AP-1 Signaling by Fra-1 Directly Regulates HMGA1 Oncogene Transcription in Triple Negative Breast

Author Manuscript Published OnlineFirst on July 12, 2019; DOI: 10.1158/1541-7786.MCR-19-0036 Author manuscripts have been peer reviewed and accepted for publication but have not yet been edited.

AP-1 signaling by Fra-1 directly regulates HMGA1 oncogene transcription in Triple Negative Breast

Cancers

1,2,4 1,2,3 1,2 1,2,3 Claire TOLZA , Fabienne BEJJANI , Emilie EVANNO , Samantha MAHFOUD , 1,2 2 2, 5 Gabriel MOQUET-TORCY , Thierry GOSTAN , Muhammad Ahmad MAQBOOL , Olivier 2,6 1, 2* 1, 2* KIRSH , Marc PIECHACZYK and Isabelle JARIEL-ENCONTRE (*) co-senior authors

1 Equipe labellisée par la Ligue contre le Cancer 2 Institut de Génétique Moléculaire de Montpellier, University of Montpellier, CNRS, Montpellier, France 3 Lebanese University of Beirut, Rafic Hariri Campus, Hadath, Beirut, Lebanon 4 Present address: Centre de recherche en Biologie Cellulaire de Montpellier, University of Montpellier, CNRS, Montpellier, France 5 Present address: Stem Cell Biology Group, Cancer Research UK Manchester Institute, The University of Manchester, UK 6 Present address: Université de Paris, EDC, UMR7216, CNRS, F-75013Paris, France

Running title: Fra-1 regulates HMGA1 expression in TNBCs

Key word : Fra-1/AP-1, HMGA1, Fra-2, transcription, triple negative breast cancer

Corresponding author : Isabelle Jariel-Encontre Address : Institut de Génétique Moléculaire de Montpellier, Université de Montpellier, CNRS, 1919 route de Mende, 34293 Montpellier Cedex, France, e-mail : [email protected] Tel : 0033(0)434359668 Fax : 0033(0)434359634,

Disclosure of conflicts of interest: The authors declare no potential conflicts of interest.

Downloaded from mcr.aacrjournals.org on September 23, 2021. © 2019 American Association for Cancer Research. Author Manuscript Published OnlineFirst on July 12, 2019; DOI: 10.1158/1541-7786.MCR-19-0036 Author manuscripts have been peer reviewed and accepted for publication but have not yet been edited.

Abstract

The architectural chromatin protein HMGA1 and the transcription factor Fra-1 are both overexpressed in aggressive Triple Negative Breast Cancers (TNBCs), where they both favor epithelial-to- mesenchymal transition, invasion and metastasis. We therefore explored the possibility that Fra-1 might be involved in enhanced transcription of the HMGA1 gene in TNBCs by exploiting cancer transcriptome data sets and resorting to functional studies combining RNA interference, mRNA- and transcriptional run-on assays, chromatin immunoprecipitation and chromosome conformation capture approaches in TNBC model cell lines. Our bioinformatic analysis indicated that Fra-1 and HMGA1 expressions positively correlate in primary samples of TNBC patients. Our functional studies showed that Fra-1 regulates HMGA1 mRNA expression at the transcriptional level via binding to enhancer elements located in the last two introns of the gene. Although Fra-1 binding is required for p300/CBP recruitment at the enhancer domain, this recruitment did not appear essential for Fra-1-stimulated HMGA1 gene expression. Strikingly, Fra-1 binding is required for efficient recruitment of RNA Polymerase II at the HMGA1 promoter. This is permitted owing to chromatin interactions bringing about the intragenic Fra-1-binding enhancers and the gene promoter region. Fra-1 is, however, not instrumental for chromatin loop formation at the HMGA1 locus but rather exerts its transcriptional activity by exploiting chromatin interactions preexisting to its binding.

Implications: We demonstrate that Fra-1 bound to an intragenic enhancer region is required for RNA Pol II recruitement at the HMGA1 promoter. Thereby, we provide novel insights into the mechanisms whereby Fra-1 exerts its pro-oncogenic transcriptional actions in the TNBC pathological context.

Introduction

Triple negative breast cancers (TNBCs) constitute a highly aggressive tumor subtype with dismal prognosis. TNBCs cannot benefit from any of the targeted therapies currently available for a number of other mammary tumors, as they neither express the estrogen- (ER) or the progesterone (PR) receptor, nor overexpress the cell membrane receptor Her2/Neu. Their treatments are based on surgery and chemotherapies with time-limited positive responses 1. Therefore, characterizing the molecular mechanisms responsible for their aggressiveness is of utmost importance, as this could enable the development of novel therapies.

The high mobility group A1 gene encodes two proteins, HMGA1a and HMGA1b generated by differential splicing of the same premessenger RNA. These are architectural nuclear proteins that interact with the minor groove of AT-rich genomic domains to regulate gene expression using a variety of mechanisms 2,3. It is often found highly overexpressed in aggressive cancers such as TNBCs 4,5, where it is considered as a master regulator of tumor progression actively contributing to neoplastic transformation and to metastasis formation 4–7. In human breast tumors, HMGA1 overexpression positively correlates with the histological grade, clinical stage, tumor size, lymph node and distant metastases, triple negative status and shorter survival 8. This points to HMGA1 as a useful diagnosis and prognosis biomarker, as well as a possible target for anticancer intervention 5,9–11.

HMGA1 overexpression in cancers is not linked to chromosome rearrangements and is mainly due to transcriptional and post-transcriptional mechanisms. Studies in various tumor types, including breast cancer, have reported that HMGA1 mRNA is the target of different miRNAs, the frequent downregulation of which leads to HMGA1 mRNA overaccumulation 12–16. Overexpression of HMGA1 pseudogenes can also account for HMGA1 increased levels by decoying the miRNAs targeting its mRNA 16. Other studies have also addressed the molecular mechanisms governing HMGA1 gene transcriptional regulation in different cancers. Activation of the Wnt/-catenin/TCF-4 pathway upon APC loss of function in colon tumors leads to upregulation of HMGA1 expression through the binding of -catenin/TCF-4 complexes to two regulatory sites located in the HMGA1 gene promoter 17. In Burkitt’s lymphoma, HMGA1 overexpression was proposed to be under the control of c-Myc also upon binding to the gene promoter region 18. In human kidney embryo- (HEK293) and glioblastoma (T98G) cell lines, the binding of E2F1 to the HMGA1 promoter increases transcription in cooperation with SP1 upon growth factor stimulation 19. HMGA1 gene expression is induced by TPA (12- tetradecanoyl-13-acetate) in the human erythroleukemia cell line K562 via an AP-1-like binding site located 246 bp downstream of the transcription start site (TSS) 20. In the JB6/P+ mouse epidermal cell line, stimulation by TPA triggers HMGA1 gene activation through an AP-1-binding site located 937 bp upstream of the TSS 21. Moreover, HMGA1 expression upon Ras activation is dependent on an AP- 1 motif located at position -1091 bp from the TSS in Rat1a cells 22. This is the same in human

HEK293 cells, as well as in various colon cancer cell lines (HCT116, SW480) expressing an oncogenic Ras 23. On their own, these observations indicate that the transcriptional mechanisms responsible for HMGA1 gene upregulation are very likely to be varied and to depend on both the tumor context and the activated oncogenic pathway(s). Therefore, understanding HMGA1 dysregulation in cancer requires context-specific investigations.

The term AP-1 defines a combinatorial family of dimeric transcription factors mainly contributed by the Fos (c-Fos, FosB, Fra-1/FosL1, Fra-2/FosL2) and Jun (c-Jun, JunB, JunD) multigene family members 24. Whereas the Jun proteins can homodimerize or heterodimerize between them, the Fos proteins must heterodimerize with Jun partners to regulate transcription. Noteworthy, Fos:Jun heterodimers are favored and more stable than the Jun:Jun dimers. AP-1 complexes bind to specific DNA motifs called AP-1/TRE (TGAG/CTCA and its variant forms) or CRE (TGACGTCA and its variant forms) found in gene regulatory elements 25,26. The fine mechanisms whereby the various AP-1 complexes regulate the transcription of their target genes are, however, still ill-understood 24. Physiologically, AP-1 is activated by numerous signaling events and constitutes a dynamic and versatile platform integrating a wide array of extra- and intracellular cues 27,28. It is also a frequent target, and a necessary effector, of activated oncogenes to exert their pathogenic actions. Importantly for the herein study, Fra-1 is frequently overexpressed at the mRNA and protein levels in many epithelial cancers, including TNBCs 29,30. In particular, undue activation of signaling cascades, such as those including the ERK or PKC- kinases, entails both overaccumulation and functional activation of Fra-1 31–36. In the specific case of breast cancers, overexpressed Fra-1 contributes to cell growth and survival, epithelial-to-mesenchymal transition (EMT), cell motility and invasion, as well as metastasis spreading 37–42.

Fra-1 and HMGA1 are both overexpressed in TNBCs where they show strong similarities in their pathogenic effects 4–7,37. Moreover, Fra-1 was reported to regulate positively the HMGA1 gene in human melanoma 43 and esophageral squamous cell carcinoma 44 though it was not addressed whether this was due to a direct transcriptional action. We have therefore addressed here whether the HMGA1 gene could a direct transcriptional target of Fra-1 in TNBCs. Our work showed that this is the case and also unveiled novel transcriptional actions for Fra-1.

Materials and Methods

Cell culture

MDA-MB-231, MDA-MB-436 and MCF-7 cells were obtained from the American Type Culture Collection (ATCC, Rockville, MD, USA). They were cultured in Dulbecco’s modified Eagle’s medium containing 10% fetal calf serum and penicillin/streptomycin (100µg/ml each) in a humidified

5% CO2 atmosphere at 37°C. Cells freshly amplified and frozen after obtention from the ATCC were used every 3-4 months. They were routinely tested for the absence of mycoplasma contamination.

Antibodies

Antibodies against Fra-1 (sc-183 for immunoblotting, sc-376148X for ChIP-seq and a mix of sc- 376148X and sc-28310X for ChIP-qPCR that are directed to the N-terminus of the protein that is not conserved in Fra-2), Fra-2 (sc-604 for immunoblotting and sc-13017X for ChIP-seq and ChIP-qPCR are directed to the Fra-2 domain spanning amino acids 180 to 282, which strongly departs from its counterpart in Fra-1), the DNA-binding domain of Fos proteins (sc-253; which is highly conserved among all Fos family proteins), GAPDH (sc-25778), JunD (sc-74), GFP (sc-8334) and Pol II (sc- 55492X) were from Santa Cruz Biotechnology. Antibodies against Pol II P-Ser5 (clone H14, # 920304) and Pol II P-Ser2 (clone 5, # 920204) were from Biolegend. Antibodies against HMGA1 (Ab4078), H3 (Ab1791), H3K4me3 (Ab8580), H3K4me1 (Ab8895), H3K27ac (Ab4729), and p300/CBP (Ab14984) were from Abcam. Antibodies against H3K36me3 (61101) were from Active Motif and those specifically directed against c-Jun (60A8), JunD (C37F9), p300 (54062) or CBP (7389) were from Cell Signaling. The anti-BrdU (clone G3N2) used as a control in ChIP-qPCR was home-purified. The anti-rabbit (sc-2313) and anti-mouse (sc-2954) HRP-conjugated secondary antibodies were from Santa Cruz Biotechnology. Mouse anti-IgM antibodies (04-6800) were from Invitrogen.

RNAi

For RNA interference studies using siRNA directed to Fra-1 or Fra-2 mRNAs, MDA-MB-231 and MDA-MB-436 cells were transfected with a total of 4.5 nM of either control (siCTL) or of the pool of 3 Fra-specific siRNAs (1.5nM of each of the 3 siRNA) using Interferin (Ozyme) according to the supplier’s specifications. For downregulation of c-Jun or JunB cells were transfected with 4,5 nM of each specific siRNA. For downregulation of JunD cells were transfected with 4,5 nM of a pool of equimolar amounts of siJunD-A and siJunD-B. For double silencing of Fra-1 and Fra-2, a total of 9 nM of either control or Fra-specific siRNA were used. For single silencing of p300 or CBP, 5 nM of

siRNA (control or specific) were used. For double knock-down, a total of 10 nM (control or specific) were used. 72 h post-transfection, cells were collected and utilized for further experiments. siRNA sequences or references are given in Supplementary Table S1D.

Plasmid tranfection

6x105 MDA-MB-231 cells were transfected for 24h with 1µg of plasmid encoding human Fra-1 or Fra-2 ORF in fusion with EGFP (pEGFP-C1, Clontech) using the TransfeXTM reagent (ACS- 80602171) from ATCC.

Immunoblotting

Immunoblotting experiments were performed as previously described 45. Proteins were detected using the Luminata Forte Western HRP Substrate from Millipore. Signal quantifications were performed using the PXi4 Syngene imaging system from Ozyme.

Analysis of breast tumor microarray data

Two breast tumor microarray data sets were analyzed. The CIT breast tumor transcriptomic data set (http://cit.ligue-cancer.net/language/fr/publications/donnees-cit-publiques) was obtained from C. Theillet and S. du Manoir (Array Express accession number E-MTA-365) 46. Transcription profiling was carried out using Affymetrix Human Genome U133 Plus 2.0 microarrays and an Affymetrix GeneChip Scanner 3000. Subsequent images were analyzed using GCOS 1.4 (Affymetrix). The R package affyQCReport was used to generate a QC report for all chips (CEL files). Raw feature data from Affymetrix HG-U133A Plus 2.0 GeneChipTM microarrays were normalized using Robust Multi- array Average (RMA) method (R package affy). More detailed information can be found in Guedj et al. 46. The publicly available breast TCGA dataset was downloaded from the TCGA data portal at https://portal.gdc.cancer.gov, where the detailed informations on the Agilent microarrays can be found. For gene expression data, we selected the level-3 microarray dataset (https://portal.gdc.cancer.gov/legacy-archive/files/44ec9e1b-e66d-4b22-92aa-ca2ac52db667), in which Agilent 244K (G4502A), a custom-designed microarray platform, was used in the experiments. The microarray data were normalized using the Lowess method and presented as calculated log2 ratio. Additional information regarding the level of the data and methods used in the process can be found at the TCGA website. For correlations between HMGA1 and Fra-1 or Fra-2 mRNA expressions in each breast cancer data set, Pearson correlation tests between paired samples were performed.

RNA extraction, reverse transcription (RT), Polymerase Chain Reaction (PCR) and quantitative PCR (qPCR)

0.2x106 siRNA-transfected cells were seeded in 6-well plates. 72h later, RNA extraction was carried out using the GenElute Mammalian Total RNA kit (Sigma-RTN70), according to the manufacturer’s instructions. After a 1h treatment with RNAse-free DNAse1, 1 µg of RNA was used for RT using the Maxima First Strand cDNA synthesis Kit (K1641, ThermoFisher). After a 10-fold dilution, 5 µl of cDNA were used for real-time PCR using the Roche Light Cycler 480 real-time PCR system and Taq Platinium polymerase (Invitrogen). Data were analyzed with the LightCycler software (Roche) and normalized to invariant S26 mRNA levels. The sequences of the primers used for amplifications are given in Supplementary Table S1A and S1B. Semi-quantitative PCRs were performed with Platinium Taq polymerase using 35 amplification cycles and analyzed by electrophoresis through 2% agarose gel.

ChIP-qPCR and ChIP-seq

ChIP were performed as previously described 45. Briefly, 2x106 siRNA-transfected or non-transfected MDA-MB-231 cells were seeded in culture plates 72h before ChIP experiments. They were fixed at room temperature (24°C) using a 1% of paraformaldehyde (Euromedex) solution for 5 minutes. Fixation was stopped by addition of 2.5 mM glycine. Cells were then incubated for at least 10 min on ice in a cell lysis buffer (PIPES 5 mM pH7.5, KCl 85 mM, NP40 0.5%, NaButyrate 10 mM, protease inhibitors). After mild centrifugation, nuclei were lysed in Nuclei Lysis Buffer (Tris-HCl 50 mM pH7.5, SDS 0.125%, EDTA 10 mM, NaButyrate 10 mM, protease inhibitors) at 4°C for 2h and, then, sonicated for 10 cycles at 4°C using the Bioruptor Pico device from Diagenode. After sonication, absorbances at 280 nm (A280) of 1/100 diluted samples were measured and A280nm-adjusted to 0.13 with the nuclei lysis buffer. For analysis of histone H3 and histone marks, 100 µl of chromatin samples were used whereas, in all other ChIPs, twice more chromatin was used. When antibody concentrations were available, 3 µg of antibodies were used. Otherwise 3 µl of commercial solutions were used per 100 µl of chromatin. Chromatin, antibodies and protein G Dynabeads (ThermoFisher) were incubated at 4°C under rotational agitation for 20 hrs. For ChIPs with the H14 (anti Pol II P-Ser5) and H5 (anti Pol II P-Ser2) antibodies (which are IgMs), beads were first incubated with 3 µg of anti-IgM antibodies for 20 min. Purification of immunoprecipitated chromatin was carried out as described in 45. Purified DNA was then subjected to qPCR analysis. The data were normalized with inputs taken from sample before the immunoprecipitation step and treated under the same conditions. The sequences of primers used to amplify different amplicons of HMGA1 and S26 genes are given in Supplementary Table S1E. For Fra-1 and Fra-2 ChIP-seqs, 2 independent ChIP experiments were carried out using 800 µl of chromatin per experiment. Sequencing of the two independent experiments for Fra-1 and

Fra-2 was carried by the MGX genomic platform (Montpellier) using the Hi-seq2500 Illumina sequencer. Data were aligned to the Hg19 genome using the BWA-backtrack software, version 0.7.12- 1039. Aligned reads were then processed using the R package PASHA 47, the replicates were merged and peak calling was performed using the IGB software with the following settings : threshold = 50, Max Gap = 50, Min Run = 50. Fra-1 and Fra-2 ChIP-seq are available on the GEO database, accession number GSE132098.

Run-on assays

Run-on assays were carried out according to Roberts et al. 48. Briefly, 2x106 MDA-MB-231 cells transfected with 4.5 nM of either the control siRNA or the siRNA pool directed against Fra-1 were seeded in culture plates. 72h later, nuclei were prepared using an ice-cold NP-40-containing lysis buffer. After addition of an in vitro transcription cocktail containing BrUTP (Sigma-Aldrich, # B7166), transcription was allowed to proceed at 30°C for 40 min. Then, nuclear RNAs were purified using the Qiagen miRNeasy mini kit (#217004). Nascent mRNAs, that have incorporated BrUTP, were purified by immunoprecipitation using anti-BrUridine antibodies (sc-32323, Santa Cruz biotechnology). Reverse Transcriptions were carried out using 10 µl of purified nascent RNAs, random primers and RT superscript III from Invitrogen. qPCRs were performed on 5 µl of cDNA using two different sets of primers specific for either HMGA1- or GAPDH mRNA for normalization. The sequences of primers are given in Supplementary Table S1C.

Chromosome Conformation Capture (3C) Assay

3C-assay was performed as described in ref. 49 up to the sonication step. In brief, 72h after transfection of MDA-MB-231 cells with 4.5 nM of either siCTL or siFra-1, cells were trypsinized and 1.1x107 cells per condition were crosslinked using 2% paraformaldehyde at 24°C for 10 min. After quenching of crosslinking with glycine and washing in PBS, cells were resuspended in a lysis buffer (Tris-HCl 10 mM pH8, NaCl 10 mM, Igepal CA-630 0.2%, protease inhibitor cocktail) and let on ice for 20 min. After mild centrifugation, pellets were resuspended in lysis buffer and, then, in the DpnII restriction enzyme buffer and homogenized with a Dounce homogenizer. After centrifugation at 14,000 rpm for 5 min, nuclei were resuspended in the DpnII buffer and the suspension was adjusted to 1.67% Triton X100. Nuclei were shacken at 37°C in a thermomixer (Eppendorf) for 1h. 1,500 units of DpnII (NEB, #R0543M) were added per 1.1x107 cells and nuclei were shaken for 6h. 1,500 units of DpnII per 1.1x107 cells were added again and shaking was pursued overnight. Addition of 1,500 units of DpnII was followed by an additional incubation of 4h. After inactivation of DpnII at 65°C for 20 min, digested chromatin was ligated (T4 DNA ligase HC, ThermoFisher, #EL0013) at 16°C overnight. After DNA extraction using phenol/chloroform/isoamyl alcohol, DNA was ethanol-precipitated. The

efficiency of cleavage by DpnII was monitored by qPCR on non-digested and digested chromatin using amplicon not sensitive (control) and sensitive to DpnII. Quantification was carried out using ∆CT and relative concentration (RC). The efficiency of digestion was calculated using the formula: efficiency = 100 x (1-control DpnII RC)/sensitive amplicon RC). Efficiencies were higher than 70%. Efficiency of library construction (ligation) was controlled by agarose gel. After quantification (Qubit, ThermoFisher), 100 ng of DNA were used for PCR amplification. The sequences of the primers used are given in Supplementary Table S1F. To test forward primer efficiency, reverse primers downstream of the forward primers were designed (see Figure 7A and Supplementary Table S1F). The efficiency of amplifications was monitored by PCR on MDA-MB-231 genomic DNA. PCR products were analyzed on 2% agarose gels. The anchor primer is located 552 upstream of the TSS. Due to the nucleotide composition of the promoter region, it was the amplification primer closest to the TSS, which we could show capable of specific PCR amplification in this region.

Statistical analyses

Data were reported as the means ± SEM. The significance between two conditions was assessed using the two-tailed Student’s paired or unpaired t-test with the GraphPad Prism5 software. P-values were considered as significant when *≤0.05 and highly significant when **≤0.01, ***≤0.005, ****≤0.001).

Results

Fra-1 and HMGA1 expressions positively correlate in breast cancers

We started our investigations by computational analysis of two independent gene expression profiling sets of human breast cancers data from two publicly accessible data banks: CIT (Tumor Identity Cards from the French National League against Cancer, Affymetrix microarrays) and TCGA (The Cancer Genome Atlas, Agilent microarrays) (see Figure 1A for breast cancer subtype classification). Our results showed that both Fra-1 and HMGA1 mRNAs (i) are globally higher in ER negative (ER-) than in ER positive (ER+) breast cancers, and (ii) are the most abundant in the aggressive basal-like breast cancers (Figure1B, upper and central panels), a subtype very similar to, but not completely overlapping, TNBCs 50. In contrast, Fra-2 mRNA levels appeared to vary little between the different breast cancer subtypes (Figure 1B, lower panels). Moreover, HMGA1 and Fra-1 mRNA levels strongly correlated in the two databanks (Figure 1C, upper panels). These observations were reminiscent of those made in melanoma 43 and esophageral squamous cell carcinoma 44 . In contrast, no such a correlation was observed between HMGA1 and Fra-2 (Figure 1C, lower panels).

Fra-1 is the main Fos family member expressed in TNBC cell lines

For functional study of possible HMGA1 gene regulation by Fra-1 in TNBCs, we mainly resorted to the MDA-MB-231 cell line, as it is one of the most widely used TNBC cellular model from the basal B subtype. Then, we validated a number of observations in another widely used basal B subtype TNBC cell line, MDA-MB-436. The ER+ MCF-7 cell line was used as a non-TNBC control when necessary.

First, we analyzed by RT-qPCR the relative mRNA levels of the Fos family members in the three cell lines (Figure 2A). The most noticeable results were as follow: (i) Fra-1 mRNA was the main Fos family mRNA expressed in the two TNBC cell lines, which contrasted with its poor accumulation in MCF-7 cells, (ii) Fra-2 mRNA was much less abundant than that of Fra-1 in MDA-MB-231- and MDA-MB-436 cells but was the most expressed Fos mRNA in MCF-7 cells and (iii) c-Fos and FosB mRNAs were hardly expressed in TNBC cells. Similar conclusions were drawn for assays conducted with different mRNA amplicons (Supplementary Figure S1A).

In a second step, we assayed the relative abundances of the Fra-1 (apparent molecular weight of 35-40 kDa) and Fra-2 (apparent molecular weight of 40-45 kDa) proteins in MDA-MB-231 cells. To this aim, we conducted immunoblotting experiments using an antibody directed to the central DNA- binding domain of c-Fos (Fos-DBD) conserved among all Fos family members. Immunoblotting analysis of total cell extracts revealed that the anti-Fos-DBD signal is 10-fold higher for Fra-1 than Fra-2 (see track indicated by an arrow in the left panel of Figure 2B and quantification of

immunoblotting signals in the right panel). As the anti-Fos-DBD antibody recognizes Fra-2 1.5-fold more efficiently than Fra-1 (see Supplementary Figure S1C for details), this indicated that Fra-1 is approximately 15-fold more abundant than Fra-2 in MDA-MB-231 cells. Fra-1 and Fra-2 were formally distinguished in immunoblotting experiments conducted with extracts from cells subjected to RNA interference using siRNAs targeting specifically Fra-1 or Fra-2 mRNAs (see below) and probed with, not only the anti-Fos-DBD antiserum, but also antibodies specific of Fra-1 or Fra-2 (Figure 2B, left panel).

Finally, we compared the relative mRNA (Figure 2C and Supplementary Figure S1B) and protein (Figure 2D) levels of Fra-1 and Fra-2, as well as those of HMGA1 in MDA-MB-231-, MDA-MB-436- and MCF-7 cells. Interestingly, the relative abundances of HMGA1 mRNA and protein between the cell lines correlated with those of Fra-1. On the contrary, no such a match was observed in the case of Fra-2. Moreover, the very low levels of HMGA1 mRNA and protein found in MCF-7 cells did not favor a role for Fra-2 in HMGA1 gene expression either, as this cell line expresses Fra-2 protein levels comparable to those in the two TNBC cell lines.

Fra-1, but not Fra-2, positively regulates HMGA1 expression

We next addressed whether Fra-1 and/or Fra-2 regulate HMGA1 mRNA accumulation in TNBCs. To this aim, HMGA1 mRNA steady-state levels were assayed in the presence or in the absence of Fra-1 and/or Fra-2 after RNAi-induced depletion. We resorted to pools of 3 siRNAs directed to either Fra-1 or Fra-2 mRNAs (thereafter named siFra-1 and siFra-2, respectively) under conditions optimized to minimize possible off-target effects (see ref. 51 for more details and Supplementary Figure S2A for more explanations). As shown in Figure 2E, siFra-1 and siFra-2 were efficient at shutting off the expression of the Fra-1 and Fra-2 proteins, respectively, both when transfected alone or together in MDA-MB-231 and MDA-MB-436 cells. Importantly, siFra-1 did not affect the expression of Fra-2 and, reciprocally, siFra-2 did not affect that of Fra-1. This excluded compensation of the loss of one protein by stimulation of the accumulation of the other, which could have flawed the interpretation of our experiments due to possible partial redundancy, as it has been documented in a few situations 52,53. Individual down-regulation of Fra-1 led to a 2-fold decrease of HMGA1 mRNA steady-state level in the 2 cell lines (Figure 2F and 2G). In contrast, that of Fra-2 had no detectable effect (Figure 2F and 2G). Moreover, the double knock-down of Fra-1 and Fra-2 did not induce any additional decrease in HMGA1 mRNA abundance, as compared to Fra-1 elimination alone (Figure 2F and 2G). This indicated that a possible contribution of Fra-2 to HMGA1 regulation was unlikely to be masked by the high abundance of Fra-1 in TNBC cells (Figure 2A-D). These data were confirmed using another HMGA1 amplicon in RT-qPCR assays (Supplementary Figure S2B). Thus, Fra-1, but not Fra-2,

appeared to be involved in the control of HMGA1 mRNA steady-state level in the TNBC cell lines used.

Next, we asked whether Fra-1 could regulate the HMGA1 gene at the transcriptional level in MDA- MB-231 cells. To this aim, we resorted to nuclear run-on assays measuring nascent RNA formed by transcribing RNA Polymerase II (Pol II) on the HMGA1 gene body in the presence and the absence of Fra-1 after RNAi-mediated knock-down. Our data showed that HMGA1 gene transcription is reduced in the absence of Fra-1 (Figure 2H and Supplementary Figure S2C. Also see Figure 6 below).

Fra-1 and Fra-2 bind to the HMGA1 locus

To unveil the mechanisms whereby Fra-1 controls HMGA1 gene transcription, we first conducted ChIP-seq experiments to identify Fra-1 binding sites (BS) on the MDA-MB-231 genome. Even though Fra-2 did not appear involved in HMGA1 gene transcription from the data presented in Figure 2, we included it in our experiments as a control, as Fra-1 and Fra-2 are expected to share the same DNA- binding specificity, at least on the basis of in vitro DNA-binding experiments 26. Our ChIP-seq experiments (see Materials and Methods for ChIP-seq accession number) indicated that both Fra-1 and Fra-2 bind to 5 sites located 12.75 and 1.06 kb upstream from the HMGA1 TSS (ensemble.org, ENST00000401473.7) and 6.45, 7.45 and 7.88 kb downstream of it, i.e. in the last 2 HMGA1 introns (Figure 3A). Each one of these Fra-1/Fra-2-binding regions contains at least one AP-1 motif (Figure 3A), supporting the idea of direct binding to DNA by the two Fra proteins at these sites. The analysis of other ChIP-seq data available from the UCSC data bank (genome.ucsc.edu) indicated that these AP- 1 BS-containing domains are recognized by diverse Fos family members in different cell- (K562, HeLa, HEPG2, HUVEC, MEF10A) and experimental contexts (Supplementary Figure S3). This further suggested that these sites are actual binding targets for the AP-1 complex in human cells. The other closest Fra-1/Fra-2 peaks were detected 350 kb upstream and 685 kb downstream of the HMGA1 TSS. Both of them are located in neighbouring topologically-associating domains (or TADs) in other cell backgrounds (Supplementary Figure S4). As TADs are essentially conserved amongst cell types and as enhancer/promoter interactions are reputed to occur essentially within TADs 54,55, these very distant Fra-1/Fra-2-BSs are most unlikely to be involved in Fra-1-dependent regulation of HMGA1. The fact that Fra-2 was found to bind the same sites as Fra-1 at the HMGA1 locus may appear puzzling, as our work did not point to a role for it in HMGA1 regulation (see Figure 2 and below). It must however be taken into consideration that binding of transcription factors to their DNA target sites can be dynamic and that binding to DNA by a transcription factor is not necessarily followed by transcriptional activation (see Discussion).

It is of note that another gene (C6orf1) lies in the close vicinity of HMGA1 (Figure 3A). We therefore asked whether its transcription could interfere with that of HMGA1. RT-qPCR assays using two

different amplicons in its 5'UTR showed that the C6orf1 mRNA accumulates to levels >100-fold less than the HMGA1 one (Figure 3B), making it unlikely that its transcription would affect that of its neighbor gene.

The HMGA1 intronic domains bound by Fra-1 and Fra-2, but not the intergenic ones, show features of active enhancers

We next asked whether the domains bound by Fra-1 and Fra-2 at the HMGA1 locus display features characteristic of enhancers. Using ChIP-qPCR, we first assayed histone H3 density and three of its post-translational modifications marking gene regulatory elements and their activity. These were H3K4me1 and H3K4me3, the ratio of which is generally low by TSSs and high at enhancers, and H3K27ac, which is usually associated with active enhancers and promoters 56–58. Twenty amplicons located at different positions of the HMGA1 locus were used in this analysis, certain of them encompassing the Fra-1/Fra-2 BSs and the gene promoter (Figure 3A). The histone H3 profile indicated lower nucleosome density by the TSS (+ 0.05 kb amplicon) (Figure 3C), consistently with the well-known depletion of nucleosomes in regions of transcription initiation. It also showed reduced nucleosome amounts at the level of the -12.63, -1.02, and +7.95 amplicons, which was suggestive of the presence of accessible regulatory and/or structural elements in these regions. As expected, H3K4me3 was high at the level of the TSS but low at those of the Fra-1/Fra-2 BSs, indicating that the latter do not correspond to non-annotated promoters (Figure 3D). In contrast, H3K4me1 was very low by the TSS and higher at other regions of the HMGA1 locus, including the Fra-1/Fra-2 BSs, especially those located in the last two introns (Figure 3E). H3K27ac was high by the TSS in agreement with active transcription of the HMGA1 gene (Figure 3F). Interestingly, it was also high around the most 3' Fra-1/Fra-2 BSs of the locus but not on those upstream of the TSS. p300 and CBP are histone acetyl transferases exerting redundant functions in acetylation of H3K27. They are often referred as a single entity due to their high sequence homology and functional similarity 59 and have been described to be part of active enhancer signatures 56,60. We therefore also assessed their presence on the HMGA1 locus by ChIP-qPCR, which revealed their binding predominantly in the intronic regions of HMGA1 gene that are bound by Fra-1 and Fra-2 (Figure 3G).

Thus, altogether, our data indicate that only the domains bound by Fra-1/Fra-2 in the last two HMGA1 introns show features of active enhancers.

c-Jun and JunB are the main Jun partners of Fra-1 at the HMGA1 locus

As the Fos family proteins bind to DNA mainly as Fos:Jun heterodimers, we addressed which Jun family member(s) might be involved in HMGA1 gene regulation. To this aim, we first downregulated

individually each one of the 3 Jun proteins using specific siRNAs and monitored HMGA1 mRNA levels 72h post-siRNA transfection. All 3 siRNAs were efficient at downregulating their respective targets (Figure 4A), but only the depletions of c-Jun and JunB led to a decrease in HMGA1 mRNA steady-state level (Figure 4B). In contrast, that of JunD led to increased HMGA1 mRNA abundance. We then monitored c-Jun, JunB and JunD binding at the HMGA1 locus in the presence or the absence of Fra-1 or Fra-2 by ChIP-qPCR, after verifying that RNAi-induced removal of either Fra-1 or Fra-2 from the HMGA1 locus (Supplementary Figure S5A and D) was not compensated by increased binding of the other Fra (Supplementary Figure S5B and C), as this might have flawed our interpretations. In the presence of the Fra proteins, all 3 Jun proteins were detected at the chromatin sites bound by Fra-1 and Fra-2 (Figure 4C, D and E; black bars). Importantly, the recruitment of c-Jun and JunB appeared to be dependent on the binding of Fra-1 (Figure 4C and D; blue bars), but not on that of Fra-2 (Supplementary Figure S5E and F; purple bars). JunD recruitement was, however, found dependent on neither Fra-1- nor Fra-2 binding (Figure 4E, blue bars, and Supplementary Figure S5G, purple bars), suggesting that Fra-1 or Fra-2 are not JunD partners at the HMGA1 locus in MDA-MB- 231 cells. Altogether, these data suggest that Fra-1:c-Jun and Fra-1:JunB are the main AP-1 dimers involved in HMGA1 high expression in MDA-MB-231 cells. Moreover, as the depletion of Fra-1 was not compensated by increased binding of c-Jun, JunB or JunD at Fra-1-binding sites, it seems unlikely that homo- or heterodimers between Jun family members contribute significantly to HMGA1 gene expression.

Fra-1, but not Fra-2, is required for p300/CBP recruitment at the HMGA1 locus

We then asked whether Fra-1 could be involved in the recruitment of p300/CBP, as we previously showed that Fra-1 is required for the recruitment of p300 on the uPA/PLAU locus in MDA-MB-231 cells 45. This was achieved by ChIP-qPCR after knock-down of Fra-1 by RNAi, as well as that of Fra- 2 as a control.

The data presented in Figure 5A and 5B show that Fra-1 down-regulation, but not that of Fra-2, entailed strong reduction of p300/CBP level in the HMGA1 intragenic enhancer regions. To address whether p300 and/or CPB could be involved in the transcriptional regulation of HMGA1, we next knocked them down individually or together using siRNAs specific for each one of them and assayed HMGA1 mRNA accumulation. Individual and double knock-down of p300 and CBP were efficient, as shown by immunoblotting (Figure 5C). We also confirmed recent observations by others 59 that the two enzymes have redundant enzymatic specificity, at least as assayed by H3K27 acetylation, as the individual deletion of either acetyltransferase had no detectable effect on the bulk of cellular H3K27ac whereas that of the two enzymes together led to a dramatic reduction in H3K27ac in the cell (Figure 5C). No change in HMGA1 mRNA abundance was observed, including when both p300 and CBP

were knocked down (Figure 5D and Supplementary Figure S6A for analysis of another HMGA1 amplicon). This contrasted with uPA/PLAU mRNA taken as a positive control, as the abundance of the latter mRNA was significantly reduced upon p300/CBP knock-down as already reported 45 (Figure 5E and Supplementary Figure 6B for analysis of another uPA/PLAU amplicon). Altogether, these data suggested that p300/CBP do not have key roles in HMGA1 expression (see Discussion).

Fra-1 is required for RNA Polymerase II recruitment at the TSS

Then, we addressed whether Fra-1 could be involved in RNA Polymerase II (Pol II) recruitment at the HMGA1 locus. We also asked whether it could affect the phosphorylations of the C-terminal domain (CTD) of its large enzymatic subunit RPB1, as they provide information on Pol II activity. Usually, (i) phosphorylation of Ser5 (P-Ser5) from the heptad repeats of the CTD is principally found at TSSs and by the beginning of genes and is indicative of Pol II transcription initiation activity and (ii) phosphorylation of Ser2 (P-Ser2) from the same repeats is low or absent at TSSs but found on gene bodies with culmination of signals by the end of transcribed genes and is indicative of elongating Pol II.

ChIP-qPCR showed a peak of Pol II around the TSS and lower signals on the gene body under the reference condition (Figure 6A and 6B, black bars). This indicated that HMGA1 is a paused gene, as observed in approximately 30% of all metazoan genes 61. Fra-1, but not Fra-2, knock-down entailed strong reduction of Pol II both at the TSS and on the gene body (Figure 6A, blue bars, and 6B, purple bars). Importantly, Pol II decrease at the HMGA1 locus in the absence of Fra-1 was specific as (i) no global decrease in neither Pol II nor its phosphorylated forms were detected by immunoblotting (Supplementary Figure S7A) and (ii) Pol II recruitment was not altered on the S26 ribosomal subunit gene promoter, the activity of which is independent of Fra-1 (Supplementary Figure S7B). These data pointed to Fra-1, but not Fra-2, as crucial for Pol II recruitment at the beginning of the HMGA1 gene, the lower Pol II levels downstream of the TSS in the absence of Fra-1 being most likely the mere consequence of reduced entry of RNA polymerases on the gene.

Pol II P-Ser5 and Pol II P-Ser2 dropped down with a fold factor similar to that of Pol II reduction upon Fra-1 knock-down but remained unchanged upon elimination of Fra-2 (Figures 6C, 6D, 6E and 6F. Also see Supplementary Figure S7C and S7D). This was consistent with, on the one hand, the absence of effect of Fra-2 on HMGA1 gene transcription and, on the other hand, a role for Fra-1 in Pol II recruitment but not on its transcription initiating and elongating activities.

Finally, further supporting a role for Fra-1, but not for Fra-2, in transcriptional regulation of HMGA1 gene, trimethylation of histone H3K36 on the gene body, which is a mark deposited by transcribing Pol II-containing complexes, was found reduced upon RNAi-mediated depletion of Fra-1 but not by that of Fra-2 (Figure 6G and 6H).

The intragenic enhancers bound by Fra-1 and Fra-2 interact with the HMGA1 promoter

Pol II P-Ser5 and Pol II P-Ser2 showed unusual profiles in the ChIP-qPCR experiments presented in Figure 6, as a high amount of Pol II P-Ser5 (i.e. transcription-initiating Pol II) was found at the 3' end of the gene (Figure 6C and 6D, black bars) and strong signals for Pol II P-Ser2 (i.e. transcription- elongating Pol II) were found in the TSS region (Figure 6E and 6F, black bars). A possibility to explain this was to infer chromatin interactions bringing into close spatial proximity the HMGA1 gene promoter and the intragenic regulatory elements bound by Fra-1 and Fra-2. To address it, we resorted to chromosome conformation capture (3C) experiments in the presence, but also in the absence, of Fra-1 to determine whether it could regulate the formation of this loop. 3C libraries using the DpnII restriction enzyme were constructed as described in Materials and Methods. According to the 3C design (Figure 7A), possible loops (doted lines numbered A1 to A14), were assessed by PCR between an "anchor" located in the promoter region (horizontal green arrow) and primers (horizontal blue arrows, see Supplementary Table S1F for sequences and exact positions) located at different positions on the locus. The efficacy of the forward primers was monitored on non-digested MDA-MB-231 genomic DNA using reverse primers (a to n; purple arrows) located 100-250 bp apart by electrophoresic analysis of PCR products (Figure 7B). 3C results of two independent experiments showed that the HMGA1 promoter region does not detectably interact with Fra-1 BS located 12 kb upstream of the TSS (Figure 7C and 7D, upper panels). This, in addition to the absence of histone marks specifying active enhancers (Figure 3), further argued for no, or weak, role of this domain in Fra-1-dependent regulation of HMGA1 gene transcription in MDA-MB-231 cells. In contrast, interactions were detected between the promoter and intragenic domains (Figure 7C and 7D, upper panels). Interestingly, the frequency of these interactions dropped beyond the intronic enhancer domains harboring the Fra-1 BSs. Altogether, these results suggested that the region containing the Fra-1-bound enhancers and the promoter are in close spacial proximity. The resolution of the 3C experiment did not allow to rule out the possibility that the signals detected in the region ±2kb around the TSS (A4 to A7) were due to the mere sequence proximity to the TSS and not to chromatin folding. A similar profiles were obtained upon Fra-1 knock-down (Figure 7C and 7D, lower panels) indicating that the formation or the maintainance of the chromatin 3D interactions on HMGA1 locus are not mediated by Fra-1.

Discussion

Work by others has previously shown that the Fra-1 transcription factor and the chromatin-associated protein HMGA1 are both overexpressed in TNBCs, where they both contribute to tumorigenesis and metastasis with widely overlapping biological effects 4–6,37,41,42. Our herein work now indicates that HMGA1 gene is a direct transcriptional target of Fra-1 in TNBCs, which is consistent with the similarity in the breast cancer traits they participate to control. Indeed, our bioinformatic analysis of transcriptomic tumor sample data from two publicly available cancer databanks (CIT and TCGA) showed that HMGA1 and Fra-1 mRNA levels are positively correlated in breast cancers, with the highest expression in TNBCs. Moreover, we provide molecular evidence that Fra-1 controls transcriptionally HMGA1 via intragenic binding owing to molecular studies conducted in one of the most widely used TNBC reference cell lines.

Numerous studies have described the pathological consequences of Fra-1 overexpression in cancers, including in TNBCs. In contrast, the transcriptional mechanisms whereby Fra-1 controls gene expression remain essentially unknown. This is largely due to the molecular and biological complexity and versatility of the AP-1 transcription complex family. In particular, a number of its components can show functional redundancy and be co-expressed in the same cells 27. This often makes it difficult to conduct functional investigations and to draw non-ambiguous molecular conclusions. From this standpoint, TNBC cell lines, including the MDA-MB-231 cell line used in this study, provide a relatively favorable experimental situation, as they principally (over)express Fra-1 30 even though they can also express Fra-2, but to a much lesser extent as we showed here. The respective roles of Fra-1 and Fra-2 in HMGA1 gene regulation are discussed successively below.

Our Fra-1 ChIP-seq data indicated 5 Fra-1 BSs at the HMGA1 locus with the next closest Fra-1 BSs located 350 kb upstream and 685 kb downstream of the gene. The latter two BSs are located in TADs different from the one bearing the HMGA1 gene. We considered that they were unlikely to control HMGA1 transcription, as TADs are essentially conserved between cell types and enhancer/promoter interactions principally occur within TADs 54,55. We also found unlikely that the -12.75 kb and -1.06 kb domains harboring Fra-1 BSs could play major parts in HMGA1 control by Fra-1. Concerning the former one, it was marked neither by p300/CBP nor by H3K27ac, which are two acknowledged features of active enhancers. Moreover, we found no evidence of spatial proximity with the TSS region in our 3C experiments. Concerning the latter one, p300/CBP and H3K27ac marks were also weak at its level. It is however of note that, even though it is not a major HMGA1 enhancer in MDA- MB-231 cells, it can manifest enhancer activity in luciferase reporter assays out of its gene context and was implicated in HMGA1 regulation by AP-1 in other cell lines 22,23 . It was however not addressed whether Fra-1 was an AP-1 component contributing to its activity in these situations. Moreover, it is important to keep in mind that TNBCs represent a pathological context with abnormally high levels of

Fra-1, which might influence enhancer selection at the HMGA1 locus, as compared to other cell contexts where other AP-1 proteins are dominant.

In contrast, strong evidence argues for transcriptional regulation of HMGA1 gene by intragenic regulatory elements bearing the 3 Fra-1 BSs located within the last two introns. Firstly, they were associated with clear marks of active enhancers, i.e. (i) a high H3K4me1/H3K4me3 ratio, (ii) high levels of H3K27ac and (iii) strong p300/CBP binding. Secondly, we demonstrated chromatin interactions between the intronic domains harboring them and the gene promoter region in 3C experiments. Noteworthy, a similar regulation by intronic enhancers bound by Fra-1 has already been documented in the case of the Fra-1/FosL1 gene itself in thyroid tumors were Fra-1 overaccumulation depends on a transcriptional feedforward mechanism in response to oncogenic Ras 62. Importantly, chromatin loop formation at the HMGA1 locus did not appear to be dependent on Fra-1, which departs from what has been observed by others in the case of a Fra-1-dependent chromatin loop forming between the P1 and P2 promoters of the Zeb2 gene 63.

An intriguing point of our work concerns p300/CBP, as binding of p300 to Fra-1 and functional cooperation between the two proteins is one of the rare Fra-1 transcriptional mechanisms that has been described so far 45,64. Similarly to what we have previously reported at the uPA/PLAU locus in MDA- MB-231 cells 45, prior Fra-1 binding is required for p300/CBP recruitment at the intronic enhancer region of HMGA1 gene. However, whereas recruitment of p300 is required for optimal uPA/PLAU gene expression 45, it does not seem necessary for HMGA1 gene expression. The latter observation is however consistent with recent work by others 59 showing that p300/CBP knock-down in mouse embryo fibroblasts entails reduction of expression of a relatively modest subset of genes (11.3 % of the expressed genes are subjected to a 2-fold factor repression) that include uPA/PLAU but not HMGA1 (See Supplemental Data, Table 5 in ref. 59). Besides this, p300 was proposed to bridge promoter-bound RNA Pol II to a Fra-2/JunD-bound enhancer lying 5 kb upstream of the TSS in the case of the iNos gene and, thereby, to contribute to gene transcription 65. We however feel a similar role for p300/CBP to be unlikely in the case of HMGA1 in TNBCs, as depletion of p300/CBP had no effect on gene expression. Further studies are therefore required to elucidate why p300/CBP is recruited at the HMGA1 locus and whether and/or when during tumorigenesis this recruitment has a functional impact.

An important point concerns the relationship between Pol II and Fra-1 at the HMGA1 locus. To our knowledge, our work describes for the first time Fra-1-dependent recruitment of Pol II at a gene TSS. This mechanism is however gene-specific, as Pol II recruitment at the uPA/PLAU locus is independent on Fra-1 in MDA-MB-231 cells 45. It is also worth underlining that, whereas Fra-1 positively influences the fraction of transcription-elongating Pol II on the uPA/PLAU gene body 45, it does not seem to control phosphorylation of Pol II CTD on Ser2 or Ser5 in the case of HMGA1. Thus, on their own, the study of functional relationships between p300/CBP and Pol II indicate that Fra-1 can exert

its transcriptional actions via a variety of mechanisms that can be gene-specific. It would therefore be interesting to address whether the Fra-1-dependent mechanisms operating on HMGA1 are also cell- and/or intracellular signaling-specific. Human melanoma 43 and esophageral squamous cell carcinoma 44 where Fra-1 was reported to regulate positively the HMGA1 gene should represent favorable situations to address this point.

As Fra-1 is strictly dependent on heterodimerization to bind to DNA, we investigated whether and which Jun member(s) could be involved in HMGA1 expression and identified c-Jun and JunB as Fra-1 main Jun partners at the HMGA1 locus in MDA-MB-231 cells. Interestingly, RNAi-mediated depletion of Fra-1 was not compensated by increased binding of c-Jun and JunB homo- or heterodimers at the Fra-1/Fra-2 BSs. This strengthened the idea that Fra-1 is essential for high HMGA1 expression in MDA-MB-231 cells. Surprisingly, we also observed that JunD represses HMGA1 expression in a Fra-1- and Fra-2-independent manner, suggesting that JunD exerts its effects through homodimerization or heterodimerization with non-Fra partners. The fact that we could see different types of AP-1 dimers involving Fra-1, Fra-2, c-Jun, JunB and JunD at the same BSs at the HMGA1 locus raises the important question of the binding dynamics of the different possible AP-1 dimers at these sites. Recent work has shown that many transcription factors might behave dynamically at their DNA-binding sites with residence time on DNA in the second range. Elucidating the biological/biochemical reasons for this is currently the objet of intense research with no unique answer (for a review, see 66). In the specific case of the HMGA1 gene in MDA-MB-231 cells, it would, for instance, be interesting to clarify the relative contributions of the various AP-1 dimers, whether their action is coordinated or not and why transcription-activating (e.g. Fra-1:JunB and Fra- 1:c-Jun dimers) and transcription-repressing dimers (JunD-containing AP-1 dimers) can bind at the same DNA sites. Along this line, a puzzling point of our investigations concerns Fra-2, as our ChIP- seq experiments show binding to the same DNA elements as Fra-1 but our functional analyses did not highlight any role for it in HMGA1 gene transcription. At this stage of investigation, we cannot formally exclude a modest transcriptional contribution for Fra-2 that would be masked by the much higher abundance of Fra-1 and that the level of Fra-2 is too low to exert any transcriptional action on HMGA1 even in the absence of Fra-1. Another explanation would refer to the notion that binding of transcription factors to their DNA targets does not necessarily mean transcription stimulation. A number of them, such as SRF 67 for example, have been described to be latent after DNA binding and to await for functional activation by intracellular signaling events. It could therefore not be excluded that cell signaling occurring within TNBCs provides activation signals for Fra-1 but not for Fra-2, at least at the HMGA1 locus. Further studies are required to clarify this point

Authors’ contributions

C. Tolza, M. Piechaczyk and I. Jariel-Encontre designed the experiments. C. Tolza carried out the experiments with the help of F. Bejjani, E. Evanno and S. Mahfoud. C. Tolza, F. Bejjani and G. Moquet-Torcy performed the setting up of ChIP-seq experiments. T. Gostan exploited breast cancer databanks. C. Tolza, F. Bejjani, M.A. Maqbool and O. Kirsh analyzed Fra-1 and Fra-2 ChIP-seq data. M. Piechaczyk and I. Jariel-Encontre wrote the manuscript.

Acknowledgements This work was supported by the CNRS and the French Ligue Nationale contre le Cancer, of which the M. Piechaczyk's group is an "Equipe Labellisée", and the GSO Cancéropole "Emergence" program. C.Tolza, F. Bejjani. and E. Evanno were recipients of fellowships from the French “Ligue Nationale contre le Cancer”, the Lebanese “Association de Spécialisation et d'Orientation Scientifique” and the EpiGenMed Labex, respectively. We thank C. Theillet and S. du Manoir for access to the CIT breast cancer data bank. We also thank H. Parrinello and M. Rohmer from the MGX genomic platform (Montpellier, France) for the Fra-1 and Fra-2 ChIP sequencing. We are grateful to J.C. Andrau for helpful discussions and critical reading of the manuscript.

Dedication This work is dedicated to Tamara Salem, our late dear friend and colleague.

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Legends to Figures

Figure 1

Fra-1 and HMGA1 mRNA expression in breast cancers. (A) Breast cancer subtype classification in the CIT and TGGA databanks. Of note, the classifications of the breast cancer subtypes in the two data banks do not overlap exactly. In the CIT databank, the basal-like and mApocrine subtypes are ER negative, the four other ones (luminal A, B, C and normal-like) being ER positive. In the TCGA databank, basal-like, Her2 and normal-like subtypes are ER negative and the luminal A, B and C subtypes are ER positive. The numbers of tumors in the diverse subtypes are as follows. CIT: basL = 47, mApo = 37, lumC = 63, lumB = 87, lumA = 81, normL = 113. TCGA: basL = 81, Her2 = 53, lumC = 0, lumB = 114, lumA = 210, normL = 8. ER+ and ER- subtypes are indicated. (B) Box plots of Fra-1 (upper panels), HMGA1 (central panels) and Fra-2 (lower panels) relative mRNA expressions in the various breast cancer subtypes, as defined by CIT (left) and TCGA (right). (C) Correlation between HMGA1 and Fra-1 (upper panels) or Fra-2 (lower panels) mRNA expressions in breast cancers. Correlations scores were calculated from CIT and TCGA databank data.

Figure 2

Fra-1-dependent HMGA1 transcription. (A) Relative mRNA levels of the Fos family members in MDA-MB-231, MDA-MB-436 and MCF-7 cells. Steady-state levels of mRNAs were quantified by RT-qPCR. The value of Fra-2 was arbitrarily set to 1 in all cell lines. (B) Fra-1 and Fra-2 protein abundances in MDA-MB-231 cells. Left panel: Typical immunoblotting experiments using antibodies specific for Fra-1 (left), for the central DNA binding domain (Fos-DBD) conserved among Fos proteins (middle) and for Fra-2 (right). Fra-1 (271 amino acids) migrates faster than Fra-2 (326 amino acids). RNAi experiments with siFra-1 or siFra-2 demonstrated the specificity of the antibodies used and allowed to discriminate Fra-1 and Fra-2 electrophoretic bands upon detection with the anti-Fos- DBD antibody. GAPDH was used as an invariant internal electrophoresis loading control. Right panel: Relative signals of Fra-1 and Fra-2 proteins. Relative signals were calculated from quantification of immunoblotting signals (see brackets in left panel) in the control condition (see arrow) from experiments conducted with the anti-Fos-DBD antibody. The values were arbitrarily set to 1 for Fra-2. The calculation of Fra-1 and Fra-2 relative abundances must take into account that anti-Fos-DBD antibody recognizes Fra-2 1.5-fold more efficiently than Fra-1 (See text and Supplementary Figure S1C). (C) Relative mRNA levels of Fra-1, Fra-2 and HMGA1 mRNAs. Steady-state levels of mRNAs were quantified by RT-qPCR. The values in MDA-MB-231 cells were arbitrarily set to 1. (D) Comparison of relative levels of Fra-1, Fra-2 and HMGA1 proteins in the 3 cell lines. GAPDH was used as an invariant internal electrophoresis control. (E) Efficacy of Fra-1 and Fra-2 protein down-

regulation by RNAi. MDA-MB-231 (left panels) and MDA-MB-436 (right panels) cells were transfected using either a control (siCTL) or pools of 3 siRNAs directed to Fra-1 (siFra-1) or Fra-2 (siFra2). Black arrows indicate non-specific bands. Fra-1 and Fra-2 protein levels were monitored by immunoblotting using GAPDH as an invariant internal loading control. (F and G) HMGA1 mRNA levels upon RNAi-mediated Fra-1 and/or Fra-2 depletion in MDA-MB-231 (F) and MDA-MB-436 (G) cells. Steady-state levels of HMGA1 mRNA were quantified by RT-qPCR 72 hrs after siFra-1 and/or siFra-2 transfection using S26 mRNA as an internal standard. (H) Transcription of HMGA1 upon RNAi-mediated Fra-1 depletion. HMGA-1 nascent RNAs in MDA-MB-231 cells were quantified by run-on assays followed by RT-qPCR using an amplicon located 1.22 kb downstream of the HMGA1 TSS. (A-H) The data presented correspond to 3 independent experiments. Error bars indicate SEM. Results of the 2-tailed Student unpaired t-test are indicated on the graphs. The sequences of primers used in RT-qPCR experiments are given in Supplementary Table S1A and S1C and those of siCTL, siFra-1 and siFra-2 in Supplementary Table S1D.

Figure 3

Fra-1 and Fra-2 binding at the HMGA1 locus in MDA-MB-231 cells. (A) Fra-1 and Fra-2 binding at the HMGA1 locus. The HMGA1 locus (RefSeq) is represented using the IGB software. Fra-1 and Fra-2 ChIP-seqs are merges of two independent experiments. The black arrow indicates the HMGA1 TSS. Fra-1 and Fra-2 peaks are indicated by red arrows. The sequences of the AP-1 binding sites in the Fra-1 and Fra-2 peaks are indicated, as well as their positions relative to the HMGA1 gene TSS. The horizontal black bars indicate the 20 amplicons used in ChIP-qPCR experiments presented in Figure 3C-G, as well as in Figures 4C-E, 5A-B and 6. The numbers indicate the center of these amplicons relative to the HMGA1 gene TSS. Amplification primer sequences are given in Supplementary Table S1E. (B) Relative steady-state levels of HMGA1 and C6orf1 mRNAs. RT-qPCR were performed using 2 sets of primers (HMGA1-3 and C6orf1-1, left panel, and HMGA1-4 and C6orf1-2, right panel) allowing the amplification of 2 amplicons from HMGA1 and from C6orf1. Mean values were calculated from 3 independent experiments with values for HMGA1 set to 1. The results of the Student’s paired t-test are indicated on the graphs. Primer sequences are given in Supplementary Table S1A and S1B. *P-value ≤0.05, **P≤0.01, ***P≤0.005, ****P≤0.0001. (C) ChIP-qPCR analysis of histone H3, (D) of H3K4me3, (E) of H3K4me1, (F) of H3K27ac and (G) of p300/CBP at the HMGA1 locus. Values are the mean of at least 3 independent experiments. For calculation of means, all values were normalized to that of amplicon +1.89 (C) and (F), amplicon - 0.32 (D), amplicon +5.63 (E) and amplicon +7.95 (G). Bars indicate SEM. ChIP-qPCR amplification primer sequences are given in Supplementary Table S1E.

Figure 4 c-Jun and JunB are the main partners of Fra-1 for the regulation of HMGA1 gene expression in MDA-MB-231 cells. (A) Efficiency of the down-regulation of c-Jun, JunB and JunD upon siRNA transfection. MDA-MB-231 were transfected with 4.5nM of either a control siRNA (siCTL) or siRNA directed to c-Jun (sic-Jun), JunB (siJunB) or JunD (siJunD, corresponding to a pool of 2 siRNAs). 72h after transfection, c-Jun, JunB and JunD protein levels were assayed by immunoblotting, GAPDH being used as an electrophoresis loading control. (B) HMGA1 mRNA steady-state level in MDA-MB- 231 cells upon Jun family members downregulation. 72h after transfection of siCTL, sic-Jun, siJunB or siJunD, mRNA levels of HMGA1 were monitored by RT-qPCR. Quantifications were performed on 3 independent experiments. (C, D and E) Binding of c-Jun (C) JunB (D) and JunD (E) at the HMGA1 locus in the presence or in the absence of Fra-1. ChIP-qPCR experiments were conducted 72h after transfection of either siCTL (black bars) or siFra-1 (blue bars). The black arrow indicates HMGA1 TSS. Vertical red arrows indicate the Fra-1 and Fra-2 binding sites (see Figure 3A). The values are the mean of 5 (c-Jun, JunD) or 7 (JunB) independent experiments and were normalized to that of amplicon +7.95 under control conditions, which was arbitrarily set to 1. Primers used for mRNA quantification are given in Supplementary Table S1A. ChIP-qPCR Amplification primer sequences are given in Supplementary Table S1E. Sequences or references of the siRNA are given in Supplementary Table S1D. Bars indicate SEM. The results of the 2-tailed Student unpaired t-test are indicated on the histograms. *P-value ≤0.05, **P≤0.01, ***P≤0.005, ****P≤0.0001

Figure 5

Effect of Fra-1 and Fra-2 depletion on p300/CBP recruitment at the HMGA1 locus and role of p300/CBP on HMGA1 expression. (A-B) Recruitment of p300/CBP at HMGA1 locus upon Fra-1 or Fra-2 depletion, respectively. ChIP-qPCR experiments were conducted on 72h after transfection of MDA-MB-231 cells with either siCTL (black bars), siFra-1 (blue bars) or siFra-2 (purple bars). The black arrow indicates HMGA1 TSS. Vertical red arrows indicate the Fra-1 and Fra-2 binding sites (Figure 3A). The values are the mean of 5 independent experiments and were normalized to that of amplicon +7.95 under control conditions, which was arbitrarily set to 1. Bars indicate SEM. The results of the 2-tailed Student unpaired t-test are indicated on the histograms. (C) Efficacy of RNAi against p300 or CBP. MDA-MB-231 cells were transfected with siCTL, sip300 or siCBP or the combination of sip300 and siCBP and protein samples were analyzed by immunoblotting with the indicated antibodies. Acetylation of histone H3 K27 was used to assess p300/CBP activity reduction. GAPDH was used as a loading control. (D and E) HMGA1 (D) and uPA/PLAU (E) mRNA levels upon depletion of p300 and/or CBP. MDA-MB-231 cells were transfected as in C and RT-qPCR assays of

HMGA1 and uPA/PLAU mRNAs were conducted (see supplementary Table S1A for primer sequences) and normalized to S26 mRNA taken as an internal control. Values are the mean of 5 independent experiments and were normalized to 1 in siCTL conditions. *P-value ≤0.05, **P≤0.01, ***P≤0.005, ****P≤0.0001.

Figure 6

Pol II, Pol II P-Ser5, Pol II P-Ser2 and H3K36me3 at the HMGA1 locus upon depletion of Fra-1 or Fra-2. (A-B) Pol II binding after depletion of Fra-1 or Fra-2, respectively. (C-D) Pol II P-Ser5 after depletion of Fra-1 or Fra-2, respectively. (E-F) Pol II P-Ser2 after depletion of Fra-1 or Fra-2, respectively. (G-H) H3K36me3 histone modification after depletion of Fra-1 or Fra-2, respectively. ChIP-qPCR experiments were conducted 72h after control (siCTL, black bars), anti-Fra-1 (siFra-1, blue bars) and anti-Fra-2 (siFra2, purple bars) siRNA transfection of MDA-MB-231 cells. HMGA1 gene TSS is indicated by a black arrow and Fra-1 and Fra-2 binding sites by red arrows. Relative signals were calculated with position -0.32 kb arbitrarily set to 1 in control conditions for Pol II and Pol II P-Ser5, with position +8.62 set to 1 for Pol II P-Ser2 and with position +3.23 set to 1 for H3K36me3. The data presented are the means of 4 independent experiments. Bars indicate SEM. The results of the 2-tailed unpaired Student t-test are indicated in histograms. Amplification primer sequences are given in Supplementary Table S1E. *P-value ≤0.05, **P≤0.01, ***P≤0.005, ****P≤0.0001.

Figure 7

3C analysis of the HMGA1 locus in the presence and in the absence of Fra-1. (A) Design of the 3C experiment. Position of the "anchor" reverse primer located in the promoter region (horizontal green arrow) and of the forward primers (horizontal blue arrows) located at the 3’ immediate proximity of DpnII restriction sites (grey arrows) are indicated. Purple arrows indicate the position of the reverse primers used to monitor the efficacy of the forward primers used for the 3C experiment analysis. All primer sequences and positions relative to TSS are given on Supplementary Table S1F. Doted lines (A1 to A14) indicate the possible interactions between the anchor and genomic regions according to the 3C experimental design. (B) Efficacy of the forward primers. Amplicons a1 to n14 were amplified by qPCR as described in Materials and Methods and analyzed by electrophoresis through a 2% agarose gel. (C and D) Independent replicates of 3C analyses of the HMGA1 locus in presence and absence of Fra-1. 3C libraries, after transfection of siCTL (C and D, upper panel) or siFra-1 (C and D, lower panels), were constructed as described in Methods. Possible interactions (A1 to A14) according to the 3C design were analyzed after qPCR on 2% agarose gels. The detection of DNA amplification

bands indicates tri-dimensional proximity between the anchor and the genomic positions defined by the DpnII sites.

Tolza et al. Figure 1

A Identity Cards of Tumours (CIT) Tumour Cancer Genome Atlas (TCGA) Affymetrix Data Bank Agilent Data Bank

ER- ER+ ER- ER+

basL mApo lumC lumB lumA normL basL Her2 normL lumC lumB lumA

mRNA mRNA 1 1 - - Fra Fra basL mApo lumC lumB lumA normL basL Her2 lumB lumA normL ER- ER+ ER- ER+ ER-

mRNA mRNA HMGA1 HMGA1 HMGA1 HMGA1 basL mApo lumC lumB lumA normL basL Her2 lumB lumA normL ER- ER+ ER- ER+ ER-

mRNA mRNA 2 2 - - Fra Fra

basL mApo lumC lumB lumA normL basL Her2 lumB lumA normL ER- ER+ ER- ER+ ER-

R2 = +0.419 R2 = +0.508 pV = 0.00E0 pV = 0.00E0 mRNA mRNA HMGA1 HMGA1 HMGA1 HMGA1

Fra-1 mRNA Fra-1 mRNA

R2 = -0.062 R2 = +0.111 pV = 0.199 pV = 1.04E-2 mRNA mRNA HMGA1 HMGA1 HMGA1

Fra-2 mRNA affymetrix profile Fra-2 mRNA Fra-2 mRNA

Tolza et al. Figure 2

MDA-MB-436 MCF-7 MDA-MB-231 8 1.5 12 levels

10 levels

levels 6 **** 1 8 6 4 mRNA mRNA mRNA 4 0.5 2 **** *** 2 **** **** **** **** **** Relative 0 **** Relative 0 0 Relative Relative

B Fra-1 Fos-DBD Fra-2 MDA-MB-231

15 1 2 1 2 1 2 ------

DBD) DBD) - MWM siCTL siFra siFra siCTL siFra siFra siCTL siFra siFra MWM MWM 10 (Fos 55 kd - Fra-2 Fra-1

35 kd - signals 5

35 kd - GAPDH

Relative Relative 0 MDA-MB-231

C D 436 231 - - Fra-1 HMGA1 Fra-2

1.5 1.5 1.5 MB MB 7 - - - levels

MDA MDA 1 1 1 MCF

*** Fra-1 mRNA *** **** 0.5 **** 0.5 0.5

**** **** Fra-2 0 0 0 Relative HMGA1

GAPDH

E MDA-MB-231 MDA-MB-436 F MDA-MB-231

1.5 1.5

1 2 1+2 1 2 1+2 ------mRNA 1 siCTL siFra siFra siCTL siFra siCTL siFra siFra siCTL siFra 1 *** ** Fra-1 0.5 0.5

Fra-2 0 0

Relative HMGA1 GAPDH

G MDA-MB-436 H MDA-MB-231

1.5 1.5 1.5

mRNA 1 1 1 mRNA ** ** 0.5 0.5 0.5 **** nascent HMGA1 Relative Relative HMGA1 0 0 0 HMGA1 Relative Relative HMGA1

Tolza et al. Figure 3

A +7.45 Kb +7.88 Kb TGAGTAA TGAGTCA TGACTCA -12.75 Kb -1.06 Kb +6.45 Kb TGAGTCA TGAGTCA TGAGTCA TGAGTCA Fra-1

Fra-2

RefGene (+) HMGA1

Coordinates 3.01 3.82 1.79 1.02 0.32 - - - - - 13.22 12.63 12.06 +0.05 +1.22 +1.89 +3.23 +4.16 +5.63 +6.52 +7.17 +7.56 +7.95 +8.62 +9.52 - - - RefGene (-) C6orf1

B MDA-MB-231 C H3

2 1.5 1.5

1.5 to HMGA1 1 1 1 level

0.5 0.5

0.5 Relative signal mRNA

**** **** 0 0 0 Relative 3.82 3.01 1.79 1.02 0.32 - - - - - +0.05 +1.22 +1.89 +3.23 +4.16 +5.63 +6.52 +7.17 +7.56 +7.95 +8.62 +9.52 HMGA1 C6orf1 13.22 12.63 12.06

HMGA1 C6orf1 - - -

D H3K4me3 E H3K4me1 1.5 2

1.5 1 1 0.5 0.5 Relative Relative signal Relative Relative signal

0 0 3.82 3.01 1.79 1.02 0.32 3.82 3.01 1.79 1.02 0.32 ------+0.05 +1.22 +1.89 +3.23 +4.16 +5.63 +6.52 +7.17 +7.56 +7.95 +8.62 +9.52 13.22 12.63 12.06 12.63 12.06 +0.05 +1.22 +1.89 +3.23 +4.16 +5.63 +6.52 +7.17 +7.56 +7.95 +8.62 +9.52 13.22 ------

F H3K27ac G p300/CBP 2.5 2.5

2 2 1.5 1.5 1 1

0.5 Relative signal 0.5 Relative Relative signal

0 0 3.82 3.01 1.79 1.02 0.32 3.82 3.01 1.79 1.02 0.32 ------+0.05 +1.22 +1.89 +3.23 +4.16 +5.63 +6.52 +7.17 +7.56 +7.95 +8.62 +9.52 13.22 12.63 12.06 +0.05 +1.22 +1.89 +3.23 +4.16 +5.63 +6.52 +7.17 +7.56 +7.95 +8.62 +9.52 13.22 12.63 12.06 ------

Tolza et al. Figure 4

A B C C MDA-MB-231 c-Jun Jun - 2.5 siCTL siJunB siJunD sic * 2.0

mRNA *** c-Jun * **** JunB 1.5 * * 1.0 JunD *** **** ** * GAPDH 0.5 HMGA1 relative relative HMGA1

MDA-MB-231 0.0 3.82 3.01 1.79 1.02 0.32 - - - - - 13.22 12.63 12.06 +0.05 +1.22 +1.89 +3.23 +4.16 +5.63 +6.52 +7.17 +7.56 +7.95 +8.62 +9.52 - - -

D E JunB JunD

* ** ***

** * **

3.82 3.01 1.79 1.02 0.32 3.82 3.01 1.79 1.02 0.32 ------13.22 12.63 12.06 +0.05 +1.22 +1.89 +3.23 +4.16 +5.63 +6.52 +7.17 +7.56 +7.95 +8.62 +9.52 13.22 12.63 12.06 +0.05 +1.22 +1.89 +3.23 +4.16 +5.63 +6.52 +7.17 +7.56 +7.95 +8.62 +9.52 ------

siCTL siFra-1

Tolza et al. Figure 5

A p300/CBP B p300/CBP

2.5 2.5

*** 2 2

1.5 1.5

1 **** 1 * *** 0.5 * *** 0.5 * Relative signal Relative signal 0 0

3.82 3.01 1.79 1.02 0.32 3.82 3.01 1.79 1.02 0.32 ------+0.05 +1.22 +1.89 +3.23 +4.16 +5.63 +6.52 +7.17 +7.56 +7.95 +8.62 +9.52 13.22 12.63 12.06 +0.05 +1.22 +1.89 +3.23 +4.16 +5.63 +6.52 +7.17 +7.56 +7.95 +8.62 +9.52 13.22 12.63 12.06 ------siCTL siFra-1 siCTL siFra-2

C D E

HMGA1 uPA/PLAU

1.5 1.5 1.5 1.5 mRNA siCTL sip300 siCBP siCTL sip300+siCBP mRNA * 1.0 1.0 1.0 *** 1.0 p300 250 kd **** CBP 250 kd 0.5 0.5 0.5 0.5 /PLAU Relative Relative /PLAU H3K27ac 15 kd 0 0 0 0 HMGA1 Relative Relative HMGA1 uPA GAPDH 35 kd MDA-MB-231

Tolza et al. Figure 6

A Total Pol II B Total Pol II 1.5 1.5

1 **** ** 1

0.5 0.5 ** ** * Relative signal 0 Relative signal 0

C Pol II P-Ser5 D Pol II P-Ser5 1.5 1.5

1 **** * 1

0.5 ** ** 0.5 * **** Relative signal Relative signal

0 0 3.82 3.01 1.79 1.02 0.32 3.82 3.01 1.79 1.02 0.32 ------13.22 12.63 12.06 +0.05 +1.22 +1.89 +3.23 +4.16 +5.63 +6.52 +7.17 +7.56 +7.95 +8.62 +9.52 13.22 12.63 12.06 +0.05 +1.22 +1.89 +3.23 +4.16 +5.63 +6.52 +7.17 +7.56 +7.95 +8.62 +9.52 ------

E Pol II P-Ser2 F Pol II P-Ser2 1.5 1.5

* **** 1 * 1 * 0.5 * ** 0.5 Relative signal Relative signal

G H3K36me3 H H3K36me3 2 2 * * 1.5 * 1.5 ** 1 **** 1

0.5 0.5 Relative signal Relative signal 0

3.82 3.01 1.79 1.02 0.32 3.82 3.01 1.79 1.02 0.32 ------13.22 12.63 12.06 13.22 12.63 12.06 +0.05 +1.22 +1.89 +3.23 +4.16 +5.63 +6.52 +7.17 +7.56 +7.95 +8.62 +9.52 +0.05 +1.22 +1.89 +3.23 +4.16 +5.63 +6.52 +7.17 +7.56 +7.95 +8.62 +9.52 ------

siCTL siFra-1 siCTL siFra-2

Tolza et al. Figure 7

A1 A2 A3 A4 A5 A6 A7 A8 A9 A10 A11 A12 A13 A14

Fra-1 Chip-seq RefGene (+) HMGA1

1 2 3 4 5 A 6 7 8 9 10 11 12 13 14 3C primers

DpnII sites 2.14 0.63 - - 15.49 14.11 12.03 +0.33 +1.98 +4.22 +6.94 +8.72 +9.91 - - - +11.72 +12.18 +13.42 Primers for 1 2 3 4 5 6 7 8 9 10 11 12 13 14 genomic DNA a b c d e f g h i J k l m n

B a1 b2 c3 d4 e5 f6 g7 h8 i9 j10 k11 l12 m13 n14

300bp Genomic

100bp DNA

C A1 A2 A3 A4 A5 A6 A7 A8 A9 A10 A11 A12 A13 A14

300bp siCTL 100bp 300bp siFra-1 100bp

D A1 A2 A3 A4 A5 A6 A7 A8 A9 A10 A11 A12 A13 A14 300pb siCTL 100pb

300pb siFra-1 100pb

AP-1 signaling by Fra-1 directly regulates HMGA1 oncogene transcription in Triple Negative Breast Cancers

Claire TOLZA, Fabienne BEJJANI, Emilie EVANNO, et al.

Mol Cancer Res Published OnlineFirst July 12, 2019.

Updated version Access the most recent version of this article at: doi:10.1158/1541-7786.MCR-19-0036

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