Interactions between BRD4S, LOXL2, and MED1 drive cell cycle transcription in triple‐negative breast cancer

Cell cycle; Gene expression; Triple-negative breast cancerCiclo celular; Expresión génica; Cáncer de mama triple negativoCicle cel·lular; Expressió gènica; Càncer de mama triple negatiuTriple‐negative breast cancer (TNBC) often develops resistance to single‐agent treatment, which can be circumvented using targeted combinatorial approaches. Here, we demonstrate that the simultaneous inhibition of LOXL2 and BRD4 synergistically limits TNBC proliferation in vitro and in vivo. Mechanistically, LOXL2 interacts in the nucleus with the short isoform of BRD4 (BRD4S), MED1, and the cell cycle transcriptional regulator B‐MyB. These interactions sustain the formation of BRD4 and MED1 nuclear transcriptional foci and control cell cycle progression at the gene expression level. The pharmacological co‐inhibition of LOXL2 and BRD4 reduces BRD4 nuclear foci, BRD4‐MED1 colocalization, and the transcription of cell cycle genes, thus suppressing TNBC cell proliferation. Targeting the interaction between BRD4S and LOXL2 could be a starting point for the development of new anticancer strategies for the treatment of TNBC.We thank the CRG genomics unit, the CRG‐UPF flow cytometry unit, and the VHIO mouse facility for their contribution. We thank Pharmaxis for the supply of PXS LOXL2 inhibitors. SS is supported by the Plan Estatal de I + D + I (COMBAT PID2019‐110598GA‐I00), and the ERC Starting Grant (ERC‐StG‐852343‐EPICAMENTE). LP‐R is supported by the Juan de la Cierva‐Formación fellowship (FJC2019‐040598‐I) and Fundación Franscico Cobos fellowship. TVT is supported by Plan Estatal de I + D + I (PID2019‐108008RJ‐I00), AECC (INVES20036TIAN), and a Ramón y Cajal investigator contract (RYC2020‐029098‐I). DC is supported by the la Caixa Foundation PhD fellowship (ID 100010434; fellowship code LCF/BQ/DI19/11730061)

Gene regulation by chromatin remodelling complexes : SWI/SNF complex in mRNA processing and B-WICH complex in ribosomal gene expression

The aim of this project is to investigate the roles of chromatin remodelling complexes in gene regulation. It is focused on two groups of chromatin complexes: the mammalian BRG1 and BRM SWI/SNF complexes and the ISWI-containing B-WICH complex. Study 1 investigates the role of SWI/SNF complexes in alternative splicing. We show that the presence of the ATPase core subunits Brg1 and Brm influence the alternative splicing outcome of a subset of genes. We show that Brg1 and Brm interact with several splicing related factors in the nascent RNA, and that the recruitment of some of these factors to their target sites is regulated by the presence of Brg1 and Brm. We propose that SWI/SNF ATPases can modulate the interactions of RNA binding factors to the nascent RNA and in that way alter alternative splicing outcome. Study 2 focuses on SWI/SNF complexes and their influence on cleavage and polyadenylation of mRNA. We show that Brg1 and Brm interact with subunits of the cleavage and polyadenylation complexes in the nascent mRNA. SWI/SNF complexes facilitate the recruitment of the cleavage and polyadenylation complex to the polyadenylation site in a subset of genes, and this results in a more efficient cleavage and polyadenylation. Study 3 shows that B-WICH is required for ribosome gene transcriptional activation upon glucose stimulation. WSTF and SNF2h, two of the B-WICH subunits, are needed to establish an active chromatin state in the RNA pol I gene promoter when the glucose concentration is raised after a period of deprivation. We propose that it counteracts the silent, poised chromatin state imposed by the silencing chromatin remodelling complex NuRD to allow for the RNA pol I machinery to bind to the promoter. These studies show that the influence of chromatin remodelling complexes upon gene expression is important for remodelling nucleosomes at the promoter, for alternative splicing, cleavage and polyadenylation and transcriptional initiation. These complexes work together with other chromatin remodelling factors, interact with other complexes and regulate their activity by affecting their recruitment dynamics.At the time of the doctoral defense, the following papers were unpublished and had a status as follows: Paper 1: Manuscript. Paper 3: Manuscript.</p

An Interaction between RRP6 and SU(VAR)3-9 Targets RRP6 to Heterochromatin and Contributes to Heterochromatin Maintenance in Drosophila melanogaster.

RNA surveillance factors are involved in heterochromatin regulation in yeast and plants, but less is known about the possible roles of ribonucleases in the heterochromatin of animal cells. Here we show that RRP6, one of the catalytic subunits of the exosome, is necessary for silencing heterochromatic repeats in the genome of Drosophila melanogaster. We show that a fraction of RRP6 is associated with heterochromatin, and the analysis of the RRP6 interaction network revealed physical links between RRP6 and the heterochromatin factors HP1a, SU(VAR)3-9 and RPD3. Moreover, genome-wide studies of RRP6 occupancy in cells depleted of SU(VAR)3-9 demonstrated that SU(VAR)3-9 contributes to the tethering of RRP6 to a subset of heterochromatic loci. Depletion of the exosome ribonucleases RRP6 and DIS3 stabilizes heterochromatic transcripts derived from transposons and repetitive sequences, and renders the heterochromatin less compact, as shown by micrococcal nuclease and proximity-ligation assays. Such depletion also increases the amount of HP1a bound to heterochromatic transcripts. Taken together, our results suggest that SU(VAR)3-9 targets RRP6 to a subset of heterochromatic loci where RRP6 degrades chromatin-associated non-coding RNAs in a process that is necessary to maintain the packaging of the heterochromatin

Genome-wide effects of RRP6 depletion on the transcriptome of S2 cells.

<p>The effects of RRP6 depletion on the steady-state expression levels were investigated by RNA-seq. Control experiments (GFP RNAi) were carried out in parallel and used as a reference. The expression levels in the control GFP cells and in RRP6-depleted cells expressed as reads per million (y-axis) are shown in green and orange, respectively. Genomic coordinates are indicated in the x-axis in B-C. (A) Pie diagram showing the effect of RRP6 depletion on the levels of different types of sequences, as indicated. (B) Examples of the effect of RRP6 depletion on the expression of repeat sequences. The upper and lower panels show subtelomeric regions of chromosome arms 2L and 3R, respectively, and the middle panel shows a region near the 2R centromere. (C) The effect of RRP6 depletion on the expression of selected transposon sequences. The genomic position of each sequence is indicated in the x-axis. (D) RNAi experiments were carried out to knock down RRP6 alone or RRP6 and DIS3 simultaneously. RNA was isolated and analysed by RT-qPCR using primer pairs designed to amplify selected sequences (the primer sequences are provided in the Supplementary Materials and Methods). The data was normalised to actin 5C RNA levels and expressed as a fold change compared to the levels observed in the GFP control. A protein-coding gene, <i>Pgk</i>, was analysed in parallel as a control. The histogram shows averages and standard deviations from three independent biological replicates. (E) ChIP experiments with antibodies against histone H3, H3K9ac, and H3K9me2 were carried out in untreated S2 cells to analyze the chromatin state of the selected genomic regions. Actin 5C was analyzed in parallel as a representative for euchromatin. The histogram shows averages and standard deviations from three independent biological replicates.</p

SU(VAR)3-9 depletion affects the association of RRP6 with chromatin.

<p>The association of selected proteins with the chromatin was analyzed by Western blotting using native chromatin preparations fractionated according to the scheme in <a href="http://www.plosgenetics.org/article/info:doi/10.1371/journal.pgen.1005523#pgen.1005523.g001" target="_blank">Fig 1F</a>. (A) Analysis of S2 cells that express the HA-tagged SU(VAR)3-9. The global levels of HP1a and SU(VAR)3-9 in the chromatin were analyzed in control cells (GFP) and in cells depleted of RRP6 and DIS3, or HP1a. The chromatin fractions were analysed using different antibodies, as indicated in the figure. An anti-HA antibody was used to detect SU(VAR)3-9. H3 and H3K9me2 served as controls. (B) Analysis of S2 cells that express the V5-tagged RRP6 depleted of HP1a. An anti-V5 antibody was used to detect RRP6 in the chromatin fractions. Depletion of HP1a does not affect the levels of RRP6 bound to the chromatin fraction. (C) Analysis of S2 cells that express the V5-tagged RRP6 depleted of SU(VAR)3-9. An anti-V5 antibody was used to detect RRP6. The quantification of the band intensities from three independent experiments is shown to the right. The standard deviations are given in parentheses. Histone H3 was used for normalization.</p

The catalytic activity of RRP6 is required for the silencing of transposon transcripts and for the maintenance of heterochromatin compaction.

<p>Wild-type RRP6-V5 or catalytically inactive mutants RRP6-Y361A-V5 and RRp6-D328A-V5 were expressed in S2 cells. Control cells that did not overexpress any protein were used in parallel for comparison. (A) Analysis of protein expression by Western blotting using an antibody against the V5 tag. Histone H3 served as loading control. (B) RT-qPCR analysis of transcript levels in cells that overexpress wither the wild-type RRP6-V5 or the catalytically inactive mutants. RNA was isolated and analyzed using primer pairs designed to amplify selected sequences, as indicated in the figure. The data was normalised to actin 5C mRNA levels and expressed as a fold change compared to the levels observed in the control cells (dark blue line). The histogram shows averages and standard deviations from three independent biological replicates. (C) PLA analysis of chromatin compaction using antibodies against HP1a and histone H3. The images show examples of PLA staining (magenta) in cells counterstained with DAPI (blue). The graph shows the number of PLA dots per cell in each condition. The mean number of dots per cell (magenta line) was 3,93 in the cells that overexpressed wild-type RRP6-V6 and 2,04 in cells that overexpressed RRP6-Y361A-V5. The difference was highly significant (P<0.0001; two-tailed Mann Whitney test; n = 150 cells analyzed in each condition, from two independent experiments).</p

Depletion of SU(VAR)3-9 influences RRP6 genomic occupancy in S2 cells.

<p>ChIP-seq experiments were carried out using S2 cells that expressed V5-tagged RRP6 under low-induction conditions. Chromatin preparations from control GFP cells and from cells depleted of SU(VAR)3-9 were used for ChIP-seq using an anti-V5 antibody. (A) Pie diagram showing the association of RRP6-rich regions with different types of sequences in control cells. (B) Chromosome distribution of RRP6 expressed in control cells as percentage of RRP6-rich regions in each chromosome (green bars). Two different scales are shown due to the lower fraction of regions in the heterochromatic scaffolds compared to the rest of the chromosome arms. The grey bars indicate the fraction of the genome corresponding to each chromosome, for comparison. (C) Depletion of SU-VAR)3-9 affects RRP6 genomic occupancy. For each chromosome or scaffold, the number of RRP6-rich regions upregulated or downregulated is expressed as percentage of the number of changed regions relative to the number of regions in that same chromosome in control cells. The percentage of affected regions is much higher in heterochromatin. (D) RRP6 occupancy in the genomic regions analyzed in <a href="http://www.plosgenetics.org/article/info:doi/10.1371/journal.pgen.1005523#pgen.1005523.g004" target="_blank">Fig 4</a>. The arrows indicate the regions amplified in the qPCR assays.</p

RRP6 is associated with heterochromatin <i>in vivo</i>.

<p>(A) Salivary gland polytene chromosomes immunostained with antibodies against RRP6 (green) and HP1a (red). The figure shows an overview micrograph. ch: chromocenter. (B) A detail showing a telomere stained with antibodies against RRP6 and HP1a, as in A. The fluorescence profile in the right part of the image shows the co-variation of both signals along the telomeric region. (C) A detail showing the chromocenter (ch) stained with antibodies against RRP6 and HP1a, as in A. (D) Co-localization of HP1a and RRP6 in dense chromatin in S2 cells analyzed by immuno-EM. An overview of a thin section through the nucleus of a cell is shown to the left. The bar represents 1 μm. A high-magnification micrograph shows co-localization of HP1a (12 nm gold) and RRP6 (6 nm gold) in the dense chromatin (<i>Dc</i>). The bar represents 100 nm. (E) The fractionation scheme used to isolate the different nuclear fractions in S2 cells: soluble (nucleoplasm), chromosomal RNP, and chromatin. (F) The distribution of HP1a, RPD3, and RRP6-V5 in the different nuclear fractions in S2 cells analyzed by Western blotting. The experiment was carried out in cells that expressed V5-tagged RRP6. Histone H3 was used as a control.</p

A metabolic map of the DNA damage response identifies PRDX1 in the control of nuclear ROS scavenging and aspartate availability

Data de publicació electrònica: 01-06-2023While cellular metabolism impacts the DNA damage response, a systematic understanding of the metabolic requirements that are crucial for DNA damage repair has yet to be achieved. Here, we investigate the metabolic enzymes and processes that are essential for the resolution of DNA damage. By integrating functional genomics with chromatin proteomics and metabolomics, we provide a detailed description of the interplay between cellular metabolism and the DNA damage response. Further analysis identified that Peroxiredoxin 1, PRDX1, contributes to the DNA damage repair. During the DNA damage response, PRDX1 translocates to the nucleus where it reduces DNA damage-induced nuclear reactive oxygen species. Moreover, PRDX1 loss lowers aspartate availability, which is required for the DNA damage-induced upregulation of de novo nucleotide synthesis. In the absence of PRDX1, cells accumulate replication stress and DNA damage, leading to proliferation defects that are exacerbated in the presence of etoposide, thus revealing a role for PRDX1 as a DNA damage surveillance factor.AM and CC were funded by the Austrian Science Fund (grant number P 33024 awarded to JIL). The Loizou lab is funded by an ERC Synergy Grant (DDREAMM Grant agreement ID: 855741). The Sdelci lab's contributions to this study were funded by an ERC Starting Grant (ERC-StG-852343-EPICAMENTE). This work was funded, in part, by a donation from Benjamin Landesmann. The funder was not involved in the study design, collection, analysis, interpretation of data, the writing of this article, or the decision to submit it for publication. CeMM is funded by the Austrian Academy of Sciences. MGVH acknowledges funding from R35CA242379, the Lustgarten Foundation, the Ludwig Center at MIT, and the MIT Center for Precision Cancer Medicine

Interactions between BRD4S, LOXL2, and MED1 drive cell cycle transcription in triple‐negative breast cancer

Abstract Triple‐negative breast cancer (TNBC) often develops resistance to single‐agent treatment, which can be circumvented using targeted combinatorial approaches. Here, we demonstrate that the simultaneous inhibition of LOXL2 and BRD4 synergistically limits TNBC proliferation in vitro and in vivo. Mechanistically, LOXL2 interacts in the nucleus with the short isoform of BRD4 (BRD4S), MED1, and the cell cycle transcriptional regulator B‐MyB. These interactions sustain the formation of BRD4 and MED1 nuclear transcriptional foci and control cell cycle progression at the gene expression level. The pharmacological co‐inhibition of LOXL2 and BRD4 reduces BRD4 nuclear foci, BRD4‐MED1 colocalization, and the transcription of cell cycle genes, thus suppressing TNBC cell proliferation. Targeting the interaction between BRD4S and LOXL2 could be a starting point for the development of new anticancer strategies for the treatment of TNBC