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Mutant p53 gain of function induces HER2 over-expression in cancer cells

Background HER2 over-expression is related with a poor prognosis in patients with invasive breast cancer tumors. Clinical associations have reported that somatic mutations of p53 more frequently detected in cases of sporadic breast cancer of the HER2 subtypes, besides a high percentage of HER2-amplifying tumors carry germline mutations of p53. The mechanisms responsible for the acquisition of oncogenic functions of p53 mutant proteins (mtp53), known as Gain of Function (GOF), over HER2 expression have not been reported. The objective of this study was to evaluate a possible relationship between p53 mutants and HER2 regulation. Methods HER2 expression (transcription and protein), as well as HER2 protein stabilization have been evaluated after inducing or silencing of p53 mutants’ expression in cell lines. Finally, we evaluated the interaction of the p53 mutants over the HER2 receptor promoter. Results Higher HER2 expression in cell lines harboring endogenous mtp53 compared with wt or null expression of p53 cell lines. Transfection of p53 mutants (R248Q and R273C) in cell lines increased the expression of HER2. Silencing of p53 mutants, decrease HER2 expression. The p53 mutants R248Q and R273C significantly increase the luciferase activity on the HER2 promoter, and both mutants also promote acetylation of H3 and H4 histones binding in it. Conclusions These findings show for the first time that p53 mutants induce over-expression of HER2 at transcriptional level of the HER2 protein. Our results could have clinical implications in breast cancer and other types of cancer where HER2 is over-expressed and used as a therapy target. Electronic supplementary material The online version of this article (10.1186/s12885-018-4613-1) contains supplementary material, which is available to authorized users.

Background
The p53 tumor suppressor protein, which is encoded by TP53 gene, exerts its biological functions mainly by its transcriptional activity, although it is accepted that wild-type p53 (wtp53) has other biological activities that are transcription independent [1,2]. Wild-type p53 levels are very low in normal cells; however, they rise rapidly in response to DNA damage, hypoxia, oxidant metabolism or oncogenic signaling, as well as in response to aging, maintaining genomic integrity and preventing tumor formation [3,4]. TP53 is the most frequent target for mutations in human cancers, with more than half of all tumors exhibiting mutation at this locus. Unlike other tumor-suppressor genes, which typically are mutated by deletion or truncation, TP53 frequently undergoes missense mutations, resulting in single amino acid substitutions in the full length protein [3]. About a third of these missense mutations are located in six residues: R175, G245, R248, R249, R273, and R282, corresponding to the p53 DNA binding domain (DBD) and known as mutational hot-spots [5]. These p53 mutations can be roughly divided into two structural subgroups: DNA contact mutants (exemplified by R273 and R248 residues) directly involved in sequence-specific DNA contact; and conformational mutants (exemplified by the R175 residue), leading to a partial or complete abrogation of the conformational wtp53 DBD. The presence of these point mutations radically alters p53 function causing not only a simple loss of wild-type function, but also a dominant negative effect by binding and inhibiting wtp53 or a Gain of Function (GOF) acquiring novel activities independent of wtp53 [3]. p53 GOF mutations have been shown to result in oncogenic and a major proliferative processes such as increased tumorigenicity, anchorage independent cell growth and increased growth rate, increased metastasis and invasiveness, decreased sensitivity to chemotherapeutic drugs, disruption of the spindle checkpoint, activated topoisomerase I activity and induction of gene amplification, reviewed in [3]. Many of the GOF data come from p53-null systems where the expression levels of re-expressed mutant p53 were comparable to those observed in cancer cells. These results suggest a true patho-physiological role of the GOF of p53-mutants, which may lead to the development of a more aggressive cancer and poorer prognosis. The molecular mechanism of GOF's phenotypes and up-regulation of gene expression by p53 mutants has not been clarified yet [6]. Among other important biomarkers implicated in several human cancers, there are the Human Epidermal growth factor Receptors (HERs), which regulate cell proliferation, angiogenesis, cell adhesion, cell motility, development and organogenesis, by activation of different downstream signaling pathways [7]. The HER family consists of four members (HER1-4) expressed in epithelial, mesenchymal, and neuronal cells, as well as in their cellular progenitors. Like all protein-tyrosine kinase receptors, the HER receptors exist as monomers on the cell surface and depend on their specific ligands for dimerization and trans-phosphorylation of their intracellular domains [8]. Although the product of the ERBB2 gene (HER2) has no known direct activating ligand due to its constitutively activated state, this receptor is known as the preferred dimerization partner for the other family members [9]. HER2 over-expression or gene amplification occurs in 20-30% of breast cancers with a correlation of poor prognosis prior to the advent of anti-HER2 therapies. Anti-HER2 therapy is currently approved for breast, gastric, and gastroesophageal cancers, when this receptor is over-expressed. However, HER2 over-expression have been reported in other malignancies, such as bladder, cervix, colon, endometrium, germ cell, glioblastoma, head and neck, liver, lung, ovarian, pancreas and salivary duct [10]. Elucidating the mechanism leading to HER2 gene up-regulation will be an important step to understand the pathogenesis of particularly aggressive subset of tumors over-expressing HER2 [11], as well as to find novel alternatives for therapy. Interestingly, Wilson et al. reported that patients carrying TP53 mutations (mtp53) show a significantly higher probability of developing breast cancer with HER2 gene amplification (83%) when compared to the cohort of early onset breast cancer cases (16%) [12]. A larger study that supports this association, found that patients with breast cancer harboring a germline TP53 mutation, have significantly higher HER2 positive tumors prevalence, compared to their counterparts who lack any mutation [13]. There are also studies where both somatic TP53 mutations and HER2 gene amplification are correlated with elevated risk of breast cancer recurrence and elevated overall mortality compared with patients with neither or only one alteration [14]. It is important to note that the studies mentioned above are just clinical associations and do not propose any mechanism responsible for this relation. In this work we found, for the first time, that p53 mutants can induce up-regulation of HER2 gene and favor the stabilization of the protein. Understanding the specific mechanisms for this relationship might reveal diagnostic and therapeutic insights in HER2 positive cancers.

Cell culture
Cancer cell lines: Saos-2, MCF-7, HeLa, C33A, OVCAR-3 and SKBR-3 were originally obtained from the American Type Culture Collection (ATCC). These cell lines were cultured in DMEM supplemented medium at 37°C in a humidified 5% CO 2 environment and were grown at 50 to 80% confluence before the next passage or further experiments.

Transient and stable transfections
Cells were plated (5 X 10 5 cells) in 60 mm × 15 mm culture dishes and transfected with: empty vector, p53R248Q, p53R273C [15], and wtp53 [16] plasmids; or with SureSilencing shRNA Plasmid Kit for TP53 (SABiosciences, Qiagen). Transfections were performed with Lipofectamine 2000 reagent (Invitrogen, CA) according to the manufacturer's instructions. Transient transfections were performed for 48 h. Stable transfections were selected for 2 weeks in a growth media containing 1200 μg/ml of G-418 (Sigma-Aldrich, St Louis MO). To keep the clones selection, cells were grown continuously in media containing 800 μg/ml of G418.

Reverse transcription and quantitative polymerase chain reaction (RT-qPCR)
Total RNA was obtained using TRIZOL reagent (Invitrogen, CA) and cDNA was reverse transcribed from 3 μg of RNA using SuperScript II reverse transcriptase (Invitrogen, CA), following the manufacturer's recommendations. Oligonucleotide primers were designed using the Primer Express Software and purchased from Sigma-Aldrich (St Louis MO). qPCR assays were performed by using SYBR green Master Mix (Thermo Scientific, Waltham MA) in an ABI 7300 Real Time PCR system (Applied Biosystems, California USA). The qPCR program was set as follow: 95°C for 5 min, followed by 40 cycles at 95°C for 30 s, 60°C for 30 s, 72°C for 30 s and a final elongation step of 72°C for 7 min, with a final dissociation curve. Each reaction was carried out in triplicate and targeted genes were normalized to beta-2-microglobulin (B2M) gene. The threshold value (CT) for each sample was used to determine gene relative expression levels by the comparative CT method (2 -ΔΔCT method) [17]. Detailed description of the primers used for HER2 and TP53 detection can be found in the Additional file 1: Table S1.

Western blot analysis
Equal amounts of whole-cell protein extracts were separated by size on discontinuous (8 and 10%) PAGE, and transferred to a 0.45 μm nitrocellulose membrane (Millipore, Billerica MA). After blocking, membranes were probed overnight with: HER2 monoclonal primary antibody (Cell signaling, Danvers, MA) and peroxidase-conjugated antibodies: p53-DO1 (Santa Cruz Biotechnology) and β-actin (Sigma-Aldrich, St Louis MO). Secondary rabbit antibody (Sigma-Aldrich, St Louis MO) required for HER2 was employed at 1:10000 dilution. Protein expression was detected by chemiluminiscence using Supersignal West Pico (Thermo Scientific, Waltham MA). The protein band intensity was analyzed from the Western blot images by using the Syngene software.

Promoter luciferase reporter assays
Stable transfected Saos-2 cells (empty vector, wtp53, p53R248Q, and p53R273C) were seeded at 1 × 10 5 cells in six-wells plates and then transiently transfected with HER2 promoter vector, pNeuLite (Addgene Inc., Cambridge MA), using Lipofectamine 2000 reagent (Invitrogen, CA) as described above. After 24 h, cells were lysed and samples probed with Dual-Glo®Luciferase Assay System (Promega, Madison WI) essentially as described in [18]. Assays were read with the Fluoroskan Ascent FL (Labsystems, Perú), 0.5 s integration/ well. Expression induction was calculated as the relative light units obtained for the Saos-2 cells expressing mtp53/average relative light units of the Saos-2 cells expressing empty vector, normalized with renilla relative light units. Results were plotted against the log of the compound concentration, using Prism 6 Software (GraphPad, San Diego CA).

ChIP promoter assay
Chromatin immunoprecipitation (ChIP) assay over HER2 promoter was performed in accordance to manufacturer's kit protocol (ChIP One Day Kit, Qiagen). Briefly, cells were fixed at room temperature and washed with PBS. To obtain the cell insoluble material, cells were gently scraped with lysis buffer, collected and pelleted by centrifugation at 4°C. DNA-protein complexes were sonicated to produce fragmented chromatin between 200 and 600 bp in length, determined by agarose gel electrophoresis. One-tenth of the fragmented chromatin was set aside as input control, and the remaining sample was pre-cleared with protein A/G. For each ChIP, either anti-p53 (DO-1) (Santa Cruz Biotechnology), anti-Histone 3 (Upstate, Millipore Merck) or anti-Histone 4 antibodies (Upstate, Millipore Merck), or non-specific control (IgG) was added and incubated overnight at 4°C. Immune complexes were precipitated at 4°C with magnetic beads and DNA-protein cross-links were reversed by adding NaCl. Samples were subsequently treated with proteinase K, and genomic DNA was recovered and purified by column affinity. Quantitation of DNA from p53, H3ac and H4ac ChIP, no antibody control, and input control was done with RT-qPCR as described above. Primers were designed by Primer Express Software (Applied Biosystems, Foster City, CA) and sequences are shown in Additional file 2: Table S2. Comparisons were normalized to input controls.

Statistical analysis
Each experiment was performed at least three times, the results were presented as mean ± Standard Error of the Mean (SEM). Statistical analysis was performed with 1-tailed paired Student's t test for means between controls and experimental data. Histograms and statistical analysis were carried out using Prism (Graphpad). * p < 0.05, ns = non-significant statistical difference analysis.

HER2 and p53 expression levels in cancer cell lines
As a first approach to the correlation that may exist between p53 mutant proteins and the HER2 receptor expression, we evaluated the mRNA and protein levels of p53 and HER2 in a panel of human cancer cells: MCF-7 and HeLa cells harboring wild-type p53 (wtp53), Saos-2 with null expression of p53 (p53-null), as well as, C33A, SKBR-3 and OVCAR-3 harboring three of the most frequent p53 hot-spot mutations (p53R273C, p53R175H and p53R248Q). For HER2 mRNA expression, quantitative PCR (RT-qPCR) assays revealed that the cell lines harboring mutant p53 (mtp53) had higher expression levels when compared with the cell lines with wild-type or null p53 expression (Fig. 1a). In agreement with the mRNA expression of HER2, we observed a higher HER2 protein expression in the cell lines harboring mutant p53 (Fig. 1b). In Additional file 3: Figure S1, (A) we showed that exogenous wtp53 expression in HeLa cell line, counteracting HPV-E6 effects, decrease HER2 mRNA expression; moreover, (B) wtp53 silencing in MCF-7 cell line increase HER2 mRNA expression; (C and D) both mRNA and protein HER2 levels are insignificantly decreased in Saos-2 cells transfected with wtp53.

The p53R248Q and p53R273C mutants increase HER2 expression
We then evaluated the effect of silencing or expressing exogenously two specific mutants (p53R248Q and p53R273C), on the expression of HER2. First, we silenced p53 expression in OVCAR-3 cell line which endogenously express p53R248Q mutant and observed a decreased expression of both the HER2 mRNA and protein, when compared to the shRNA negative control plasmid ( Fig. 2a and b). On the contrary, when we expressed exogenously the mutant p53R248Q by stably transfection in the Saos-2 cell line (p53-null) ( Fig. 2c and d), we observed that the p53R248Q mutant, lead to an elevated mRNA and protein expression of HER2. We also transfected transiently p53R248Q mutant in the HeLa cell line with similar results than in Saos-2 cell line (Data not shown). Next, also based on our previous results where we observed an elevated expression of HER2 in C33A, a cervical cancer cell line harboring the p53R273C mutant; we silenced the endogenous expression of the p53R273C mutant in C33A cell line, and this silencing lead to a decreased in the expression in both mRNA and protein levels of HER2 as well as in the case of p53R248Q mutant ( Fig.  2e and f). In counterpart, stably transfection in the p53-null cell line (Saos-2) ( Fig. 2g and h), lead to an increased HER2 protein expression. We also transiently transfected the p53R273C mutant, in the HeLa cell line (data not shown), in this case we also found an increase of HER2 protein level between HeLa and Saos-2 cells, but in HeLa we did not observe an increase of HER2 mRNA expression. These results suggest that the p53R248Q mutant, regulates positively HER2 expression at both transcriptional and protein level while regulation by p53R273C mutant could be cell context dependent.
The p53R248Q and p53R273C mutants induce transcriptional activation of HER2 promoter in Saos-2 cell line Because we also observed that p53R248Q and p53R273C mutants induce an increase in the HER2 mRNA expression, we evaluated the effect of both mutants over the HER2 , as well as cells containing wild-type p53 (wtp53) protein (MCF-7 and HeLa), and null p53 expression (Saos-2) were analyzed to determine HER2 mRNA and protein expression. HeLa cell line expresses wtp53 but its expression is decreased by Human Papillomavirus 18 (wtp53 • ). a HER2 relative mRNA expression was measured by quantitative RT-PCR (RT-qPCR); final data were calculated by the comparative threshold cycle (CT) method (2 -ΔΔCT ), employing the β2-microglobulin (β2M) transcript as normalization control. The results are shown as the mean ± the standard error of the mean (SEM) of three independent assays. Statistical analysis was performed by comparing each cell line against Saos-2 cell line with t-student test, *p < 0.05; ns = non-significant statistical analysis. b HER2 protein levels were measured by Western Blot assay. Equal amounts of total cell lysates were blotted and proteins were identified as described in the Materials and Methods, with specific antibodies against HER2 and p53. Blots were stripped and re-probed for anti-β-actin as loading control. Image is representative of three independent experiments for: e HER2 mRNA expression assay by RT-qPCR, and f protein expression assay by Western Blot. g-h The Saos-2 cell line (null-p53) was stably transfected with mutant p53R273C, or with empty vector, as indicated in Materials and Methods. Transfected cells were processed after stable selection for: g HER2 mRNA expression assay by RT-qPCR, and h protein expression assay by Western Blot. The RT-qPCR results are the means of at least three independent experiments ± SEM and the image of the blot is representative of three independent experiments. Statistical analysis was performed with t-student test by comparing the results obtained in: shp53 vs scramble transfected C33A cells; or p53R273C vs empty vector transfected Saos-2 cells: * p < 0.05 promoter using a luciferase transactivation assay in the HER2 promoter. We used the Saos-2 cells expressing stably either the p53R248Q or p53R273C mutants and we co-transfected the plasmid containing the HER2 promoter sequence in a luciferase reporter vector (pNeuLite) [19]. The luciferase activity was normalized with data obtained for renilla activity (Fig. 3). These results showed that, under our experimental conditions, p53R248Q and p53R273C significantly increased up to 50% the HER2 promoter activity, with respect to the control (empty vector). We also evaluated the transient ectopic expression of wtp53 in Saos-2 cell line and in this case, the results showed that exogenous wtp53 significantly diminished the HER2 promoter activity, in contrast to the results observed with the mutants of p53 analyzed (Fig. 3).
The p53R248Q and p53R273C mutants bind and induce histone acetylation on the HER2 promoter Due to our previous results indicating that p53 mutants up-regulate HER2 promoter activity, we decided to gain additional insight into the molecular mechanisms underlying a possible physical interaction between the HER2 promoter and the p53R248Q or p53R273C mutants. We performed chromatin immunoprecipitation (ChIP) assays, using antibodies directed against p53 and against acetylated forms of histones H3 and H4. These assays were performed with chromatin extracted from the Saos-2 cells stably expressing p53R248Q or p53R273C, as well as with the empty vector (control). We designed specific primers to dissect the HER2 proximal promoter region into three PCR products, denominated as HER2a, HER2b and HER2c, according to the description reviewed by Hurst (Fig. 4a). We found that in HER2a and HER2b regions, both p53R248Q and p53R273C mutants had a similar fold enrichment after p53 immunoprecipitation: HER2a (2.3 and 3.0 fold enrichment, respectively) and HER2b (2.4 and 2.5, respectively); moreover, in both regions there is an important increase in the acetylation marks of H3 and H4 histones ( Fig. 4b and c). For HER2c we found that p53R248Q mutant has the highest enrichment, while p53R273C mutant had a similar enrichment both HER2a and HER2c (10.4 and 2.2 fold enrichment, respectively); the most important enrichment in acetylated H3 and H4 marks was also observed in this region (Fig. 4d). The Saos-2 cell line transfected with p53R248Q presented both the highest p53 protein level and acetylated histones (particularly H4) in the three proximal regions. For the HER2 distal promoter, we used the same set of primers reported by Delacroix et al., 2005 [22]. These primers were denominated HER2d and HER2e, which respectively cover two AP-2 binding sites located in the − 4000 bp and − 4500 bp of the HER2 promoter. Another region upstream of the HER2 promoter, with no HER2 activator binding sites, denominated as HER2f was included (Fig. 5a). We observed that HER2d has the major fold enrichment for the p53R273C mutant in the HER2 promoter. In general, there was a modest enrichment of acetylated histone marks for HER2d and HER2e (Fig. 5b and c). The region denominated HER2f had the lowest enrichment for acetylation marks in H3 and H4 histones, coinciding with the lowest p53 recruitment in the promoter (Fig. 5d). It is worth noticing that in distal regions (HER2d and HER2e) the enrichment of acetylated H3 and H4 have differences between the two evaluated mutants; while p53R273C induced a higher increase in the enrichment of acetylated histone marks for HER2d region, the p53R248Q favor a higher increase of these marks in the HER2e region. Taken together, these results suggest that both the p53R248Q and p53R273C mutants are directly involved in HER2 promoter activation. Moreover, both p53 mutants are contributing to the modification of the HER2 promoter towards a Transcriptional activity of the HER2 promoter was assessed by a Luciferase reporter assay in Saos-2 cells stably transfected with R248Q and R273C p53 mutants, as well as with wild-type p53 plasmid and empty vector. Stably transfected Saos-2 cells were co-transfected with pNeuLite luciferase reporter plasmid for the HER2 promoter. After 48 h, luciferase activity was measured as described in the Materials and Methods. The results of the luciferase activity were normalized to renilla activity and are reported as the mean ± SEM of three independent experiments. Statistical analysis was performed with t-student test by comparing the results obtained for the Relative Activity in Saos-2 cells transfected with p53R48Q or p53R273C or wtp53 against empty vector: * p < 0.05 more activated chromatin by enriching H3 and H4 acetylation marks.

Discussion
It is generally accepted that mutant p53 (mtp53) proteins promote carcinogenesis by exerting a dominant-negative effect over wtp53 protein, as well as by new oncogenic gain-of-function (GOF) activities independent of wtp53. However, the mechanisms of the GOF phenotype by p53 mutants, are only starting to be defined [23]. In previous works, it has been observed that p53 mutants are more frequently detected in a subset of breast and gastric cancer patients with poor prognosis, such as in those presenting the HER2 subtype [24,25]. We choose the cell lines mentioned in the Material and Methods section, because Saos-2 is a cell line known for be p53-null, which is common used to study new roles for the wild-type and mutants of p53 [26]. In the case of MCF-7 and SKBR-3 breast cancer cell lines, Xiao et al., observed that MCF-7 cells have a typical number of gene copies (i.e., 2) and very low HER2 protein content, whereas SKBR-3 cells contain approximately 10-fold more copies of the HER2 gene and 15-fold higher HER2 protein content [27]. Consistent with Xiao's results, we observed the most expression of mRNA and protein expression in SKBR-3 cell line. We decided not to use SKBR-3 cell line or p53R175H mutant in Fig. 4 The p53R248Q and p53R273C mutants associate with chromatin and induce histone H3 and H4 acetylation in HER2 promoter proximal region. Saos-2 cells stably transfected with p53R248Q, p53R273C or empty vector were analyzed by Chromatin immunoprecipitation (ChIP) as described in the Materials and Methods. Antibodies against acetylated histone H3, H4, p53 protein and IgG as non-relevant antibody were used for the immunoprecipitation. a Shows a graphic representation of the regions encompassed by the primers used for ChIP assay. Precipitated DNA was amplified with primers described in Additional file 2: Table S2. HER2a (b) region from − 22 to − 75 bp; HER2b (c) -217 bp, and (d) region with not known HER2 regulatory sequences. Relative amounts of acetylated H3 and H4 histones, as well as p53 interaction with HER2 promoter regions were determined on the precipitated DNA by quantitative PCR (qPCR). Normalized data were calculated relative to each primer set (b-d), comparing p52R248Q or p53R273C vs IgG ChIP 1.0% input sample. The results of Fold Enrichment are reported as the mean ± SEM of three independent experiments. Statistical analysis was performed with t-student test by comparing the results obtained for the Fold Enrichment in Saos-2 cells transfected with p53R48Q or p53R273C against empty vector transfected cells: * p < 0.05 subsequent experiments indeed the amplification of HER2 in this cell line [28]. In case of cervical adenocarcinomas, there are reports that conclude that amplification of HER2 and mutations in TP53 are rare in cervical adenocarcinomas and that, low level chromosome 17q copy number gains are not associated with HER2 over-expression [29]. Beaufort et al., described several ovarian cancer cell lines and confirms the p53 mutation for the OVCAR-3 cell line but no HER2 amplification reported [30]. In the present study, we provided evidence to support the notion that mtp53 is implicated in the positive regulation of HER2 expression. We showed that in two different cell lines harboring wtp53 (HeLa and MCF-7), HER2 mRNA is expressed in very low level, which agrees with the results of Yang et al. (2006) indicating that wtp53 could negatively regulate HER2 [31]. In this regard, HeLa is a cervical cancer cell line containing the human papillomavirus-18 (HPV-18) genome that encodes for the E6 oncoprotein and causes p53 degradation [32]; which might explain the HER2 protein expression in HeLa cells. On the other hand, in spite that MCF-7 also harbors wtp53, the high basal HER2 protein levels observed in this cell line is probably inherent to the normal genetic background needed to maintain the function of breast cells; Fig. 5 The p53R248Q and p53R273C mutants increase histone acetylation and p53 bind to HER2 promoter distal region. Chromatin immunoprecipitation (ChIP) assays were carried out in p53R248Q, p53R273C and empty vector stably transfected Saos-2 cells as described in the Material and Methods. Antibodies against acetylated histone H3, H4, p53 protein and IgG as non-relevant antibody were used for the immunoprecipitation. a Shows a graphic representation of the regions encompassed by the primers used for ChIP assay. Precipitated DNA was amplified with primers described in Additional file 2: Table S2. HER2d (b) -4000 bp; HER2e (c) -4500 bp, and HER2f (d) up-steam the HER2 promoter. Relative amounts of acetylated H3 and H4 histones, as well as p53 interaction with HER2 distal promoter regions were determined by quantitative PCR (qPCR). Normalized data were calculated relative to each primer set (b-d), comparing p52R248Q or p53R273C vs IgG ChIP 1.0% input sample. The results of Fold Enrichment are reported as the mean ± SEM of three independent experiments. Statistical analysis was performed with t-student test by comparing the results obtained for the Fold Enrichment in Saos-2 cells transfected with p53R48Q or p53R273C against empty vector transfected cells: *p < 0.05 additionally, the p53 pathway has been suggested to be impaired in this tissue [33,34]. In our work, we found that cell lines harboring three of the most common p53 mutants (p53R273C, p53R175H and p53R248Q) express higher levels of HER2 (mRNA and protein) compared to the cell lines with functional wtp53 (MCF-7), HPV-inactivated wtp53 (HeLa) or null-p53 (Saos-2). Our results are consistent with studies that suggest a clinical association between the presence of p53 mutations and HER2 over-expression [12][13][14]. We demonstrated that the exogenous expression of the p53R248Q or p53R273C mutants in two different cell types, HeLa and Saos-2 induces a significant increase in the levels of HER2 protein. On counterpart, we analyzed the effect of inhibiting the expression of the endogenous p53R248Q or p53R273C mutants harbored in the cell lines OVCAR-3 and C33A, respectively. We observed differences in the action efficiency of the shRNA for p53 silencing, which may be due the efficiency of transfection intrinsic to each cell line. The most p53 expression inhibition was for the p53R273C mutant expression compared with the expression for the p53R248Q mutant. Despite this, in both cases the inhibition of the p53 mutants' expression was enough to decrease the expression of HER2, both at the level of mRNA and protein. The regulation of HER2 by p53 mutants, suggests important clinical implications in the diagnostic and therapy of HER2 positive breast cancer. To gain some insight on the molecular mechanisms by which p53 mutants positively regulate the expression of HER2, we evaluated the possibility of induction of HER2 promoter activity. In this sense, previous reports have demonstrated the promoter activation of genes such as MDR1, EGFR, NF-kB and Axl by mtp53 activity [6,35]. Our results showed that both p53 mutants (R248Q and R273C) stably expressed in the Saos-2 cell line induced HER2 gene promoter activity employing a luciferase reporter assay. Up-regulation of the HER2 promoter activity could be the result of direct mutant p53 DNA-binding, induction or recruitment of other transcription factors and transcriptional activators such as histone acetyl transferases. Among known HER2 promoter activators are the transcription factors AP-2, YY1 and members of the ETS family [21,36,37]; there are also repressors such as FOXP3, PEA3 and GATA4 [19,38]. According with the HER2 promoter described by Hurst in 2001, we evaluated if the observed up-regulation in HER2 is achieved through binding of mutant p53 to DNA fragments containing AP-2 or ETS sequences [21]. We demonstrated the specific recruitment of p53 mutants to both, proximal and distal regions of the promoter by ChIP assay but not all the regions were enriched at the same level. Interestingly, we observed that the highest enrichment for the p53R248Q mutant was in the HER2 proximal region while the mutant p53R273C shows its highest recruitment level in the distal HER2 promoter region even though both protein enrichments were located at regions with AP-2 binding sites. It is possible that the p53 mutants alone or bound with AP-2 proteins have different affinity or bind selectively to different sequences near AP-2 binding sites in the HER2 promoter. AP-2 factors have been demonstrated to bind to wtp53 in vitro and in vivo [39,40]; for example, in a p53-dependent binding AP-2 can regulate the p21WAF1/CIP1 gene promoter [39,41].
Recently it was also determined, using p53 point mutants, that the AP-2 binding region of p53 is located in amino acids 305-375 [42]. It is worth mentioning that the p53R248Q and p53R273C mutants are capable of activating AP-2 binding regions in other promoters, suggesting that these p53 mutants could cooperate with AP-2 to activate HER2 transcription. Other transcription factors have been shown to interact with p53 mutants, including SP1, ETS1, ETS2, NF-Y, and the Vitamin D receptor [43], altering their target genes expression. Mutant p53R273H might exert its oncogenic gain-of-function by binding to ETS human genomic sequences, both within proximal and distal promoters [44]. In agreement with these studies, in the HER2 proximal promoter there is an ETS sequence, and we observed that p53R248Q and p53R273C mutants bind to this region, suggesting that these p53 mutants interact with ETS transcription factors. On the other hand, we evaluated if these p53 mutants had any repercussion on H3 and H4 histone acetylation, enhancing the epigenetic marks and inducing directly or indirectly HER2 promoter activity [45,46]. As expected, histone H3 and H4 acetylation enrichment induced by p53 mutants coincides with the level of HER2 promoter activation observed in the luciferase reporter assays. In general, the major increase in the level of the acetylated marks in H3 and particularly in H4 histones was due to p53R248Q mutant, and observed in the HER2 proximal promoter, indicating a more relevant participation of the proximal promoter for the HER2 activation. In agreement with our results, Zhu et al. showed that p53 GOF mutants bind to and up-regulate chromatin regulatory genes, such as the methyltransferases MLL1, MLL2 and acetyltransferase MOZ, resulting in genome-wide increases of histone methylation and acetylation. Analysis of The Cancer Genome Atlas shows a specific tendency of up-regulation of MLL1, MLL2 and MOZ in selected patient tumors that express GOF p53-mutants [47]. The MOZ gene, encodes an enzyme that adds an acetyl group to K9 of H3 histone, allowing increased gene expression [47]. In agreement with this report, we observed that p53 mutants, R248Q and R273C, increase the level of acetylated marks in H3 histone at the HER2 promoter. In addition, we observed that the acetylation level of histone H4 was higher than that for histone H3 in all the regions analyzed, coinciding with Sandip K et al., 2001 observation, using HDAC inhibitors, suggesting that H4 acetylation is more important for HER2 activation [46]. A schematic representation for the histone arrangement and p53 binding in the proximal and distal HER2 promoter regions is shown in Fig. 6.
On the other hand, mutant p53 also could increase the stability of HER2 protein, previous reports had demonstrated that p53R175H can regulate some genes like EGFR at the transcriptional level, and EGFR contributes to the HER2 protein stability by increasing its dimerization state [7,28]. In addition, it has been previously reported that HER2 protein stabilization is also induced by Hsp90 [48]. The chaperone system controls the stability of the nascent forms of both EGF-R and HER2, but only the mature form of HER2. Interestingly, it has been also reported that Hsp90 also plays a key role in the folding of p53 mutants and stabilization, suggesting that a target therapy to Hsp90 could inhibit at the same time the HER2 signaling pathway and mutant p53 gain of function [49]. Recently, it was also demonstrated that mutants of p53 promote HSF1 phosphoactivation, protein stabilization and specific DNA-binding of this protein to its target gene promoters; HSF1 is the master transcriptional regulator of heat shock chaperones, including HSP90, suggesting a mtp53-HSF1-HSP90 oncogenic cooperation to stabilize the HER2 protein [50].

Conclusion
In conclusion, our study has demonstrated for the first time that HER2 over-expression is induced by p53 mutants through HER2 transcriptional activation. Acetylated H3 and H4 histones are associated to the HER2 proximal promoter in the presence of both p53R248Q and p53R273C mutants. However, more studies are required to completely elucidate the mechanism by which p53 mutants can induce HER2 mRNA up-regulation and its protein over-expression. Exploring the oncogenic gain of functions of p53 mutants may reveal new opportunities for diagnosis and therapy. In this regard, our results could have clinical implications in breast cancer and other cancer types where HER2 is over-expressed and used as a therapy target. As we mentioned in background section, Wilson et al. reported that patients carrying TP53 mutations show a significantly higher probability of developing breast cancer with ERBB2 gene amplification in human tumors. It will be necessary to demonstrate if the p53R248Q and p53R273C mutants are associated to HER2 over-expression in cancer or if there are specific mutations of p53 associated to overexpression of HER2. Fig. 6 Model of HER2 up-regulation by p53 mutants. Three different molecular mechanisms of how mutant p53 may activate transcription from HER2 promoter are depicted. a Mutant p53 may induce histone (s) acetylation and/or inhibit histone deacetylase (s) causing enrichment of H3 and H4 acetylated forms and chromatin reorganization on the HER2 promoter; b Mutant p53 may induce interaction of one or more transcription factor (s), with the HER2 promoter, such as AP-2 and ETS; c Mutant p53 itself can interact directly on the HER2 promoter. These 3 possibilities are not mutually exclusive and would lead to the HER2 promoter activation with the consequent HER2 gene expression. In this representation, the histones are large blue spheres, the histone H3 acetylated marks are small green circles, the histone H4 acetylated marks are small yellow circles, the mutant p53 proteins are shown as light blue ovals, the AP-2 transcription factor are yellow ovals and ETS are green ovals. The DNA sequence is shown in red


extfieldsofstudy: Medicine Chemistry

text:
Crystal structure of a SFPQ/PSPC1 heterodimer provides insights into preferential heterodimerization of human DBHS family proteins

Members of the Drosophila behavior human splicing (DBHS) protein family are nuclear proteins implicated in many layers of nuclear functions, including RNA biogenesis as well as DNA repair. Definitive of the DBHS protein family, the conserved DBHS domain provides a dimerization platform that is critical for the structural integrity and function of these proteins. The three human DBHS proteins, splicing factor proline- and glutamine-rich (SFPQ), paraspeckle component 1 (PSPC1), and non-POU domain–containing octamer-binding protein (NONO), form either homo- or heterodimers; however, the relative affinity and mechanistic details of preferential heterodimerization are yet to be deciphered. Here we report the crystal structure of a SFPQ/PSPC1 heterodimer to 2.3-Å resolution and analyzed the subtle structural differences between the SFPQ/PSPC1 heterodimer and the previously characterized SFPQ homodimer. Analytical ultracentrifugation to estimate the dimerization equilibrium of the SFPQ-containing dimers revealed that the SFPQ-containing dimers dissociate at low micromolar concentrations and that the heterodimers have higher affinities than the homodimer. Moreover, we observed that the apparent dissociation constant for the SFPQ/PSPC1 heterodimer was over 6-fold lower than that of the SFPQ/NONO heterodimer. We propose that these differences in dimerization affinity may represent a potential mechanism by which PSPC1 at a lower relative cellular abundance can outcompete NONO to heterodimerize with SFPQ.

The Drosophila behavior human splicing (DBHS) 2 family of proteins are ubiquitous nuclear proteins implicated in many aspects of nuclear functions (1). Defined by the highly conserved DBHS region that consists of two tandem RNA recognition motifs (RRMs), a NonA/paraspeckle domain (NOPS) and a C-terminal coiled-coil, the members of this protein family have been reported as multifunctional proteins, playing important roles in RNA biogenesis and transport, and subnuclear body (paraspeckle) formation via direction interaction with structural noncoding RNA as well as DNA damage repair (1).
The DBHS family of proteins are exclusively found in the animal kingdom; invertebrate or lower vertebrate typically encode a single DBHS protein, whereas higher vertebrates encode more than one (2). In humans, three DBHS proteins: SFPQ, PSPC1, and NONO, share greater than 70% identity within the DBHS region, flanked by low complexity N-and C-terminal domains that vary in sequence and length among the three paralogues (Fig. 1). DBHS proteins are obligatory dimers; the three human DBHS proteins form either homodimer or heterodimers among the three proteins in vitro (3)(4)(5) and all three possible heterodimers (SFPQ/NONO, SFPQ/ PSPC1, and PSPC1/NONO) have been shown in mouse Sertoli cells (6).
The dimeric nature of the DBHS family of proteins has been confirmed by the crystal structures of human NONO/PSPC1 heterodimer (7), SFPQ homodimer (5), and Caenorhabditis elegans NONO-1 (2) in our previous studies. The crystal structures of the DBHS proteins consistently revealed an intimate dimer interface contributed by all four domains within the DBHS region. The dimer formation is largely driven by the hydrophobic interactions of the second RRM with the NOPS domain as well as the antiparallel coiled-coil interaction, whereas the first RRM is dispensable in dimer formation (2,5,7). Dimerization of the DBHS proteins is not only critical to their structural integrity but also to their functions such that dimerization incompetent DBHS proteins generated by mutation of key hydrophobic residues failed to form subnuclear bodies termed paraspeckles (7).
Combinatorial heterodimerization among the three DBHS proteins diversifies the functional repertoire and also contributes functional overlap and redundancy, evidenced by functional compensation of the loss of DBHS protein by the other paralogues. Specifically, NONO deficiency in murine embryonic fibroblasts derived from NONO knockout mice led to spontaneous up-regulation of PSPC1, which formed a complex with SFPQ in place of NONO and compensated the loss of NONO in DNA repair (8). Similar up-regulation of SFPQ and PSPC1 was observed in patients with syndromic intellectual disability with little or no NONO protein expression due to the mutations in NONO (9). The data suggest that up-regulation of SFPQ and PSPC1 may compensate the down-regulated NONO at least partially; however, the distinct contributions of NONO as in SFPQ/NONO or PSPC1/NONO are irreplaceable by SFPQ/PSPC1 in regulation of synaptic transcription, leading to intellectual disability (9). Taken together, these observations indicate that dimerization among the three DBHS paralogues is likely interdependent as well as spatially and temporally dynamic. Thus, different DBHS heterodimers may have different tissue-or cell-type specific functions, contributing to the myriad of nuclear functions assigned to the DBHS proteins. However, the relative affinity and mechanistic details of the combinatorial dimer states are still unclear.
As a first step toward better understanding the relative affinity and mechanism of combinatorial dimerization of the three human DBHS proteins, we have carried out analytical ultracentrifugation on the SFPQ containing dimers (SFPQ homodimer, SFPQ/PSPC1, and SFPQ/NONO). In addition, we have determined the crystal structure of SFPQ/PSPC1 and analyzed the structural differences of the SFPQ/PSPC1 heterodimer with the previously characterized SFPQ homodimer.
In this study, His-tagged PSPC1 (residues 61-320) was coexpressed with untagged SFPQ (residues 276 -535) and the resulting SFPQ/PSPC1 heterodimer was successfully purified, confirming that the three human DBHS proteins can form all three possible heterodimers. One of the important observations from the course of purification of all six possible dimeric complexes (three homodimers and three heterodimers) is that human DBHS proteins display a preference for heterodimerization; when two DBHS proteins are co-expressed, the less expressed protein is exclusively found in the heterodimer (supporting Fig. S1), indicating that dimerization affinities of het-erodimers are stronger than those of homodimers and/or heterodimers are more stable than homodimers.

Differential dissociation constants of SFPQ-containing dimers
As a first step toward characterizing the relative affinities for dimer formation among the SFPQ-containing dimers, we conducted sedimentation velocity experiments in the analytical ultracentrifuge comparing all three possible SFPQ-containing dimers: SFPQ homodimer, SFPQ/PSPC1, and SFPQ/NONO heterodimers at an initial total protein concentration of 5 M. All three dimers have the same construct boundaries defined by the dimerization domain (RRM1, RRM2, NOPS, and partial coiled-coil) (Fig. 1). It should be noted that the exclusion of the extended coiled-coil domain beyond the dimerization domain was necessary to avoid complication of data analysis because of oligomerization mediated by the extended coiled-coil (5).
The data from sedimentation velocity experiments were fitted to a continuous size (c(s)) distribution model and the results demonstrated that all three samples at an initial concentration of 5 M exist as two major species in solution with sedimentation coefficients of 2.3 and 3.8 S, consistent with monomer and dimer, respectively (Fig. 2, A-C, Table 1). There is also a small proportion of a higher order oligomer with a modal sedimentation coefficient of ϳ5.7 S, particularly in the SFPQ homodimer sample ( Fig. 2A). More importantly, the relative proportion of dimers and monomers differed in each sample with significantly more dimeric species present for the heterodimeric complexes, particularly SFPQ/PSPC1 (Fig. 2C), compared with the SFPQ homodimer, indicating that the dimerization affinities of the SFPQ-containing heterodimers are indeed stronger than that of the SFPQ homodimer. This assertion was confirmed by performing sedimentation equilibrium experiments at multiple rotor speeds and initial protein concentrations. The data were analyzed by a global nonlinear regression method and best fitted to a monomerdimer equilibrium for the SFPQ homodimer and to the 1:1 complex (i.e. AϩB N AB) for the SFPQ-containing heterodimers. The resulting apparent dissociation constants showed that the SFPQ/PSPC1 heterodimer (K D AB3 AϩB of 0.55 M) is almost 20-fold tighter than the SFPQ homodimer (K D 231 of 10.54 M) (Fig. 2, D-F, Table 1). By contrast, the SFPQ/NONO heterodimer (K D AB3 AϩB of 3.65 M) is ϳ3-fold tighter than the SFPQ homodimer (Table 1). A noticeable difference (Ͼ6-fold) in the apparent dissociation constant between the two SFPQ-containing heterodimers is also observed ( Table 1).

Crystal structure of a SFPQ/PSPC1 heterodimer
To gain insight into the preferential heterodimerization of human DBHS proteins, we have crystallized the SFPQ/PSPC1 heterodimer encompassing the dimerization domain (  Crystal structure of a SFPQ/PSPC1 heterodimer and supporting Fig. S2). The structure of the SFPQ/PSPC1 heterodimer was solved by molecular replacement, and refined to 2.3-Å resolution (Fig. 3). Despite high sequence similarity between the two proteins within the dimerization domain (74 and 93% sequence identity and similarity, respectively) (supporting Fig. S2), several key residues that are different in the two proteins allowed unambiguous assignment of SFPQ and PSPC1 (supporting Fig. S3). The final model consists of one heterodimer in the asymmetric unit.
Consistent with the previously characterized structures of human DBHS proteins, the structure of the SFPQ/PSPC1 heterodimer displays a pseudosymmetrical dimer, reflecting high sequence identity within the dimerization domain (Fig. 3). The second RRM domain (RRM2), NOPS, and coiled-coiled domain provide the major dimerization interface. Largely hydrophobic in nature, the interactions of the NOPS domain with the RRM2 and coiled-coil domain of the dimer partner are followed by the right-handed coiled-coil interactions between the two proteins.
The dimerization domain without the first RRM domain (RRM1) superposes well with each other (root mean square difference (r.m.s. deviation) of 1.05 Å for 146 common C␣ positions), whereas the r.m.s. deviation for the overall superposition of the two protein chains is 2.53 Å (224 C␣). The major devia- Plotted are the data at 12,000 rpm (circles) and 18,000 rpm (triangles) for each protein at an initial concentration of 1.5 M with solid lines representing the nonlinear best fits to homo-or heterodimerization equilibrium models. Insets, residuals plotted as a function of radial position for the best fits obtained at both rotor speeds. Note that data and fits shown are representative of global nonlinear least squares analyses employing multiple protein concentrations (0.5-15 M) and rotor speeds that yielded the dimerization dissociation constants reported in Table 1. Global reduced 2 values were Ͻ1.0 for all nonlinear best fits. a Molecular mass of a dimer based on amino acid composition. b Taken from the ordinate maximum of the relevant peak in the c(s) distribution best-fit ( Fig. 2). c Taken from the ordinate maximum of the relevant peak in the c(M) distribution best-fit (data and fits not shown). d Dimerization dissociation constant determined from global nonlinear least squares analyses of sedimentation equilibrium experiments at multiple speeds and initial protein concentrations (data and fits not shown). Estimated error was determined from Monte-Carlo error analysis of the nonlinear best-fit parameters (1000 iterations, 95% confidence).

Crystal structure of a SFPQ/PSPC1 heterodimer
tion from the 2-fold symmetry is due to the arrangement of the two N-terminal RRM1 domains relative to each other. Although the RRM1 domains from each protein superpose well individually (r.m.s. deviation of 0.685 Å, 77 C␣), the observed rotation angle of the two RRM1 domains is 170.4 o , breaking the 2-fold symmetry. In addition, the relative position of RRM1 in SFPQ to the dimer core is tilted by 27 o toward the dimer core compared with that of RRM1 of PSPC1, contributing to further deviation from the 2-fold symmetry. This observation is consistent with the structure of a PSPC1/NONO heterodimer (PDB code 3SDE), the first DBHS protein structure we have characterized, where similar pseudo-symmetry was observed with the major deviation from the 2-fold symmetry contributed by the dispositions of the two RRM1 domains (7). In contrast, in the structure of the SFPQ homodimer (PDB code 4WII), the two RRM1 domains are positioned in a more symmetrical way (rotation angle of 178.8 o ) with no significant deviation in the dispositions of each RRM1 to the dimer core (Fig. 4, A and B) (5). It is, therefore, not surprising that the use of the structure of PSPC/NONO (PDB code 3SDE) as a search model in phasing by molecular replacement found a straightforward structure solution, whereas a more complicated search strategy was required to produce readily interpretable electron density maps when the SFPQ homodimer (PDB code 4WII) was applied as a search model.

Structural comparison between the SFPQ/PSPC1 heterodimer and SFPQ homodimer
Due to high sequence similarity between SFPQ and PSPC1 within the dimerization domain (74 and 93% sequence identity and similarity, respectively), the overall structure of the SFPQ/ PSPC1 heterodimer is indistinguishable from that of the SFPQ homodimer (PDB code 4WII) (Fig. 4A) (5). Analysis of interface using PISA (11) shows the common principal interface involving all four domains in both structures with over 40% of residues directly involved in the dimer interface (supporting Table  S1). The principal dimer interface is largely driven by hydrophobic interactions evidenced by a highly negative gain of solvation energy on complex formation (⌬ i G ϭ Ϫ50.4 kcal/mol with p value 0.027 for the SFPQ/PSPC1 heterodimer; ⌬ i G ϭ Ϫ46.1 kcal/mol with p value 0.061 for the SFPQ homodimer) as calculated by PISA (11).
Closer inspections on the amino acid sequences of the two proteins and superposition of the structures of the SFPQ-containing dimers, however, reveal subtle differences in local primary and tertiary structures. Although most of the primary sequence variations are scattered throughout the dimerization domain, four regions that have a stretch of amino acids with sequence variations are observed (supporting Fig. 2A). With exception of Region 2, these regions are involved in dimer interfaces with the resulting local structures showing subtle differences in the two structures; Region 1, located prior to the RRM1, contributes to the asymmetry of the two RRM1 arrangements described in the previous section, whereas Regions 3 and 4 are located in the NOPS and C-terminal coiled-coil, respectively, where further local structural differences are observed (Fig. 4A). Although most of the intermolecular interactions in these regions made by networks of hydrogen bonds and salt  Table  S2) and detailed below.
Region 1 (residues 276 -287 in SFPQ; 61-80 in PSPC1), showing the most sequence divergence between SFPQ and PSPC1, takes on a different secondary structure. In the struc- Crystal structure of a SFPQ/PSPC1 heterodimer ture of the SFPQ homodimer (PDB code 4WII), this region forms a short ␣-helical structure that is not involved in the dimer interface (5). The corresponding residues of PSPC1 in the structure of the SFPQ/PSPC1 heterodimer form a strand with no secondary structure. Interestingly, an additional hydrophobic residue (Phe-70) in Region 1 of PSPC1 participates in the dimer interface between the two RRM1 domains in the heterodimer, contributing to the asymmetry in the arrangement of the RRM1 domains (Fig. 4, B and C). The different hydrophobic environment associated with this asymmetry at the interface of the two RRM1 domains may contribute toward favoring heterodimerization.
Region 3 (residues 464 -471 in SFPQ; 249 -256 in PSPC1), located in the middle of the NOPS domain, appears to be flexible with different conformations observed in the two structures (Fig. 4, A, D, and E). The electron density of Region 3 of SFPQ (residues 462-474) is in fact disordered in the structure of the SFPQ/PSPC1 heterodimer and thus the residues in this region are not modeled in the final structure (Fig. 4D). Side chains of the majority of the residues in Region 3 of PSPC1 and the SFPQ homodimer are found in the solvent-accessible area, thus not contributing to the dimer interface. However, the first residue of Region 3 (Ala-464 in SFPQ and Met-249 in PSPC1) is directly involved in the dimer interface through hydrophobic interaction with a residue from RRM2 of the dimer partner (Fig.  4, D and E). Interestingly, the partnered hydrophobic residue in RRM2 shows amino acid variation: Tyr-381 in SFPQ, whereas Val-166 in PSPC1. The resulting hydrophobic interaction in the heterodimer shows Met-249 of PSPC1 interacting with Tyr-381 from RRM2 of SFPQ, whereas Ala-464 in Region 3 of SFPQ (not observed in the SFPQ/PSPC1 structure) is likely to interact with Val-166 from RRM2 of PSPC1 when extrapolated from the structure of the SFPQ homodimer (Fig. 4, D and E).
Region 4 (residues 525-535 in SFPQ; 310 -320 in PSPC1) is located at the end of anti-parallel coiled-coil interface. Although all of the residues of Region 4 in PSPC are observed, residues beyond 527 of SFPQ in the heterodimer structures are not observed with a similar observation made in the SFPQ homodimer. In addition, the majority of the residues in this region belong to the beginning of the "coiled-coil interaction motif" (residues 528 -555 in SFPQ) characterized in our previous study, which mediates polymerization of the SFPQ dimers (5). Therefore, it is unlikely that the sequence variations in this region directly influence the dimer interface. Variations in these residues may have, if any, impacts on choice of polymerization partners among the DBHS dimers, which are yet to be characterized.
However, closer inspection of the dispositions of the distal coiled-coil domain reveals that highly conserved residues at the C-terminal end of the NOPS domain (Phe-486, Tyr-490, and Trp-494 for SFPQ; Phe-271, Tyr-275, and Trp-279 for PSPC1) take on drastically different side chain conformations in the two structures (Fig. 4, D-F). Although the structure of the SPFQ homodimer shows similar side chain conformations for these residues in each chain of the dimer (Fig. 4E), the side chains of the corresponding residues take on drastically different conformations in the heterodimer setting (Fig. 4, D and F). Importantly, two of these residues (Tyr-275 and Trp-279 for PSPC1 numbering) have been shown previously to be critical for DBHS dimerization; mutation of these residues to alanine in NONO and PSPC1 prevents homo-and heterodimer formation in yeast two-hybrid analyses (7). Similar observations on the different side chain conformations of these residues were also made in the structure of a NONO/PSPC1 structure. Thus, observation of different side chain conformations in the structure of the SFPQ/PSPC1 heterodimer further reinforces the idea that structural plasticity associated with these residues may influence choice of dimerization partner.
Due to the intimate interaction between NOPS and RRM2 of the partner DBHS protein (RRM2Ј), we further inspected the dimer interface between NOPS and RRM2Ј and found a cluster of hydrophobic interactions between NOPS and RRM2Ј that show differences in the amino acid identity (Fig. 4, G-I). The residues involved in this hydrophobic pocket belong to the RRM2Ј (Val-435 for SFPQ numbering) and the distal end of NOPS (Ala-481, His-483, Tyr-488, and Ser-491 for SFPQ numbering). This region of NOPS also harbors the three conserved aromatic residues mentioned above that show different side chain conformations. The two clusters of hydrophobic residues are found in the different faces of the helix, however; whereas the three conserved aromatic residues face the distal coiled-coil domain of the partner protein (CCЈ), two of the residues (Tyr-488, and Ser-491 for SFPQ numbering) involved in the latter hydrophobic pocket are located in the outer face of the helix with the other two residues (Ala-481 and His-483 for SFPQ numbering) sitting on the hinge region before the beginning of the helix. With the exception of the alanine residue (Ala-481 for SFPQ; Ala-266 PSPC1), the amino acid identity of the four residues involved in this hydrophobic pocket varies among the three DBHS proteins. In combination with the three conserved aromatic residues, the variations of amino acid sequence in this hydrophobic pocket may contribute to the apparent differences in the dissociation constants of the homodimer and heterodimers.
In summary, it appears that preferential heterodimerization of the SFPQ-containing dimers is governed by synergistic effects of subtle variations in the local structures observed in the dimer interfaces encompassing all four domains (two RRMs, NOPS, and coiled-coil) within the dimerization domain.

Discussion
Since the first identification and characterization of members of the DBHS protein family in the early 1990s, a myriad of nuclear functions have been designated to this family of proteins (1). Although exhibiting a large overlap of functions, the three human DBHS paralogues, SFPQ, PSPC1, and NONO, are incapable of complete functional compensation after loss of the other members in some cases (9). This suggests specific roles for each paralogue in different temporal and spatial contexts, presumably acting through the divergent N and C termini flanking the conserved DBHS domain. It has been established that the functional unit of the DBHS proteins is a dimer (3)(4)(5)7); the three paralogues, a product of evolutionary duplication and divergence of an ancestral gene, homoand heterodimerize with each other to add extra layers to the Crystal structure of a SFPQ/PSPC1 heterodimer functional complexities of the DBHS proteins in higher vertebrates.
In this study, built on our previous studies where preference to heterodimerization has been observed from the co-expression and purification of the three human DBHS proteins using a bacterial expression system, we confirmed through sedimentation analyses that the dimerization affinity for the SFPQ-containing heterodimers is indeed tighter than that of the SFPQ homodimer. Surprisingly, the apparent dissociation constant for the SFPQ/PSPC1 heterodimer is more than 19-fold lower than that of the SFPQ homodimer, whereas the SFPQ/NONO heterodimer is ϳ3-fold tighter than the SFPQ-homodimer, suggesting that the majority of DBHS proteins are most likely to exist as heterodimers in vivo. This imposes significant implications in interpretation of the past literature on the human DBHS proteins where individual proteins have been addressed separately to probe their nuclear function. Thus, future work may need to revisit and validate the roles assigned to each individual paralogue taking into account their dimerization states.
The observed 19-fold difference in the apparent dissociation constant between the SFPQ/PSPC1 heterodimer and the SFPQ homodimer was unexpected given that the two paralogues share high sequence similarity within the dimerization domain. Reflecting this similarity, the structural determinants for preferential heterodimerization of SFPQ/PSPC1 compared with the homodimeric SFPQ cannot be attributed to a few key residues; contributed by all four domains (RRM1, RRM2, NOPS, and coiled-coil domain), synergistic effects from small variations in the identity or conformation of many residues largely in the hydrophobic pockets of the dimerization interface appear to favor heterodimerization.
Intrinsic differences in dimerization affinity among all six possible human DBHS dimers are most likely to be the major determinant of the dimerization partner choice in vivo. However, other factors including the differential expression level of the three paralogues and post-translational modifications may also contribute to the equilibrium and dynamics of homo-and heterodimerization in vivo.
Although all three paralogues are ubiquitously expressed in many cell types, relative expression levels are different; SFPQ is highly expressed across all tissue types, whereas varied expression levels of NONO and PSPC1 are observed in the Human Protein Atlas (www.proteinatlas.org) 3 (12). In particular, the expression level of PSPC1 is highly tissue-and cell-type specific; very low expression of PSPC1 is observed in heart muscle and ovary tissue. In HeLa cells, PSPC1 is much less abundant than SFPQ and NONO (6), whereas a similar expression level of PSPC1 was observed in mouse Sertoli cell (13). In addition, the presence of multiple transcript variants producing isoforms of each paralogue adds extra layers of complexities in the combinatorial dimers of the three DBHS paralogues. Some of these variants lack a nuclear localization signal located at the C terminus (14, 15), suggesting that localization of DBHS proteins can be altered depending on the identity and isoform of the dimerization partner. This is particularly relevant with the emerging function and importance of the cytoplasmic pool of DBHS family proteins in neuronal development. Localization of the DBHS family proteins is predominantly nuclear; however, recent studies have shown that a non-nuclear pool of SFPQ is essential for axon motor development and viability (16,17).
The effects of post-translational modifications on dimerization of DBHS proteins are yet to be characterized; however, various post-translational modifications on DBHS proteins including phosphorylation and methylation have been shown to alter their nuclear activities and localization of the proteins from the nucleus to the cytoplasm in some cases (18 -20). It is plausible that the altered cellular activities and localization of DBHS proteins may directly reflect altered dimerization due to post-translational modifications.
In summary, we demonstrate that the apparent dissociation constants of the SFPQ-containing heterodimers are lower than that of the SFPQ homodimer, confirming that heterodimerization is indeed preferred to homodimerization among the SFPQ-containing dimers. A noticeable difference (Ͼ6-fold) in the apparent dissociation constant between the SFPQ/PSPC1 and SPFQ/NONO heterodimers is also observed. These differences in dimerization affinity may represent a potential mechanism by which less expressed PSPC1 can outcompete more abundant NONO to heterodimerize with the most abundant SFPQ in many cell and tissue types. Despite the noticeable difference in the apparent dissociation constants, very subtle differences in the dimerization interfaces are observed in the crystal structure of the SFPQ/PSPC1 heterodimer compared with that of the SFPQ homodimer, reinforcing the concept that the synergistic effects of subtle variations in the sequence and local structures result in stronger affinity in the DBHS heterodimers.

Protein expression and purification
For coexpression and purification of the SPFQ/PSPC1 heterodimer, pETDuet-1-PSPC1(61-320) and pCDF-13-SFPQ(276 -535) were co-transformed into Rosetta2 (DE3) (Novagen). The cells were grown in LB growth medium supplemented with 100 g ml Ϫ1 of ampicillin, 50 g ml Ϫ1 of spectinomycin, and 50 g ml Ϫ1 of chloramphenicol and protein expression was induced with isopropyl 1-thio-␤-D-galactopyranoside (0.5 mM) at an OD of 0.8 -1.0. The cells were harvested after further incubation at 25°C for 16 h. The cell pellet was suspended in buffer A (20 mM Tris-HCl (pH 7.5), 250 mM NaCl, 10% glycerol) and disrupted using a TS series benchtop cell disruptor (Constant Systems Ltd.). The soluble fraction was applied to a nickel-chelating column (Hi-Trap HP, GE Healthcare) and the bound protein was eluted by applying a linear gradient of 25-500 mM imidazole in buffer A. Fractions containing His-PSPC1/SFPQ were pooled and subjected to TEV protease digestion with 1:100 ratio (w/w) at 4°C overnight. Uncleaved heterodimer and His-TEV protease were removed by the second nickel-affinity chromatography. Flow-through fractions containing the cleaved PSPC1/SFPQ heterodimer were concentrated and applied to a HiLoad 16/600 Superdex 200 size exclusion column, pre-equilibrated with 20 mM Tris-HCl (pH 7.5), 0.25 M NaCl. Expression and purification of the SFPQ homodimer (residues 276 -535) and SFPQ(276 -535)/ NONO(53-312) heterodimer was described previously (5). The purified proteins were typically concentrated to 7-10 mg/ml and stored at Ϫ80°C until further use.

Sedimentation velocity
Sedimentation velocity experiments were conducted at 20°C using a Beckman Coulter XL-A or XL-I analytical ultracentrifuge with an An-50 Ti 8-hole rotor using similar methods reported previously (21,22). Briefly, double-sector quartz cells were loaded with 380 l of protein samples at an initial concentration of 5 M and 400 l of reference buffer (20 mM Tris-HCl (pH 7.5), 500 mM NaCl). Data were collected at 230 nm and rotor speed of 40,000 rpm in continuous mode using a using a step size of 0.003 cm without averaging. The partial specific volume of SFPQ, SFPQ/NONO, and SFPQ/PSPC1, buffer density, and buffer viscosity were computed using SEDNTERP (23). Data were analyzed using the continuous size-distribution model employing the program SEDFIT (24 -27) with a continuous size distribution model.

Sedimentation equilibrium
Sedimentation equilibrium experiments were performed in a Beckman Coulter XL-I analytical ultracentrifuge in an An-50 Ti 8-hole rotor employing similar methods reported previously (21,22). Briefly, 120 l of protein samples (0.5-15 M) and 140 l of reference buffer (20 mM Tris-HCl (pH 7.5), 500 mM NaCl) were centrifuged at rotor speeds of 12,000 and 18,000 rpm until sedimentation equilibrium was attained (t ϭ 36 and 48 h for each speed). Experiments were carried out using a step size of 0.001 cm with 10 averages at a wavelength optimized at 234 nm. Data were fitted to a monomer-dimer equilibrium model using SEDPHAT (21,22,28).

Crystallization and X-ray diffraction data collection
Crystals of the SFPQ/PSPC1 heterodimer were grown by hanging-drop vapor diffusion experiments at 20°C. 1.5 l of SFPQ/PSPC1 (10 mg/ml) was mixed with 1.5 l of reservoir solution (50 mM MES (pH 5.5), 0.2 M NaCl, 3% (w/v) aminohexanoic acid, 25% (w/v) PEG3350) and equilibrated against 0.5 ml of the reservoir solution. Crystals were transferred briefly to a drop containing artificial reservoir solution with 30% (w/v) glucose before cryocooling. Diffraction data were recorded on beamline MX2 at the Australian Synchrotron at a wavelength of 0.954 Å at 100 K. The data were processed with XDS (29), and merged and scaled with AIMLESS (30). Data collection and merging statistics are summarized in Table 2.

Structure solution and refinement
The crystal structure of the SFPQ/PSPC1 heterodimer was solved by molecular replacement using PHASER (31) within the CCP4 suite (32). The structure of the PSPC1/NONO heterodimer (PDB code 3SDE (7)) was used as search model after removing all nonprotein atoms. The structure of PSPC1/NONO found a solution with a log likelihood gain of 792 and a Z score of 25.3, locating one dimer in the asymmetric unit. Difference Fourier maps (F o Ϫ F c ) from the first round of refinement with auto-BUSTER (33) revealed several key residues that are different among SFPQ, PSPC1, and NONO, which allowed unambiguous assignment of SFPQ and PSPC1. At this stage, the correct amino acid sequence was incorporated, and iterative model building with COOT (34) and refinement with autoBUSTER was carried out. The final model consists of two chains with SFPQ in Chain A (residues 290 -461 and 475-527) and PSPC1 in Chain B (residues 66 -320), and 136 water molecules. The quality of the model was validated using MOLPROBITY (35). The refinement statistics are included in Table 2. The atomic coordinates have been deposited in the Protein Data Bank as entry 5WPA.