage and mRNA splicing [4]. The RAR forms a heterodimer with the retinoid X * Corresponding author. Address: Department of Biochemistry, University of Wisconsin-Madison, 433 Babcock Drive, Madison, WI 53706-1544, USA. Fax: +1 608 262 7122. E-mail address: [email protected] (M. Clagett-Dame). 1 Abbreviations used: NEDD9, neural precursor cell expressed, developmentally down-regulated 9; RAINB2, retinoic acid inducible in neuroblastoma cells 2; HEF1, human enhancer of filamentation 1; Cas-L, Crk-associated substrate lymphocyte type; atRA, all-trans retinoic acid; RARE, retinoic acid response element; EMSA, electrophoretic mobility shift assay; ChIP, chromatin immunoprecipitation; RAR, retinoic acid receptor; RXR, retinoid X receptor; DMEM, Dulbecco’s modified Eagle media; Cas, CRK-associated substrate; PBS, phosphate buffered saline; DR, direct repeat; RXRE, retinoid X response element; Hoxa1, homeo box A1; COUP-TF, chicken ovalbumin upstream promoter transcription factor; HNF4, hepatocyte nuclear factor 4; SV40, simian virus 40; Act D, actinomycin D; CHX, cycloheximide; GAPDH, glyceraldehyde3-phosphate dehydrogenase; pCMVb, porcine cytomegalovirus beta; CRABPII, cellular retinoic acid binding protein II; C3, chromosome capture conformation assay; mTGRRE1, mouse tissue transglutaminase response element; HRE, hormone response element; BAC, bacterial artificial chromosome. 0003-9861/$ - see front matter Ó 2008 Elsevier Inc. All rights reserved. doi:10.1016/j.abb.2008.06.005

receptor (RXR), and it is this complex that shows high-affinity binding to specific DNA sequences called retinoic acid response elements (RARE). The consensus RARE sequence to which RARs bind is composed of two direct repeats of PuG(G/T)TCA separated by a variable number of bases, most often 5, 2, or 1 [5]. However, natural RAREs do not always adhere to the canonical half-site consensus sequence and spacing rules, and more complex elements have also been described. Regulation of gene transcription via the binding of the RXR/RAR complex to these RARE enhancer sequences is the major mechanism whereby atRA directly regulates target gene expression. In the current model, the receptor heterodimer is bound to the RARE both in the presence and absence of ligand and atRA binds only to the RAR partner of the RAR/RXR complex [6,7]. In cells, nuclear receptor activity is modulated via chromatin remodeling by coactivators and co-repressors [8]. In the absence of atRA, the RXR/RAR complex recruits co-repressor proteins that keep the chromatin condensed and gene expression silenced. atRA binding by RAR signals a conformational change leading to the dissociation of corepressor proteins from the RXR/RAR heterodimer and recruitment of coactivator proteins that open up the chromatin and facilitate transcriptional activity [9]. atRA regulates the expression of genes in a tissue and cell-typespecific fashion, and acts as a regulatory molecule during the development of many developing systems including the nervous system [10]. atRA signaling in the developing nervous system is required for normal hindbrain patterning and development of the

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post-otic cranial nerves. The development of this brain region is highly sensitive to both vitamin A deficiency and conversely, to excess atRA [11]. Hindbrain patterning involves the development of transient structures called rhombomeres (‘‘territories” of cells with a particular developmental fate) whose identity can be transformed through the expression of different mRNAs and proteins. Thus, atRA is thought to regulate the expression patterns of target genes in the hindbrain thereby influencing rhombomeric identity as well as downstream developmental fate [12]. Many of the atRA target genes that play a role in hindbrain development remain to be identified; their identification is critical to understanding the mechanisms underlying the biological effects of atRA in the developing nervous system. Recently, our group identified NEDD9 as a gene that is rapidly induced in SH-SY5Y cells by treatment of cells with atRA [1,2]. Both neurite outgrowth and cell spreading occur in SH-SY5Y cells exposed to atRA. Work from our group shows that early in embryogenesis NEDD9 mRNA is expressed in the node region, and prior to the onset of overt segmentation it is found in the developing hindbrain where its expression is altered both by retinoid deficiency and excess (by 6 h after exposure to atRA) [1]. NEDD9 is a member of the CRK-associated substrate (Cas) family of multidomain docking proteins, and contains multiple protein–protein interaction domains with which it may assemble proteins and regulate signaling cascades. Cell culture studies have shown that this protein plays important roles in regulating cell shape, cell migration, apoptosis and the cell cycle [13–17]. We are interested in determining how genes that lie downstream of atRA and its receptor are regulated, with specific interest in the genes that play a role in nervous system development and are directly regulated by atRA. In the present report we extensively characterize a RARE 869 bp 50 of the published HEF1 (NEDD9) sequence (GenBank Accession No. L43821), and show that it is more complex in nature than many RAREs in other known atRA target genes. Materials and methods DNA constructs for transient transfections Primers, listed in Table 1, containing the appropriate cut sites (SacI and XhoI) for cloning were used to amplify two lengths of the NEDD9 promoter; from (1670 to +15) and from (2756 to +15) from a human bacterial artificial chromosome (BAC; RP11263D22, CHORI; Oakland, Ca). Products were gel purified, subcloned into the pGEM-T easy vector system and sequenced. The inserts were excised via (SacI and XhoI) and ligated into the pGL3-basic vector cut with the same enzymes. To generate the human RARb RARE positive control plasmid, synthetic oligonucleotides (Table 1) were annealed and ligated into the pGL3-promoter plasmid cut with MluI and NheI. The human and mouse 1X RARE constructs were constructed by annealing synthetic oligonucleotides (Table 1), which contained SacI and XhoI overhangs. They were then cloned into the pGL3-promoter vector cut with the same enzymes. The 2X RAREs were constructed in a similar fashion; oligos were annealed which contained SacI and XhoI overhangs as well as a SacI cut site. This was then cloned into the pGL3-promoter vector which had been cut with SacI and XhoI. The SacI overhang was designed to obliterate this SacI cut site when cloned into the vector. To construct the 4X RAREs, the 2X RARE constructs were cut with SacI (at the site added by the 2X oligo) and XhoI and a second annealed 2X RARE oligonucleotide was ligated into this site. The human 31 bp deletion, hM-1, hM-2, hM-3, hM-4, hM-1/2/3/4 and the h-M1/4 mutant were all prepared with the Stratagene QuikChangeÒ Site-Directed Mutagenesis Kit (Stratagene; San Diego, CA) using the oligonucleotides listed in Table 1;

mutations are in lowercase. The human 31 bp deletion, hM-1, hM-2, hM-3, and hM-4 all used the (1670, +15) construct as template. The hM-1/4 construct was made using the hM-4 primers on the hM-1 plasmid as template. The hM-1/2/3/4 was made using the hM-2/3 primers on the hM-1/4 plasmid as template. The NEDD9 containing region of constructs subjected to mutagenesis was sequenced, and was then re-subcloned into the parent pGL3basic vector backbone that had not undergone PCR mutagenesis and was then sequenced. All constructs were prepped with the QIAprep Spin Miniprep Kit (QIAGEN; Valencia, CA) prior to sequencing, and using the QIAfilter Plasmid Midi and Maxi Kit for transfection studies. Cell culture and transfections MCF-7 (human breast carcinoma) and COS-1 (Green African monkey kidney) cells were obtained from the American Type Culture Collection (ATCC, Rockville, MD) and SH-SY5Y (human neuroblastoma) cells were a gift from Dr. Biedler (Memorial Sloan-Kettering Cancer Center, New York, NY). MCF-7 and SH-SY5Y cells were cultured as described previously [18,19]. COS-1 were cultured in high glucose DMEM and supplemented with sodium bicarbonate (1.5 g/ L) and 10% heat inactivated fetal bovine serum. SH-SY5Y cells were harvested by gentle agitation; MCF-7 and COS-1 cells were treated with Accutase (Innovative Cell Technologies, Inc.; San Diego, CA). Eighteen to twenty-two hours prior to transfection, cells were plated at a density of 105 cells/well in 12-well tissue culture plates. Cells were transfected with reporter construct (0.3 lg) and pCMVb (0.015 lg) as an internal control for transfection efficiency; where indicated, hRARa [20] and mRXRa [21] were each used at 0.1 lg. FuGENE (Roche Diagnostics, Mannheim, Germany) was used at 0.75 ll/well. Cells were dosed 24 h following the transfection with vehicle (ethanol, 0.1%, or a combination of ethanol 0.1% + DMSO 0.1% for the RAR/RXR ligand experiment) or to a final concentration of atRA (1010 to 106 M; Spectrum Chemical Company (New Brunswick, USA) and was deemed greater than 99% pure by reverse-phase HPLC [22], TTNPB (109 to 105 M; RAR-selective agonist; Sigma, St. Louis, MO) and/or LGD1069 (108 to 106 M; RXR-selective agonist; kind gift from Dr. R.W. Curley Jr., Ohio State University). The cells were washed twice with 4 °C phosphatebuffered saline (PBS, pH 7.4), followed by the addition of 200 ll reporter lysis buffer (Promega, Madison, WI) and incubation at room temperature for 5 min. Cell extracts were collected after they were loosened from the plate by agitation and scraping using a plastic pipette tip, and the lysate was stored at 20 °C until use. On the day of the assay, cells were thawed, quickly vortexed and subjected to centrifugation at room temp in a table top microfuge (Eppendorf centrifuge 5415 c) at 14,000 rpm for 2 min. Cell supernatent (20 ll) was transferred to a 96 well plate, 50 ll of luciferin substrate was added (20 mM Tricine, 1 mM Mg(CO3)4Mg(OH)2, 2.7 mM MgSO4, 0.1 mM EDTA, 1 mM DTT, 270 lM acetyl CoA, 470 lM D-luciferin, 526 lM ATP) and luciferase activity was assayed using a MLX Microtiter Plate Luminometer (Dynex Technologies; Chantilly, VA). b-galactosidase activity was assayed using the Galacto-LightTM b-Galactosidase Reporter Gene Assay System, (Applied Biosystems; Foster City, CA). Luciferase values were normalized to the b-galactosidase values from the pCVMb internal control plasmid. Normalized reporter construct values were then divided by normalized values for the appropriate parent vector (pGL3-basic or pGL3-promoter), except Fig. 5C where the pGL3basic data is graphed. Electrophoretic mobility shift assay (EMSA) The assay was performed as previously described [1]. The oligonucleotides used in the gel shifts are described in Table 1. Human

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Table 1 Primer Sequences

Mutation substitutions are in small bold type PCR product size: h2A-3 (123 bp), h2B-3 (201 bp), hExon 6 (159 bp), hRAR(b RARE (339 bp), hGAPDH (520 bp), hNEDD9 RARE region (178 bp), hNEDD9 Exon 6 region (159 bp) Annealing temperature: h2A-3 (64°), h2B-3 (64°), hExon 6 (64°), hRARb RARE (64°), hGAPDH (60°) Extension time: h2A-3 (5 s), h2B-3 (8 s), hExon 6 (7 s), hRAR(b RARE (15 s), hGAPDH (20 s)

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RARc1 and murine RXRc were prepared as described [23,24]. The human RARc1-specific antibody was purchased from ABR-Affinity BioReagents (Golden, CO). RNA Isolation and RT-PCR Cells were treated with either vehicle (ethanol, 0.2% final), or atRA (106 or 107 M) for the times indicated. Total and poly(A)+ RNA was isolated from SH-SY5Y and MCF-7 cells as described [25]. mRNA (1.0 or 0.5 lg, respectively) was reverse transcribed with SuperScript III reverse transcriptase (Invitrogen), random hexamers and diluted to 125 ll. PCR studies (5 ll/rxn) were conducted in triplicate, and each sample was normalized to GAPDH. Human primers anchored in exons 2A and 3, 2B and 3, and in exon 6, as well as RARb-specific primers are listed in Table 1. Quantitative PCR studies were performed as described previously using the real-time LightCycler system (Roche, Indianapolis, IN) with the LightCycler faststart DNA master SYBR green 1 kit. Annealing temperatures and extension times used are listed in Table 1. All primers produced products that had a single melting peak and the appropriate sized band when analyzed on an ethidium bromidestained agarose gel. PCR product quality was monitored using post-PCR melt curve analysis at the end of the amplification cycles. mRNA half-life was calculated with GraphPad Prism software (San Diego, CA) and the nonlinear regression (curve fit), in order to calculate the value for k (shown below). Half-life (T1/2) was then calculated using the equation: Y = (TOP–BOTTOM)  e(kx) + BOTTOM; T1/2 = ln (2)/k. ChIP assays SH-SY5Y cells were seeded in 100-mm plates and allowed to become confluent. Cells were then treated for 4 h with either vehicle or atRA (106 M). Nuclear proteins were cross-linked to DNA by adding formaldehyde directly to the medium to a final concentration of 1% for 15 min at room temperature on a rocking platform. Cross-linking was stopped by adding glycine to a final concentration of 0.125 M and shaking for 5 min. The medium was removed, the cells were washed twice with ice cold PBS and collected by scraping into ice cold PBS supplemented with complete protease inhibitor tablets (cat. no. 11836153001) used as per manufacturers directions (Roche, Mannheim, Germany). After centrifugation at 1200 rpm (Eppendorf centrifuge 5415 c) for 5 min at 4 °C the cell pellets were frozen, thawed, and then resuspended in nuclei lysis buffer (1% SDS, 10 mM EDTA, 50 mM Tris–HCl, pH 8.1, and Roche protease inhibitor pellet) and the lysates were sonicated on ice to produce DNA fragments of 300 to 1000 bp in length with the Fisher

Scientific Sonic Dismembrator, Model 100 sonicator (power setting: 4, 3  15 s). Cellular debris was removed by centrifugation (14,000 rpm) and the lysates were diluted 1:10 in ChIP dilution buffer (0.01% SDS, 1.1% Triton X-100, 1.2 mM EDTA, 16.7 mM NaCl, Roche protease inhibitor pellets, 16.7 mM Tris–HCl, pH 8.1). Chromatin solutions were precleared with protein A sephrose overnight at 4 °C with rotation. The solution (3%; inputs) was set aside until cross-links were reversed. Chromatin solutions (17% of the solution for each immunoprecipitation) were incubated with 30 ll of the indicated antibodies for 3 h at 4 °C with rotation. The antibodies against RXR (DN 197) sc-774, RARa (C-20) sc-551, or normal rabbit IgG sc-2027, were obtained from Santa Cruz Biotechnologies (Santa Cruz, CA). The immune complexes were collected with 30 ll protein A agarose for 2 h at 4 °C with rotation. The beads were pelleted by centrifugation for 1 min at 4 °C at 7500 rpm in a microfuge and washed sequentially with 500 ll of the following buffers: twice with low salt wash buffer (0.1% SDS, 1% Triton X-100, 2 mM EDTA, 50 mM NaCl, 20 mM Tris–HCl, pH 8.1), and LiCl wash buffer (250 mM LiCl, 1% Nonidet P-40, 1% sodium deoxycholate, 1 mM EDTA, 10 mM Tris–HCl, pH 8.1). Finally, the beads were washed twice with 1 ml TE buffer (1 mM EDTA, 10 mM Tris–HCl, pH 8.1). The immuno-complexes were eluted by adding 150 ll elution buffer (1% SDS, 100 mM NaHCO3), vortexing 20 s and placing at 65 °C for 10 min and then vortexing again for 20 s. After centrifugation at 7500 rpm for 1 min at room temperature, the supernatant was collected, the entire elution process was repeated a second time for a total elution of 300 ll. The cross-linking was then reversed by adding NaCl to a final concentration of 0.3 M and RNaseA to a final concentration of 3 pg/ll and incubating overnight at 65 °C. The remaining proteins were digested by adding proteinase K (final concentration 40 lg/ml) and incubating for 3 h at 45 °C. The DNA was recovered using the QIAquick PCR Purification Kit (QIAGEN; Valencia, CA) according to directions. Inputs and immunoprecipitations were eluted in water and then subjected to PCR using primers designed to amplify appropriate fragments of either the proximal human NEDD9 promoter region or exon 6 region. The PCR primers are listed in Table 1. The PCR products were separated by electrophoresis through 2.0% agarose that was post stained with ethidium bromide. Chromatin sonication was evaluated by running inputs on a 1.6% agarose gel supplemented with 0.5 lg/ml ethidium bromide. In silico search for transcription binding motifs The region from 485 to 435, which included the NEDD9 RARE, was used to search for additional transcription factor binding motifs. The region was searched using 3 programs; Consite

Fig. 1. Schematic of the NEDD9 gene structure showing the conserved location and sequence of the RARE in human, mouse and rat. The exons in the NEDD9 gene are indicated by white boxes (non-coding) and grey boxes (coding) and are shown to scale; exons are numbered above according to [26]. Transcriptional initiation of the 2A transcript starts with exon 1A whereas the initiation site for the 2B transcript starts with exon 2B. The two major transcripts (2A and 2B) are indicated by connecting lines with intron distances listed below. Translation initiation sites in exons 2A and 2B (black arrows) and the translation stop site (black hatch mark) in exon 8 are indicated. The coding sequence in exons 3–8 is common to both the 2A and 2B transcripts, the 4 amino acids specific to each transcript are shown above the translational start site. Shown above in bold text is the putative RARE with the hexameric half-sites boxed in black, it is located from 475 to 445 bp (relative to the transcriptional start site) upstream of the 2B-exon. The element is comprised of overlapping 50 DR1, DR5 and 30 DR1 sequences.

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(http://asp.ii.uib.no:8090/cgi-bin/CONSITE/consite/) and Alibaba2 and Match both available through http://www.gene-regulation.com/. Results atRA Specifically upregulates the NEDD9 transcript that initiates in exon 2B The two mRNA splice variants (2A and 2B) are produced from the NEDD9 gene, and encode for proteins that differ in only three amino acids at the amino terminus (Fig. 1) [26]. The two variants arise from two translation initiation sites in exons 2A and 2B (Fig. 1, black arrows). Thus, there are two distinct NEDD9 promoters which can generate mRNAs that encode for proteins that are largely conserved in structure. Previous work from our group showed that NEDD9 mRNA is induced in SH-SY5Y and MCF-7 cells by atRA, however, the probes used in these studies did not discriminate between the 2A and 2B transcripts [1,2]. In order to determine whether atRA regulates one or both of these transcripts in cells, we designed primers spanning from exon 2A to exon 3, from 2B into exon 3, or primers common to both the 2A and 2B transcripts anchored in exon 6 (Fig. 2A). RT-PCR analysis of mRNA isolated from SH-SY5Y cells revealed that the 2A transcript was expressed at low but detectable levels in untreated cells, and exposure to atRA did not increase the amount of transcript produced. The 2B transcript was expressed at higher levels in vehicle-treated cells, and showed a 2.2-fold induction by atRA, mirroring the induction seen using the common exon 6 primer pair (Fig. 2B, left). Similarly, in MCF-7 cells, a 3-fold increase in the expression of the 2B transcript was observed in atRA-treated cells, whereas the 2A transcript was not induced (Fig. 2B, right). Thus, mRNAs that include the exon 2B-exon 3 region are induced by atRA both in SYSY5Y and MCF-7 cells. Furthermore, the fold-induction of the 2B3 product, measured after the exposure of cells to atRA, closely matches that observed using the exon 6 primer pair that amplifies both transcripts showing that induction of the 2B transcript can account for the full retinoid effect on increased NEDD9 transcription. RA induction of the NEDD9 2B-3 mRNA transcript requires transcription but not new protein synthesis, and retinoid does not alter mRNA stability To assess the relative importance of RNA transcription and new protein synthesis on the induction of the NEDD9 2B transcript, SHSY5Y cells were treated with actinomycin D for 4 h in the presence

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or absence of atRA, or were pretreated for 30 min with cycloheximide (a protein synthesis inhibitor that blocks translational elongation) followed by an additional 4 h with atRA or vehicle, and NEDD9 mRNA was measured by quantitative PCR. The induction of NEDD9 mRNA by atRA at 4 h was blocked in SH-SY5Y cells when actinomycin D was added (Fig. 3, inset). Similar results were obtained using the exon 6 primers as well as those specific to the 2B transcript (not shown). atRA-induction of a control mRNA, RARb, showed the expected induction by atRA in SH-SY5Y cells (8.1-fold), and this effect was eliminated in the presence of actinomycin D (data not shown). In contrast, neither the induction of the NEDD9 or RARb mRNAs by atRA was changed when cycloheximide was present. These results show that the induction of NEDD9 mRNA does not require new protein synthesis and may represent a direct transcriptional effect of atRA on the expression of this transcript. In order to determine whether atRA could also be increasing the steady-state levels of NEDD9 by stabilizing the mRNA, the rate of transcript decay was compared in vehicle and atRA-treated cells. SH-SY5Y cells pretreated with vehicle or atRA for 24 h followed by the addition of actinomycin D showed no difference in the rate of NEDD9 mRNA disappearance over the course of an 8 h experiment (Fig. 3). A second study in which samples were evaluated at shorter time intervals (0, 0.5, 1.0, and 1.5 h) confirmed that atRA did not alter NEDD9 mRNA stability (not shown). The NEDD9 mRNA had a half-life of 29 ± 2 min in vehicle-treated cells and did not differ from that in cells treated with atRA (34 ± 3 min). Taken together, these results show that that the induction of the NEDD9 gene by atRA results from an increase in transcription. The NEDD9 2B promoter contains a RARE that confers responsiveness to atRA We previously identified a putative RARE consisting of four hexameric half-sites separated by 1, 5 and 1 bp upstream of the transcriptional initiation site of the NEDD9 2B transcript, and showed that this 31 bp sequence bound specifically to the RXR/RAR in a gel mobility shift assay [1]. This element is highly conserved across human, mouse and rat (Fig. 1). To test whether this RARE alone was sufficient to drive transcription in response to atRA, reporter constructs were prepared which placed 1, 2, or 4 copies of the human or mouse NEDD9 RARE in series, upstream of a luciferase reporter and SV40 promoter in the pGL3-promoter vector (Fig. 4A). As a positive control, a 70 bp region of the human RARb promoter containing a well-defined DR5-type RARE [27,28] was also cloned into the same vector. MCF-7 and COS-1 cells were chosen for study because

Fig. 2. atRA induces the NEDD9 2B and not the 2A transcript in SH-SY5Y and MCF-7 cells. (A) The positions of primers used to detect the NEDD9 transcripts, 2A, 2B and exon 6, are shown (arrows); the product size is shown to the right. (B) Fold-induction of NEDD9 mRNAs in response to atRA treatment is shown for primer sets to exon 6 (detects both the 2A and 2B transcripts), primers spanning exon 2B to exon 3 (2B transcript only), and for exon 2A to exon 3 (2A transcript only). SH-SY5Y and MCF-7 cells were dosed for 24 h with vehicle (0.1% ethanol) or atRA (106 M) followed by Poly(A)+ RNA isolation and analysis by quantitative PCR assay. Primer sequences and PCR conditions are listed in Table 1. Fold-changes from vehicle were calculated after normalization to GAPDH.

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Fig. 3. The induction of NEDD9 mRNA by atRA occurs at the level of transcription. atRA does not alter the stability of the NEDD9 transcript, and NEDD9 induction by atRA is eliminated by actinomycin D but not by cycloheximide (inset). To assess the importance of new gene transcription versus protein synthesis, SH-SY5Y cells were treated for 4 h with vehicle (ethanol), atRA (106 M), vehicle + actinomycin D (Act D, 5 lg/ml) or were pretreated for 30 min with cycloheximide (CHX, 10 lg/ml) followed by ethanol or atRA (106 M) for an additional 4 h. Total RNA was reverse transcribed and analyzed by quantitative PCR using the exon 6-specific primer set (Table 1). Fold-changes from vehicle were calculated after normalization to GAPDH and are shown. For RNA stability studies, SH-SY5Y cells were treated for 24 h with vehicle (ethanol) or atRA (106 M) at which time Act D (5 lg/ml) was added, and the quantity of NEDD9 mRNA remaining at each time point was assessed by RT-PCR. All data was normalized to GAPDH. mRNA half-life was calculated with Prism software and nonlinear regression (curve fit).

they are transfected at a higher efficiency than SH-SY5Y cells. Transfection of the MCF-7 cell line with all three constructs resulted in an increase in luciferase activity in atRA-treated compared to vehicle-treated cells. Induction of reporter gene activity by atRA was observed both in the presence of co-transfected RARa

and RXRa nuclear receptors (Fig. 4C) and in MCF-7 cells containing only their native complement of nuclear retinoid receptors (Fig. 4B). In the absence of receptor co-transfection, there was an increase in the reporter gene activity as the number of RARE elements increased. With receptor co-transfection, maximal gene induction was observed in the presence of the 2X element. In a second cell line (COS-1) transfected with human or mouse 1X, 2X, and 4X RARE constructs, reporter gene activity was increased in cells exposed to atRA compared to vehicle (Fig. 4D). Thus, a single copy of either the human or mouse NEDD9 RARE is sufficient to mediate responsiveness to atRA in both MCF-7 and COS-1 cells. Next, the ability of the RARE, within the context of the normal flanking sequence and driven by the endogenous NEDD9 promoter, to mediate an increase in gene transcription in response to atRA was assessed. We placed 2756 and 1670 bp of sequence upstream of the human NEDD9 2B transcriptional start site into the promoterless pGL3-basic luciferase reporter vector, and measured reporter gene activity resulting from each of these two constructs after exposure to vehicle and atRA (Fig. 5A). Both the longer (2756 to +15) and shorter (1670 to +15) regions of NEDD9 2B native promoter were inducible by atRA (5.9 ± 0.9 and 12.5 ± 1.9fold, respectively; Fig. 5B). While the basal luciferase activity of both constructs was similar (not shown), the shorter promoter piece showed a more robust response to atRA. In a time-course experiment, the shorter construct was significantly induced by 2.3-fold after 4 h (Fig. 5D) whereas it took the longer piece 12 h to achieve the same level of response (2.5-fold, Fig. 5E). In a dose response experiment, the shorter construct showed significant response to as little as 108 M atRA (2.7-fold; p < 0.05), whereas a similar fold-induction of the larger construct required 10-fold more atRA (Fig. 5C). Taken together, this work shows that both reporter constructs within the context of the native NEDD9 promoter are responsive to atRA, and further, suggests that the longer con-

Fig. 4. The NEDD9 RARE placed upstream of a heterologous promoter and a reporter gene confers responsiveness to atRA. (A) Schematic of constructs containing 1, 2, or 4 copies of the NEDD9 RARE placed 50 of the SV40 promoter and luciferase reporter gene; the sequence of the human NEDD9 RARE is shown for reference and the hexameric half-sites are in black, each grey box represents one copy of the 31 bp RARE sequence. (B) Reporter gene activity in atRA-treated MCF-7 cells transfected with the 1X, 2X, or 4X NEDD9 RARE or the hRARb RARE in the absence of exogenous RAR and RXR. (C) As described in B but cells were also co-transfected with RARa and RXRa expression plasmids. (D) Reporter gene activity of human and murine 1X, 2X, and 4X constructs in COS-1 cells co-transfected with RARa and RXRa expression plasmids. In experiments, cells were seeded in 12-well plates and were transfected with reporter plasmid, pCMVb (internal control), and where indicated, RARa and RXRa nuclear receptors using FuGENE 6 and evaluated for both luciferase and b-gal activity as described in Methods. Data has been normalized to the pGL3-promoter parent vector. Values are mean ± standard error.

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Fig. 5. The native NEDD9 2B promoter containing the RARE is capable of mediating the response to atRA. (A) Schematic of the longer and shorter construct containing the native NEDD9 promoter and containing the RARE (depicted by the grey box). The numbers below the construct indicate the bp location relative to the transcriptional start site for the 2B mRNA. (B) Fold-induction of the constructs (2756, +15) and (1670, +15) in response to atRA (106 M) after 24 h. (C) Response of a NEDD9 reporter construct or the pGL3-basic parent vector after 24 h of exposure to increasing doses of atRA. Statistical analysis was performed using a one-way ANOVA with Bonferroni posthoc analysis (P < 0.05) comparing atRA to vehicle. (D) Time-course of NEDD9 reporter activity in cells exposed to 106 M atRA compared to vehicle. Statistical analysis was performed using a two-way ANOVA with Bonferroni posthoc analysis (P < 0.05). In all panels MCF-7 cells were transfected with reporter plasmid, pCMVb (internal control), and RARa and RXRa nuclear receptors using FuGENE 6 and evaluated for both luciferase and b-gal activity as described in Methods. In B and D data has been normalized to the pGL3basic parent vector.

struct could include negative regulatory element(s) that is (are) absent in the shorter construct. In order to determine whether the RARE in the NEDD9 promoter construct could account for all of the atRA-increase in reporter activity, the 31 bp RARE was removed from the 1670 bp construct by mutagenesis and the resulting construct was tested for atRA-responsiveness in the reporter gene assay. After 24 h of exposure to atRA, the mutant construct showed less than a 2-fold increase in reporter activity whereas the wild-type construct containing the intact RARE showed a 13fold induction (Fig. 6). A second time-course experiment in which atRA response was measured after 16, 24, and 48 h confirmed these initial observations (data not shown). Taken together, these results show that the 31 bp RARE is responsible for mediating the atRA regulatory effects on the NEDD9 2B promoter.

Both the full-length RARE and individual DR sequences can bind to the RXR/RAR in vitro, but the intact RARE is required for fullresponsiveness to atRA In order to determine whether the DR sequences contained within the NEDD9 RARE were individually capable of interacting with the RXR/RAR complex, each DR sequence was radiolabeled

Activation of the RARE occurs effectively with a RAR-specific ligand, but not a RXR-specific ligand The ability of RAR and RXR-specific ligands to activate luciferase in the 1670 to +15 bp NEDD9 promoter construct was tested, and showed that activation occurred over a range of concentrations when the RAR-specific ligand, TTNPB, was used (108 to 105 M). In contrast, much less of an increase in reporter gene activation was noted at the highest concentration of RXR-specific ligand, LGD1069 (106 M) when used alone. This suggests that the majority, if not all of the response, can be accounted for by the formation of a transcription activating complex containing the RAR protein, as opposed to a RXR homodimer. Interestingly, the addition of both the RAR and RXR-selective ligand at suboptimal doses resulted in an increase in activity over that produced by either agent alone (Fig. 7).

Fig. 6. The 31bp RARE region is required for induction of the NEDD9 2B promoter by atRA. Reporter gene activity in MCF-7 cells transfected with either native NEDD9 promoter containing the RARE (1670, +15) construct or the same construct in which the 31 bp RARE is deleted, pCMVb (internal control), and RARa and RXRa nuclear receptors using FuGENE 6. Data has been normalized to the pGL3-basic parent vector.

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Fig. 7. Activation of the RARE occurs effectively with a RAR-specific ligand, but not a RXR-specific ligand. The shorter NEDD9 promoter construct (1670, +15) was used in transient transfection assays and cells were dosed with either vehicle (0.1% ethanol, 0.1% DMSO), atRA (109 to 106 M), a RAR-selective ligand (TTNPB 109 to 105 M), a RXR-selective ligand (LGD1069 108 to 106 M) or both (107 LGD1069 plus 108 to 105 M TTNPB). Cells, seeded in 12-well plates, were transfected with either the RARE containing (1670, +15) construct, or pGL3-basic pCMVb (internal control), and RARa and RXRa nuclear receptors using FuGENE 6 and evaluated for both luciferase and b-gal activity as described in Methods. Data has been normalized to the pGL3-basic parent vector.

and studied by electrophoretic mobility shift assay. The results showed that all three radiolabeled DR elements (50 hDR1, hDR5, and 30 hDR1) were individually capable of producing a shift when combined with RAR and RXR, whereas relatively no binding occurred when only one receptor type was added (Fig. 8A, data not shown). The addition of excess unlabeled probe (human full-length complex element, 50 hDR1, hDR5, or 30 hDR1, respectively) competed specifically with the radiolabeled element for binding to the RXR/RAR complex, whereas an unlabeled mutant probe (hM1/2/3/4, 50 hDR1-M1, hDR5-M2, or 30 hDR1-M4, respectively) did not. The presence of RAR nuclear receptor in the shifted complex was confirmed using an RARc-specific antibody which supershifted the complex. In order to test the importance of each of the four half-sites in RXR/RAR complex binding, individual mutations were made in each half-site sequence (Fig. 9; hM-1, hM-2, hM-3, and hM-4), and another oligonucleotide containing mutations in all 4 halfsites (hM-1/2/3/4) was also synthesized (the sequence of individual mutants is shown in Fig. 9). When used in gel shifts, both the wild-type RARE as well as the individual hM-1, hM-2, hM-3, and hM-4 mutants showed a band shift in the EMSA, although mutation in either half-site of the DR5 element appeared to reduce the affinity of the RXR/RAR/DNA interaction. When AcNPV nuclear extract was used in place of that containing the over-expressed nuclear receptors in the incubation, neither the wild-type nor mutant (hM-1, hM-2, hM-3, and hM-4) elements shifted, and furthermore, neither receptor when added alone produced a shift (data not shown). In contrast, mutation of all four half-site elements completely eliminated the formation of a complex between the RXR/ RAR and the radiolabeled mutant (Fig. 8B). Thus, at least one of the three DR elements (50 DR1, DR5, or 30 DR1) must be intact to preserve RXR/RAR binding in vitro. In order to evaluate whether the entire 31 bp element was needed to confer full-responsiveness to atRA, constructs containing mutations in each individual half-site of the 31 bp RARE in the NEDD9 2B native promoter luciferase construct (1670 to +15 bp) were tested for responsiveness to atRA. Additional con-

structs containing mutations in both the 50 and 30 DR1 elements (hM-1/4), which leaves the DR5 intact, and a construct with mutations in all 4 half-sites (hM-1/2/3/4), which leaves no DR intact, were also studied. As a control, the DR elements containing individual half-site point mutations were first radiolabeled and tested directly in the EMSA assay and none were capable of forming a complex (50 hDR1-M1, hDR5-M2, or 30 hDR1-M4; data not shown). When the contribution of each of the four half-sites was examined in a functional transient transfection assay, mutation of each individual half-site significantly decreased the responsiveness to atRA (Fig. 9). The effect of mutating the 50 DR1 element reduced response to half that of the wild-type element (4.4- compared to 9.1-fold), whereas all the remaining mutant constructs showed activity ranging from 1.3- to less than 1.9-fold. These results show that despite the ability of individual half-site mutants to bind to RXR/RAR in vitro, all four half-sites are required for maximum induction by atRA in cultured cells. Since our construct contains a highly conserved DR5, we designed a construct in which the 1st and 4th half-sites were mutated leaving the DR5 intact. Surprisingly, this construct had essentially no atRA induced activity indicating that base pairs in addition to those that compose the two half-sites of the DR5 are essential for induction. A construct in which all four half-sites were mutated showed no induction by atRA, again showing that all the atRA activity could be accounted for with the 31 bp region, and more specifically, with the four half-site elements. The NEDD9 promoter binds RAR and RXR nuclear receptors In order to investigate whether retinoid receptors associate with the NEDD9 promoter in live cells, ChIP assays were performed in atRA and vehicle-treated SH-SY5Y cells. For this purpose we designed primer pairs for the detection of the NEDD9 promoter region that contains the RARE as well as a region in exon 6 selected to be a negative control for the specificity of the antibodies (Table 1). Chromatin was extracted from SH-SY5Y cells that had been exposed to vehicle or atRA (106 M) for 4 h. In the absence of atRA the NEDD9 RARE region, but not the downstream exon 6 region, were immunoprecipitated by antibodies to the RARa and RXR receptors. Treating cells with atRA for 4 h did not appear to significantly affect the level of association between nuclear receptors and chromatin (neither positively nor negatively) at the RARE nor exon 6 region (Fig. 10). Discussion In previous work, we identified NEDD9 as a novel atRA-responsive gene in SH-SY5Y cells [1]. These cells respond to atRA by differentiating and extending neurites in culture and the induction of NEDD9 is one of the earliest gene changes that can be detected in these cells. In vivo, the expression of NEDD9 mRNA is also rapidly altered in the developing hindbrain of embryos exposed to excess atRA [2]. In the present report, we show that of the two possible NEDD9 mRNA splice variants (2A and 2B); only the 2B form is induced by atRA in SH-SY5Y and MCF-7 cells. We also show that the induction of the 2B NEDD9 mRNA results from an increase in transcription, and we now report on the activity of a complex element consisting of four half-sites located in the 50 region of the NEDD9 2B promoter that serves as a functional retinoic acid response element (RARE). The NEDD9 RARE is a highly conserved complex element composed of four half-sites separated by 1, 5, and 1 base pairs. The hexameric half-sites within many RAREs are spaced by 5 base pairs, although they can also be spaced by 2 or 1 [5,28]. One of the most potent RAREs is the DR5 sequence found in the promoter of the hRARb gene. The four individual half-sites within the complex NEDD9 RARE show a high degree of similarity to the defined min-

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Fig. 8. The 50 DR1 and 30 DR1 and the DR5 element are all capable of binding RAR and RXR nuclear receptors. (A) EMSA of the intact 31 bp RARE, as well as each individual DR pair: h50 DR1, hDR5 and h30 DR1, in the presence of added RARc1 and RXRc protein. Specificity of the complex (indicated by black arrow) was tested by adding a 100-fold excess of each specific oligonucleotide, either a wild-type or mutant version, in unlabeled form (sequences listed in Table 1, mutations in small case). The complex is super shifted with the addition of RARc1-specific antibody, indicated by black bracket, but not a nonspecific antibody (data not shown for all constructs.) (B) The effects of individual half-site mutations in the context of the complex element were tested by the addition of RARc1 and RXRc nuclear receptor in EMSA assays. The position of the RAR/ RXR complex is indicated by an arrow. The ability to compete away complex formation with cold RARb DR5 probe is also shown. A construct in which all four half-sites have been mutated, M-1/2/3/4, does not produce a shift. Complexes were resolved on 4% non-denaturing polyacrylamide gels, dried, and visualized using autoradiography.

imal RARE half-site consensus sequence (PuG(G/T)TCA). Two of the four half-sites are a perfect match (2nd and 4th), whereas the 1st and 3rd, each vary from the consensus sequence at one position. All four of these half-sites are found in RAREs residing within the promoter region of other RA regulated genes (see Table 1 in [29] and references therein; [30–35]). When tested by EMSA in the presence of both RAR and RXR proteins, each individual DR (50 hDR1, hDR5, and 30 hDR1) is efficiently shifted, as is the intact complex 31 base pair element indicating that at least in vitro, each of these isolated sequences can serve as a binding site for the receptor heterodimer. However, it is clear that more than a single direct repeat sequence is important, as mutational analysis within the context of the native NEDD9 promoter sequence shows that all four half-sites are required for full-responsiveness to atRA in a cell reporter assay. Interestingly, the two central hexameric half-sites of the NEDD9 RARE which comprise a DR5, are identical in sequence to the two half-sites in the DR5 of the human CRABPII RARE [32], with only the intervening and flanking nucleotides being different. However, when the first and fourth half-sites of the NEDD9 RARE are mutated so that only the two half-sites matching the CRABPII RARE remain, we still observe a nearly complete loss of atRA-responsiveness in reporter assays (from 9.5- to 1.3-fold). This

shows that the DR5 alone is insufficient to mediate atRA-responsiveness within the context of the native NEDD9 promoter sequence, and that both the 50 and 30 flanking sequences, each containing an additional half-site element, have an important influence on activity. Not all RAREs appear as simple DR5, DR2 and DR1 type elements, and more complex elements have been described. In particular, RAREs have been found which have variations in sequence, spacing, and complexity [5,29,36–41]. In the Cyp26A1 gene, two DR5s are both required for optimal RA induction, and although these two RAREs are spaced approximately 1500 bp apart, they appear to act synergistically [42,43]. A number of other genes contain elements comprised of more than two closely spaced half-sites, and as is seen with NEDD9, all the individual half-site elements are required to maintain full-responsiveness to atRA. The a-fetoprotein promoter contains a complex element composed of three direct repeats and one inverted repeat. Mutation analysis and transient transfection indicate a requirement for all four of the half-sites of the complex element for full transcriptional response to RA, although this element has been described to function as a RXRE as opposed to a RARE [44]. The human surfactant protein B promoter contains a complex element composed

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Fig. 9. All four half-sites of the complex HEF1 RARE are necessary for maximal induction by atRA. Point mutations were made to each of the individual half-sites in the context of the complex element and the (1670, +15) promoter, sequence is shown in left panel mutations are underlined. Constructs which contained mutations in both the 1st and 4th half-site leaving the DR5 intact and mutations in all four half-sites were also tested. The transcriptional activity of each construct after transfection into MCF-7 cells, as described in Fig. 7, is shown in the right panel. Point mutations to the individual half-sites significantly reduce the response to atRA. Mutation to all four half-sites or deletion of the element further eliminates the response to atRA. Data has been normalized to the pGL3-basic parent vector.

of three half-sites organized as a DR5 and DR9, and mutations in each of the three half-sites reduce atRA-induction in transient transfection assays [35]. Similarly, the mouse transglutaminase gene promoter contains a RARE (mTGRRE1) consisting of three half-sites separated by 5 and 7 bp each [45]. The full tripartite element is required for RARE activity in culture, whereas the distal two half-sites are sufficient to confer RAR/RXR binding in vitro. What sets the NEDD9 RARE apart from many previously reported complex type elements is that it does not show similar degeneracy of many of the half-site sequences, and furthermore, other complex elements are often only weakly responsive to atRA in cell transfection assays [5,28]. Thus, while the NEDD9 RARE is complex, it retains half-site sequences that adhere closely to the consensus RARE half-site motif, and the 31 base pair element within the context of the intact promoter is highly responsive to atRA. Non-canonical or complex RAREs are being recognized as a more common feature of naturally occurring hormone response elements (HRE). This type of element may, in fact, be under reported due to bias in the methods used to search for and assess the activity of HREs. Importantly, for many RAREs, half-site elements may not exactly match the consensus, making in silico screening difficult. Transfection reporter analysis of an isolated putative DR would not take into consideration the influence of flanking sequence or more remotely located elements that might exert a critical effect on responsiveness to RA. Likewise, attributing the hormone responsiveness of a longer stretch of DNA to only a DR element identified in silico would not rule out a contribution from unknown flanking elements including non-canonical half-sites unless more extensive deletion and mutational analysis was performed. There are new approaches that have emerged that should improve the identification and characterization of genomic sites of protein–DNA interaction, including ChIP-based and chromosome capture conformation (3C) assays. Several recent reviews discuss the extent to which these and other newer techniques have been applied in the nuclear receptor field [46,47]. The mechanism whereby the four half-sites are needed to confer full activation by retinoic acid is unclear. In transient transfec-

Fig. 10. RAR and RXR are associated with the RARE in the NEDD9 2B promoter in intact cells. ChIP analysis of RAR and RXR binding to regions of the NEDD9 gene in confluent SH-SY5Y cells treated for 4 h with either vehicle or atRA (106 M). Crosslinked cell lysates were subjected to immunoprecipitation with antibodies to RARa, RXR, or IgG (negative control) as indicated in Methods. DNA precipitates were isolated and then subjected to PCR (33 cycles) using the primers specific to the NEDD9 RARE or to a control region (NEDD9 exon 6) shown in Table 1. Primers to RAREs of hRARb and hCRABPII RAREs were used as positive controls and primers to downstream regions of hRARb were used as additional negative controls. We consistently observed very high levels of nuclear receptor association at both the hRARb RARE and hCRABPII RARE, these levels were 35–45 times the level of association detected at hRARb exon 2 and exon 5 (data not shown).

tion assays using a RAR or RXR-specific ligand alone, activation of the RARE occurs effectively with the RAR-specific ligand TTNPB but not the RXR-specific ligand LGD1069. However, when the ligands are used together, addition of RXR ligand potentiates the activity of suboptimal levels of TTNPB. This type of agonist effect is characteristic of activation through a RAR/RXR heterodimer [48]. The fact that the RXR ligand has little effect on its own, except at high concentrations (that are known to elicit some binding to the RAR [49]) suggests the RXR/RXR homodimers are not involved. Finally, synergistic transcriptional activation has been observed previously when agonists for both the RAR and RXR partners are added [50,51]. EMSA provides further evidence that NEDD9 activation is mediated through a RAR/RXR heterodimer, since only a combination of RAR and RXR produced a shift. In addition the ChIP assay,

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which detects receptor binding within the living cell, clearly shows that both the RAR and RXR are bound to the NEDD9 RARE. As previously reported, we find that the nuclear receptors are bound to the RARE, both in the absence and presence of atRA. This is consistent with work by Gillespie et al. (2007) [7], in which RARa and RXRc were associated with the Hoxa1, Cyp26A1, and RARb2 RAREs in the both the absence and presence of ligand. The cell-based ChIP assay, however, does not provide sufficient resolution to determine which of the NEDD9 halfsite sequences are bound to receptor. Mutational analysis suggests that the effect of mutating the first half-site element is the least detrimental to atRA response. However, the remaining half-sites are all clearly essential to confer a full response. It is possible that only one of the direct repeat elements interacts with the retinoid receptors, and that overlapping elements for other transcription factors may play a role in this response. An in silico search of the region from 485 to 435, which includes the NEDD9 RARE, identified three additional highly conserved motifs. Consite, Alibaba2 and Match programs, all identified motifs for the general transcription factor AP-1, and the chicken ovalbumin upstream promoter transcription factor (COUP-TF); the Match program also identified a hepatocyte nuclear factor 4 (HNF4) motif. All three of these motifs have been described in numerous retinoid-responsive promoters; AP-1 [33,52–54], COUP-TF [55,56], and HNF4 [53,57–59]. It is also possible that there is a cooperative interaction with nearby transcription factors outside of this 31 bp element. Whether additional transcription factors play a role in modulating the response of the NEDD9 RARE to atRA and its receptors remains to be determined. Alternatively, it is possible that the receptors may bind in a higher order structure as a complex to more than two of the half-site elements. At present, we cannot distinguish between these possibilities. In addition to the upregulation of NEDD9, atRA treatment of SH-SY5Y cells causes them to differentiate and undergo morphological changes including the extension of neurites and changes in adhesion [60]. Correspondingly, ectopic over expression of NEDD9 in PC12 cells has been shown to produce neurite-like processes [61]. It has also been shown that the NEDD9 2B transcript (shown here to be atRA regulated), but not the 2A transcript, is induced in neurons of the cerebral cortex and hippocampus 1–14 days following ischemia [26]. Therefore, it is possible that NEDD9 could be a directly regulated participant in the mechanism where atRA induces neurite outgrowth in SHSY5Y cells. This effect of atRA in SH-SY5Y cells is unlikely to result from the change in only one gene product and therefore it will be interesting to determine how the NEDD9, a protein known to play role in cell adhesion and shape changes, may be functioning. In summary, we have characterized the human NEDD9 2B promoter. Although NEDD9 contains two unique promoter regions, only one is regulated by atRA. Analysis of the promoter reveals the presence of a conserved complex RARE composed of four hexameric half-sites approximately 450 bp 50 of the transcriptional start site, all of which are required for full induction by atRA. Thus, the NEDD9 RARE is more complex in nature than the classic two hexameric half-site RARE. Finally, upregulation of NEDD9 is direct and results from an increase in transcription mediated via the binding of RAR and its heterdimeric partner RXR. Acknowledgments We thank Dr. Hector F. DeLuca, Dr. Richard S. Eisenstein and Dr. Emery H. Bresnick for editorial review of this manuscript.

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References [1] R.A. Merrill, A.W.-M. See, M.L. Wertheim, M. Clagett-Dame, Dev. Dyn. 231 (2004) 564–575. [2] R.A. Merrill, J.M. Ahrens, M.E. Kaiser, K.S. Federhart, V.Y. Poon, M. ClagettDame, Biol. Chem. 385 (2004) 605–614. [3] L.J. Gudas, M.B. Sporn, A.B. Roberts, in: M.B. Sporn, A.B. Roberts, D.S. Goodman (Eds.), The Retinoids: Biology, Chemistry, and Medicine, Raven Press, New York, 1994, pp. 443–520. [4] P. Germain, P. Chambon, G. Eichele, R.M. Evans, M.A. Lazar, M. Leid, A.R. De Lera, R. Lotan, D.J. Mangelsdorf, H. Gronemeyer, Pharmacol. Rev. 58 (2006) 712–725. [5] D.J. Mangelsdorf, K. Umesono, R.M. Evans, in: M.B. Sporn, A.B. Roberts, D.S. Goodman (Eds.), The Retinoids: Biology, Chemistry, and Medicine, Raven Press, New York, 1994, pp. 319–349. [6] P. Chambon, FASEB J. 10 (1996) 940–954. [7] R.F. Gillespie, L.J. Gudas, J. Mol. Biol. 372 (2007) 298–316. [8] D.M. Lonard, B.W. O’Malley, Mol. Cell 27 (2007) 691–700. [9] C.K. Glass, M.G. Rosenfeld, Genes Dev. 14 (2000) 121–141. [10] M. Clagett-Dame, E.M. McNeill, P.D. Muley, J. Neurobiol. 66 (2006) 739–756. [11] M. Clagett-Dame, H.F. DeLuca, Annu. Rev. Nutr. 22 (2002) 347–381. [12] A. Gavalas, R. Krumlauf, Curr. Opin. Genet. Dev. 10 (2000) 380–386. [13] M.K. Singh, L. Cowell, S. Seo, G.M. O’Neill, E.A. Golemis, Cell Biochem. Biophys. 48 (2007) 54–72. [14] S.F. Law, J. Estojak, B. Wang, T. Mysliwiec, G. Kruh, E.A. Golemis, Mol. Cell. Biol. 16 (1996) 3327–3337. [15] Y. Ohashi, S. Iwata, K. Kamiguchi, C. Morimoto, J. Immunol. 163 (1999) 3727– 3734. [16] G.A. van Seventer, H.J. Salmen, S.F. Law, G.M. O’Neill, M.M. Mullen, A.M. Franz, S.B. Kanner, E.A. Golemis, J.M. van Seventer, Eur. J. Immunol. 31 (2001) 1417– 1427. [17] S.J. Fashena, M.B. Einarson, G.M. O’Neill, C. Patriotis, E.A. Golemis, J. Cell Sci. 115 (2002) 99–111. [18] J.S. Chapman, K.L. Weiss, R.W. Curley Jr., M.A. Highland, M. Clagett-Dame, Arch. Biochem. Biophys. 419 (2003) 234–243. [19] M. Clagett-Dame, T.J. Verhalen, J.L. Biedler, J.J. Repa, Arch. Biochem. Biophys. 300 (1993) 684–693. [20] M. Petkovich, N.J. Brand, A. Krust, P. Chambon, Nature 330 (1987) 444–450. [21] D.J. Mangelsdorf, U. Borgmeyer, R.A. Heyman, J.Y. Zhou, E.S. Ong, A.E. Oro, A. Kakizuka, R.M. Evans, Genes Dev. 6 (1992) 329–344. [22] M.G. Motto, K.I. Facchine, P.F. Hamburg, D.J. Burinsky, R. Dunphy, A.R. Oyler, M.L. Cotter, J. Chromatogr. 481 (1989) 255–262. [23] J.J. Repa, J.A. Berg, M.E. Kaiser, K.K. Hanson, S.A. Strugnell, M. Clagett-Dame, Prot. Expr. Purif. 9 (1997) 319–330. [24] M. Munder, I.M. Herzberg, C. Zierold, V.E. Moss, K. Hanson, M. Clagett-Dame, H.F. DeLuca, Proc. Natl. Acad. Sci. USA 92 (1995) 2795–2799. [25] R.A. Merrill, L.A. Plum, M.E. Kaiser, M. Clagett-Dame, Proc. Natl. Acad. Sci. USA 99 (2002) 3422–3427. [26] T. Sasaki, S. Iwata, H.J. Okano, Y. Urasaki, J. Hamada, H. Tanaka, N.H. Dang, H. Okano, C. Morimoto, Stroke 36 (2005) 2457–2462. [27] H. de Thé, M.M. Vivanco-Ruiz, P. Tiollais, H. Stunnenberg, A. Dejean, Nature 343 (1990) 177–180. [28] D.J. Mangelsdorf, Nutr. Rev. 52 (1994) S32–44. [29] M. Clagett-Dame, L.A. Plum, Crit. Rev. Eukaryot. Gene Expr. 7 (1997) 299–342. [30] G. Vasios, S. Mader, J.D. Gold, M. Leid, Y. Lutz, M.-P. Gaub, P. Chambon, L. Gudas, EMBO J. 10 (1991) 1149–1158. [31] G.W. Vasios, J.D. Gold, M. Petkovich, P. Chambon, L.J. Gudas, Proc. Natl. Acad. Sci. USA 86 (1989) 9099–9103. [32] A. Åström, U. Pettersson, P. Chambon, J.J. Voorhees, J. Biol. Chem. 269 (1994) 22334–22339. [33] L. Panariello, L. Quadro, S. Trematerra, V. Colantuoni, J. Biol. Chem. 271 (1996) 25524–25532. [34] Y.-S. Piao, H. Peltoketo, J. Oikarinen, R. Vihko, Mol. Endocrinol. 9 (1995) 1633– 1644. [35] A. Naltner, M. Ghaffari, J.A. Whitsett, C. Yan, J. Biol. Chem. 275 (2000) 56–62. [36] S. Kato, H. Sasaki, M. Suzawa, S. Masushige, L. Tora, P. Chambon, H. Gronemeyer, Mol. Cell. Biol. 15 (1995) 5858–5867. [37] J.E. Balmer, R. Blomhoff, J. Lipid Res. 43 (2002) 1773–1808. [38] J. Wang, A. Yen, Mol. Cell. Biol. 24 (2004) 2423–2443. [39] L.J. Donato, J.H. Suh, N. Noy, Cancer Res. 67 (2007) 609–615. [40] K. Han, H. Song, I. Moon, R. Augustin, K. Moley, M. Rogers, H. Lim, J. Endocrinol. 192 (2007) 539–551. [41] N.G. Pedigo, H. Zhang, A. Mishra, J.R. McCorkle, A.K. Ormerod, D.M. Kaetzel, Gene Express. 14 (2007) 1–12. [42] O. Loudig, C. Babichuk, J. White, S. Abu-Abed, C. Mueller, M. Petkovich, Mol. Endocrinol. 14 (2000) 1483–1497. [43] O. Loudig, G.A. MacLean, N.L. Dore, L. Luu, M. Petkovich, Biochem. J. 392 (2005) 241–248. [44] C. Li, J. Locker, Y.-J.Y. Wan, DNA Cell Biol. 15 (1996) 955–963. [45] L. Nagy, M. Saydak, N. Shipley, S. Lu, J.P. Basilion, Z.H. Yan, P. Syka, R.A.S. Chandraratna, J.P. Stein, R.A. Heyman, P.J.A. Davies, J. Biol. Chem. 271 (1996) 4355–4365. [46] L.E. Tavera-Mendoza, S. Mader, J.H. White, Nucl. Recept. Signal. 4 (2006) 1–8. [47] G. Deblois, V. Giguère, Mol. Endo. (2008). [48] J. Bastien, C. Rochette-Egly, Gene 328 (2004) 1–16.

174

D.C. Knutson, M. Clagett-Dame / Archives of Biochemistry and Biophysics 477 (2008) 163–174

[49] H. Umemiya, H. Kagechika, H. Fukasawa, E. Kawachi, M. Ebisawa, Y. Hashimoto, G. Eisenmann, C. Erb, A. Pornon, P. Chambon, H. Gronemeyer, K. Shudo, Biochem. Biophys. Res. Commun. 233 (1997) 121–125. [50] J.-Y. Chen, J. Clifford, C. Zusi, J. Starrett, D. Tortolani, J. Ostrowski, P.R. Reczek, P. Chambon, H. Gronemeyer, Nature 382 (1996) 819–822. [51] B. Roy, R. Taneja, P. Chambon, Mol. Cell. Biol. 15 (1995) 6481–6487. [52] R. Schüle, K. Umesono, D.J. Mangelsdorf, J. Bolado, J.W. Pike, R.M. Evans, Cell 61 (1990) 497–504. [53] B.D. Raisher, T. Gulick, Z. Zhang, A.W. Strauss, D.D. Moore, D.P. Kelly, J. Biol. Chem. 267 (1992) 20264–20269. [54] Q. Tao, Y. Cheng, J. Clifford, R. Lotan, Genomics 83 (2004) 270–280.

[55] D.K. Scott, J.A. Mitchell, D.K. Granner, J. Biol. Chem. 271 (1996) 31909–31914. [56] P.M. Barger, D.P. Kelly, J. Biol. Chem. 272 (1997) 2722–2728. [57] T.R. Magee, Y. Cai, M.E. El-Houseini, J. Locker, Y.-J.Y. Wan, J. Biol. Chem. 273 (1998) 30024–30032. [58] H. Nakshatri, P. Chambon, J. Biol. Chem. 269 (1994) 890–902. [59] S.N. Lavrentiadou, M. Hadzopoulou-Cladaras, D. Kardassis, V.I. Zannis, Biochemistry 38 (1999) 964–975. [60] S. Påhlman, A.-I. Ruusala, L. Abrahamsson, M.E.K. Mattsson, T. Esscher, Cell Differ. 14 (1984) 135–144. [61] S.D. Bargon, P.W. Gunning, G.M. O’Neill, Biochim. Biophys. Acta 1746 (2005) 143–154.