The nucleolus: Coordinating Stress Response and Genomic Stability

BBA - Gene Regulatory Mechanisms 1867 (2024) 195029 Available online 19 April 2024 1874-9399/© 2024 The Author. Published by Elsevier B.V. This is an open access article under the CC BY-NC-ND license (http://creativecommons.org/licenses/bync-nd/4.0/). Review The nucleolus: Coordinating stress response and genomic stability Katiuska Gonz´ alez-Arzola * Centro Andaluz de Biología Molecular y Medicina Regenerativa (CABIMER), Universidad de Sevilla, Consejo Superior de Investigaciones Científicas, Junta de Andalucía, Universidad Pablo de Olavide, 41092 Seville, Spain Departamento de Bioquímica Vegetal y Biología Molecular, Universidad de Sevilla, 41012 Seville, Spain ARTICLE INFO Keywords: Cellular stress coordination Nucleolar stress Nucleolus rDNA transcription Ribosome biogenesis ABSTRACT The perception that the nucleoli are merely the organelles where ribosome biogenesis occurs is challenged. Only around 30 % of nucleolar proteins are solely involved in producing ribosomes. Instead, the nucleolus plays a critical role in controlling protein trafficking during stress and, according to its dynamic nature, undergoes continuous protein exchange with nucleoplasm under various cellular stressors. Hence, the concept of nucleolar stress has evolved as cellular insults that disrupt the structure and function of the nucleolus. Considering the emerging role of this organelle in DNA repair and the fact that rDNAs are the most fragile genomic loci, therapies targeting the nucleoli are increasingly being developed. Besides, drugs that target ribosome synthesis and induce nucleolar stress can be used in cancer therapy. In contrast, agents that regulate nucleolar activity may be a potential treatment for neurodegeneration caused by abnormal protein accumulation in the nucleolus. Here, I Abbreviations: 3D FISH, tridimensional fluorescence in situ hybridization; A-bodies, amyloid bodies; APE1, apurinic/apyrimidinic endonuclease 1; ARF, alternative reading frame; ATM, ataxia telangiectasia mutated; ATR, ataxia telangiectasia and Rad3-related protein; BAX, BCL2-associated x protein; BER, base excision repair; BLM, Bloom’s syndrome protein; BRCA1, breast cancer type 1 susceptibility protein; CDK, cyclin-dependent kinase; CKII, casein kinase II; CLEM, correlative light and electron microscopy; CRISPR/Cas9, clustered regularly interspaced short palindromic repeats/caspase9; CRM1, chromosome region maintenance 1; CtBP, carboxy-terminal binding protein; CtIP, CtBP interacting protein; DDR, DNA damage response; diRNA, damage-induced small RNA; DJ, distal junction; DKC1, dyskerin pseudouridine synthase 1; DNA-PKcs, a catalytic subunit of DNA-dependent protein kinase; DRB, 5,6-dichloro-1-β-D-ribofuranosylbenzimidazole; DSB, double-strand break; E2F1, E2F transcription factor 1; EM, electron microscopy; ETS, external transcribed spacer; FBL, fibrillarin; GAR, Gly-Arg-rich domain; GLTSCR2, glioma tumorsuppressor candidate region gene 2; GNL2, guanine nucleotide binding protein-like 2; GNL3, guanine nucleotide binding protein-like 3; H3K27me3, histone H3 Lys27 trimethylation; H3K4me3, histone H3K4 trimethylation; H3K9me2/3, histone H3 Lys9 di/trimethylation; H4K20me2/3, histone H4 Lys20 di/trimethylation; HMG, high-mobility group; HMGB1, high mobility group box 1; HP1 α /β, heterochromatin protein 1 α /β; HR, homologous recombination; HS-AFM, high-speed atomic force microscopy; IDP, intrinsically disordered protein; IDR, intrinsically disordered region; IGS, intergenic spacer; IGS16RNA, lncRNA at 16 kb downstream of the rDNA TSS; IGS22RNA, lncRNA at 22 kb downstream of the rDNA TSS; IGS28RNA, lncRNA at 28 kb downstream of the rDNA TSS; INB, intranucleolar body; I-Ppol, Physarum polycephalum endonuclease I; ITS, internal transcribed spacer; K-rich, Lys-rich; lncRNA, long non-coding RNA; L-rRNA, large subunit rRNA; MDC1, mediator of DNA damage checkpoint protein 1; MKI67, marker of proliferation Ki-67; MLO, membrane-less organelle; MRE11, meiotic recombination 11 homolog 1; MRN, MRE11, RAD50 and NBS1 complex; NAC, NAM/ATAF/CUC transcription factor family; NAD, nucleolar associated domain; NBS1, Nijmegen breakage syndrome protein 1; NCL, nucleolin; nDDR, nucleolar DNA damage response; NER, nucleotide excision repair; NHEJ, non-homologous end joining; NHP2L1, chromosome protein 2-like 1; NOL10, nucleolar protein 10; NoLS, nucleolar localization sequence; NoRC, nucleolar remodeling complex; NPM, nucleophosmin; NuRD, nucleosome remodeling and deacetylation; PAPAS, promoter and pre-rRNA antisense; PARP1, Poly ADP-ribose polymerase 1; PCNA, proliferating cell nuclear antigen; PJ, proximal junction; PML, promyelocytic leukemia protein; POL-I/II/III, RNA polymerase-I/II/III; pre-rRNA, ribosomal RNA precursor; RAD50, radiation sensitive protein 50; RAD51/52, radiation sensitive protein 51/52; RBD, RNA binding domain; RBP, RNA binding protein; rDNA, ribosomal DNA; R-DPR, R-rich dipeptide-repeat protein; RIEP, ribosomal IGS encoded protein; RNF8, RNF8: ring finger protein 8; RP, ribosomal protein; RPA194, RNA polymerase-I subunit A; RPA2, replication protein A2; RPF2, ribosome production factor 2; RPL, ribosomal protein large; RPS, ribosomal protein small; RPSA, ribosomal protein SA; R-rich, arginine-rich; rRNA, ribosomal RNA; RRP1A, ribosomal RNA processing protein 1 homolog A; RRS1, ribosome biogenesis regulatory protein homolog; SL1, promoter selectivity factor; snoRNA, small nucleolar RNA; snoRNP, small nucleolar ribonucleoprotein; snRNP, small nuclear ribonucleoproteins; SRM, super-resolution microscopy; S-rRNA, small subunit rRNA; SUMO 1/2/3, small ubiquitin like modifier 1/2/3; SURF6, surfeit 6; TAF, TBP-associated factors; TBA, thrombin-binding aptamer; TBP, TATA-binding protein; TCOF1, Treacher Collins-Franceschetti syndrome; TFIIIB, transcription factor-IIIB; TIF-IA, transcription initiation factor-IA; TIP5, TTF-1-interacting protein-5; TOPO I/II, type I/II topoisomerase; TSS, transcription start site; TTF1, transcription termination factor 1; UBF, upstream binding factor; WRN, Werner syndrome helicase; XRCC1, X-ray cross complementing protein 1; XRCC4, X-ray cross complementing protein 4; ZTF-7, zinc finger putative transcription factor family 7. * Avenida Am´ erico Vespucio 24, 41092 Seville, Spain. E-mail address: [email protected]. Contents lists available at ScienceDirect BBA - Gene Regulatory Mechanisms journal homepage: www.elsevier.com/locate/bbagrm https://doi.org/10.1016/j.bbagrm.2024.195029 Received 15 November 2023; Received in revised form 25 March 2024; Accepted 12 April 2024 BBA - Gene Regulatory Mechanisms 1867 (2024) 195029 2 explore the roles of nucleoli beyond their ribosomal functions, highlighting the factors triggering nucleolar stress and their impact on genomic stability. 1. Introduction Floating in the chromatin solution, nucleoli are the most prominent and visible structures within the nucleus of eukaryotic cells. They are membrane-less organelles (MLOs), as are nuclear speckles, Cajal bodies, histone locus bodies, promyelocytic leukemia protein (PML) nuclear bodies, heterochromatin, the nuclear pore complex, among other assemblies existing in cell nuclei that contribute to compartmentalization of specific biological functions [1–3]. For many years, the sole function of the nucleolus was thought to be a factory for ribosomal RNA (rRNA) synthesis and assembly of ribosomal subunits [4]. Nevertheless, this organelle has gained attention over the last two decades as a central hub for protein trafficking control of critical cellular processes, e.g., cell cycle progression [5], tumorigenesis [6], and the cellular response to a large number of stressors [7], including DNA damage [8]. Nucleoli exhibit a highly dynamic structure that is disassembled and reassembled in each cell cycle [9]. Such a dynamic behavior is behind its lately ascribed role in the coordination of cellular response to injuries. Not in vain, different types of cellular stressful situations are often accompanied by dramatic changes in the organization and composition of the nucleolus [7]. Beyond the fact that nucleoli have been considered novel players in sensing and managing the cellular stress response, the concept of ‘nucleolar stress’ has been increasingly employed in recent years. This term is now used to refer to abnormalities in the structure and function of the nucleolus caused by stressful events that impair the ribosome biogenesis function and, consequently, affect cellular homeostasis [10]. Considering the growing role of the nucleolus in DNA repair and that ribosomal DNA (rDNA) genes, which reside in nucleoli, are especially unstable regions, nucleolar stress has been linked to several pathologies, ranging from cancer and neurodegenerative diseases to overall aging [11]. Consequently, therapies targeting nucleoli are vertiginously emerging. In this review, I revisit the roles of nucleoli beyond their ribosomal functions, emphasizing the factors that trigger nucleolar stress and their effects on gene stability regulation. 2. Nucleolar structure and functions Nucleoli encompass rDNA, diverse types of RNAs, and over 300 proteins [4]. These compartments were initially described as the sites for ribosome biogenesis, implicating the transcription of rDNA genes, rRNA maturation, and ribosomal subunit assembly [4]. Proteins and rRNAs involved in ribosome synthesis are highly concentrated in the nucleolus, creating a unique compartment or distinct body without a delimiting membrane. The high concentration of rRNAs comes from the transcription of the ribosomal genes, and these genes comprise about 60 % of total cellular transcription in eucaryotic cells, guaranteeing that the translational machinery meets the protein synthesis demand [12]. To increase rRNA synthesis and sustain the high demand for ribosomes, eukaryotic cell genomes keep a high copy number of their rRNA genes, varying from 100 to 10,000 across species [13]. Even though cells possess an elevated copy number of rDNA genes, only 50 % of them are transcribed; even metabolically active cells with a high demand of ribosomes keep a portion of their rDNA silenced, which has led some researchers to consider these genomic sites as sensors for DNA damage [14–16]. DNA methylation of cytosine residues in CpG dinucleotides is the principal epigenetic mechanism of silencing of rDNA genes [13,17]. In addition, repressive histone marks are often found in the promoter regions of silenced rDNA genes. These marks include histone H3 Lys27 trimethylation (H3K27me3), H3K9me3, as well as deacetylated histone H4 Lys16 (H4K16) and H4K20 [18,19]. On the contrary, active rDNA genes are enriched in active histone marks, i.e., H3K4 trimethylation (H3K4me3), H3 and H4 acetylation, and the histone variant H2A.Z [19]. Nucleoli form around transcriptionally active repeats of tandemly repeated rDNAs genes, a.k.a. nucleolar organizer regions (NORs Fig. 1) [20,21], resulting in a subnuclear compartment that concentrates the transcription and processing machinery responsible for rRNA synthesis, processing, and pre-ribosome subunit assembly [22]. In humans, the rDNA gene copies are positioned on the short arm of the acrocentric chromosomes (13, 14, 15, 21, and 22), between telomeric and centromeric heterochromatin, and distributed over the five chromosomes [21] (Fig. 1). rDNA gene array at NORs is flanked by two sequences named distal and proximal junctions (DJ and PJ, respectively, Fig. 1) that are conserved among the above chromosomes [23]. NORs can exist in either active or silent states. At active NORs, the rDNA copies are extensively complexed to the main rDNA transcription factor named upstream binding factor (UBF), a high-mobility group (HMG) box containing protein that stimulates rRNA synthesis. Thus, rDNA genes remain undercondensed. NORs are visible as secondary constrictions on chromosomes [24] (Fig. 1). On the contrary, transcriptionally silent or inactive NORs are not bound to UBF and, thus, entirely condensed [25]. 2.1. The nucleolus as a multilayer biomolecular condensate: the nucleolar sub-structure Growing evidence supports that the nucleolus represents a biomolecular condensate, a term increasingly used to describe MLOs, whose formation is driven by reversible liquid-liquid phase separation (LLPS) [26,27] similar to oil-in-water emulsions. Thus, nucleoli are dynamic assemblies with liquid-like properties whose tendency to fuse explains their broad size distribution [26]. The fusion of nucleoli is a highly active and dynamic process [28]. During interphase, the number of nucleoli gradually decreases as they fuse or associate with one another, causing the remaining nucleoli to become larger and denser [28]. Consequently, one or two nucleoli are typically observed during this period. However, at the beginning of cell division (prophase), the nucleolus begins to dissociate, and then nucleoli reappear at the end of mitosis (telophase) [28]. The maximum number of nucleoli per nucleus depends on the maximum number of NORs present in a genome, which is 10 in human cells [29]. The nucleolus’s highly dynamic architecture, which is disassembled and reassembled in each cell cycle around active NORs, results from the immiscibility of their different fluid-like components. Nucleoli consist of several RNAs complex to different ribonucleoproteins (RNA binding proteins or RBPs) involved in ribosome production and many other functions. It has been proposed that the liquid-like behavior of ribonucleoproteins enables the spontaneous formation of MLOs since purified nucleolar proteins phase separate into droplets resembling nucleoli [27]. Nevertheless, the representation of nucleoli as purely liquid-like compartments has been challenged. A study conducted by the Brangwynne team used quantitative imaging and mathematical modeling to investigate the properties of nucleolar material and their implication in the nucleolar form and function. Their findings suggest that the nucleolar shape is tuned by the constrained mobility of rRNA and mechanical constraints arising from the surrounding heterochromatin [30]. Therefore, nucleoli can be considered complex fluids with viscoelastic properties, exhibiting features of both solids and liquids [30]. The nucleolus contains separated layers or internal subcompartments, owing to differences in sequence-encoded biophysical properties of its macromolecule constituent, e.g., droplet surface tension [27]. The mammalian nucleolus is internally organized in at least four K. Gonz´ alez-Arzola BBA - Gene Regulatory Mechanisms 1867 (2024) 195029 3 concentric sub-compartments: the innermost structure is the fibrillar center (FC), followed by the dense fibrillar component (DFC), then by the granular component (GC), whereas the outermost layer corresponds to a ring of condensed perinucleolar chromatin (PC) (Fig. 2). The FC and DFC form functional units or modules with multiple copies immersed in a unique GC [31], and each FC-DFC unit contains two or three transcriptionally active rDNAs at the FC/DFC border [32] (Fig. 2). Interestingly, most eukaryotic nucleoli, including those in yeast, plants, all invertebrates, and some vertebrates, have only two main sublayers: a central fibrillar zone and a peripheral granular region [33,34]. It is believed that during evolution, the fibrillar zone of nucleoli separated into two domains, leading to the formation of the FC and the DFC [33]. Consequently, the differentiated FC sublayer emerged in reptiles at the transition between the anamniotes and amniotes, following a substantial increase in the size of the rDNA intergenic region [33,34]. A self-assembling scaffold protein, Treacher Collins- Franceschetti syndrome (TCOF1, a.k.a. Treacle), is thought to be responsible for FC formation and may provide insight into the emergence of the FC across evolution [35,36]. The multilayered nucleolar structure has been conserved in modern Eukaryotes. The emergence of additional nucleolar sub-compartments may have imparted regulatory functions to nucleoli, offering the potential for rapid responses to environmental stimuli through the selective sequestration of response factors [34]. Nucleolar sub-compartmentalization perfectly reflects the role of this organelle in the different steps of rRNA synthesis and processing, as well as pre-ribosomal subunit assembly, occurring during ribosome biogenesis (Fig. 3). FC holds the rDNA transcription machinery, including RNA polymerase I (POL-I), UBF, and promoter selectivity factor complex (SL1), among other factors [7,22]. DFC docks factors for the early stages of pre-rRNA processing, such as small nucleolar ribonucleoproteins (snoRNPs), small nucleolar RNAs (snoRNAs), fibrillarin (FBL), and others [7,22]. As nascent rRNAs emerge from the border between FC and DFC (where POL-I is most abundant), they bind to FBL [31]. FBL is a selfassociated RBP responsible for the early processing and methylation of rRNAs [37], which is endowed with an N-terminal glycine- and argininerich (GAR) domain that contains several intrinsically disordered regions (IDRs) and a C-terminal methyltransferase region which, in turn, comprises an RNA binding domain (RBD) [32]. The binding of the newly synthesized pre-rRNAs to the C-terminal RBD of FBL (along with the selfassembly tendency of this latter via its GAR domains) promotes FBL phase separation, the establishment of the DFC phase, and the prerRNAs sorting into the DFC to initiate their processing [32]. Once the polycistronic pre-rRNA cleavages into 28S, 18S, and 5.8S rRNAs, these Fig. 1. Schematic representation of rDNA genes in mammalian cells. The rDNA gene array is located at the nucleolar organizer regions (NORs). It is positioned between telomeres and centromeres on the short arms (p-arms) of the acrocentric chromosomes 13, 14, 15, 21, and 22 in humans. They are flanked by heterochromatic distal and proximal junctions (DJ and PJ, respectively). NORs can be observed as achromatic gaps on chromosomes due to reduced condensation of rDNA. NORs are situated between telomeric and centromeric heterochromatin. Each rDNA repeat unit (~43 kb) consists of a transcribed region (encoding the 45S pre-mRNA for 18S, 5.8S, and 28S mature rRNAs) and an intergenic spacer (IGS). The rDNA transcribed region in humans is 13 kb, while the IGS region has 30 kb. The coding region is flanked by external transcribed spacers (5 ′ ETS and 3 ′ ETS), and rDNA genes are separated by internal transcribed spacers (ITS1 and ITS2). A transcription termination site is located downstream of the transcribed region. The IGS comprises a pre-rRNA (gene) promoter and a spacer promoter. Distal to the spacer promoter are situated enhancer elements. The IGS region may contain nucleosomes. Fig. 2. Internal organization of mammalian nucleoli. Quadripartite structure of a mature nucleolus in interphase: fibrillar center (FC, dark blue), dense fibrillar component (DFC, medium blue), granular component (GC, light blue), and perinucleolar chromatin (PC, gray). The FC-DFC modules are present in several dozen copies in humans and are immersed in a single GC sub-compartment. Each FC-DFC bipartite structure contains two or three transcriptionally active rDNAs (orange) at the FC/DFC interface. Distal junction (DJ) sequences locate at the PC nucleolar layer. (For interpretation of the references to colour in this figure legend, the reader is referred to the web version of this article.) K. Gonz´ alez-Arzola BBA - Gene Regulatory Mechanisms 1867 (2024) 195029 4 latter are modified by snoRNPs and additional processing factors [22]. Although the RBD of FBL is situated inside its methyltransferase region, the methylation function of this latter domain is not relevant for prerRNA sorting [32]. Super-resolution electron microscopy analyses recently revealed that FBL and other ribosome assembly factors form a network of 18–24 regularly spaced clusters or mini condensates into the DFC [31,32]. Among these factors are non-histone chromosome protein 2-like 1 (NHP2L1) and dyskerin pseudouridine synthase 1 (DKC1) [32]. The significance of such nanosized clusters inside the DFC is unknown, but their presence suggests compartmentalizing the different RNA processing reaction steps. The outer GC encloses the FC-DFC units, and it is the place where the late stages of ribosome synthesis take place, i.e., the late rRNA processing and the pre-ribosomal subunit assembly. The 5.8S and 28S rRNAs in the GC assemble with the 5S rRNA and ribosomal proteins (RPs) to form the 60S subunit. In contrast, the 18S rRNA and other RPs build the 40S ribosome subunit [22] (Fig. 3). Nucleophosmin (NPM, a.k.a. NPM1 and B23) is an abundant histone chaperone residing in the GC that stimulates the rDNA transcription by regulating the histone density around the rDNA genes [38]. Besides its role in the nuclear export of the 60S and 40S ribosomal subunits [39], NPM has lately been assigned a role in separating nascent 60S particles from the chromatin fraction of the FC-DFC module towards GC, where further maturation proceeds [40]. This chaperone encompasses a structured N-terminal oligomerization domain, a middle-disordered region responsible for histone chaperoning, and a folded C-terminal nucleic acid binding domain [41]. In addition to ribosome synthesis, mRNA transport, chromatin remodeling, and DNA repair [41], NPM regulates the alternative reading frame (ARF)-p53 tumor suppression pathway [42,43]. Pioneering biophysical approaches performed by Kriwacki’s group demonstrated that NPM dramatically contributes to the nucleolar sub-structure using its ability of LLPS [44,45]. To drive phase separation, NPM displays a succession of three acidic and two basic tracts, most located in the middle IDR and one in the pentameric domain, providing a solid interaction multivalency. Thus, multivalent interactions of NPM with nucleolar proteins containing arginine-rich (R-rich) linear motifs and rRNA confer the liquid-like feature to the GC [44]. A recent study found that in addition to R-motifs, lysine-rich (K-rich) proteins also trigger NPM phase transitions to regulate the trafficking of nucleolar proteins [43]. Another abundant protein in the GC is nucleolin (NCL), a histone chaperone that facilitates rDNA transcription by promoting a euchromatic state [46]. NCL consists of three acidic stretches at the aminoterminal end, four RBDs, and a carboxy-terminal GAR domain [47]. According to a recent investigation, the GAR domain of NCL drives its subcellular localization in the axon of neurons and the plasma membrane of tumor cells, thus contributing to placing NCL within the nucleolus [48]. Considering the involvement of the GAR domains of FBL in the phase transitions at the nucleolar DFC, it is tempting to hypothesize a similar role for the GAR domain of NCL in the GC, in addition to the phase transitions accomplished by NPM. The nature and exact cellular function of the fourth nucleolar layer (PC) is still controversial. 3D fluorescence in situ hybridization (3D FISH) analysis has led to the proposal that PC concentrates inactive chromosomal regions enriched in genes involved in response to other organisms (defensin, immunoglobulin, and T-cell receptor genes), sensory perception (olfactory genes), tissue development and embryo implantation, as well as a large number of snoRNA genes, 5S rDNA and tRNA genes [49]. Such gene families positioned at PC are arranged in large gene clusters and may be tissue-specific expressed [49]. According to the apparent inactive chromatin structure of PC, several repressive Fig. 3. Nucleolar organization and its function in ribosome biogenesis. The overall structure of the mammalian nucleolus shows the internal fibrillar center (FC), a dense fibrillar component (DFC), and an external granular component (GC), all of them surrounded by condensed perinucleolar chromatin (PC). RNA polymerase-I (POL-I) transcribes the rDNA genes to form a 47S pre-rRNA at the FCDFC border. A pre-initiation complex formed by POL-I, transcription factor upstream binding factor (UBF), the promoter selectivity factor complex (SL1), transcription initiation factor-IA (TIF-IA), and Treacle is required for rDNA cluster transcription. Type-I/II topoisomerases (TOPO I/II) action is also needed to initiate rDNA transcription. In the DFC, pre-rRNA is further processed in mature 18S, 5.8S, and 28S rRNAs. DFC harbors small nucleolar RNAs (snoRNAs), small nucleolar ribonucleoproteins (snoRNPs), fibrillarin (FBL), and other factors involved in pre-rRNA processing. In the nucleoplasm, the 5S rDNA and the ribosomal proteins (RPs) genes are transcribed by RNA polymerase III (POL-III) and RNA polymerase II (POL-II), respectively. In the GC, RPs associate with rRNAs to form the ribosomal subunits. IGS: intergenic spacer. K. Gonz´ alez-Arzola BBA - Gene Regulatory Mechanisms 1867 (2024) 195029 5 histone marks can be detected (H3K27me3, H3K9me3, and histone H4 Lys20 trimethylation or H4K20me3), whereas active histone marks, such as H3K4me3, are nearly depleted at these regions [49]. Although the chromatin adjacent to nucleoli, the so-called nucleolar-associated domains or NADs, displays heterochromatic features, other nucleolar genome organization landscapes cannot be ruled out due to methodological limitations of NAD identification [50]. Curiously, one region located to the periphery of the PC is flanking the NORs, the DJ [23] (Fig. 2). These sequences positioned in the PC display transcriptionally active chromatin features, thus suggesting a role for DJs in nucleolar formation and functioning [51–53]. DJ regions also organize the rDNA array when it is damaged, implying their participation in rDNA gene stability, as mentioned below. Spatial mapping of the nucleolar proteome revealed that some nucleolar proteins anchor the nucleolus to the PC and form a “nucleolar rim” [54]. The nucleolar rim holds several proteins, such as marker of proliferation Ki-67 (MKI67), which is needed for proper chromosome segregation during mitosis, and guanine nucleotide binding protein-like 3 (GNL3) that is required for cell proliferation, among other factors [54]. The quadripartite structure of the nucleolus is visible by electron microscopy (EM), but the other lately described nucleolar sub-structures are not easily observed by EM; these include the periphery of DFC (involved in the maturation of rRNAs [36,55]) and the nucleolar rim whose biochemical function remains a mystery. Until recently, it was believed that NPM and NCL in the GC and FBL in the DFC were the only nucleolar proteins strongly implicated in nucleolar assembly; however, other factors may contribute to the nucleolar architecture. An integrative software approach calculates the iNo score, determining the degree of nucleolar disruption after depleting specific nucleolar proteins [56]. This analysis determined that very few RPs are required to maintain the nucleolar structure, and only lateassembled RPs of the 60S large ribosomal subunit significantly contributed to nucleolar structure [56]. Among 60S RPs, ribosomal protein large 5 (RPL5, a.k.a. uL18) and RPL11 (a.k.a. uL5) are essential to nucleolar structure maintenance [56–58]. Curiously, RPL5 and RPL11 are necessary for activating the p53 pathway upon disruption of ribosome biogenesis [59]. RPL5 and other R-rich RPs, such as guanine nucleotide binding protein-like 2 (GNL2), surfeit 6 (SURF6) and RPL23 (a.k.a. uL14), display liquid phase transitions with NPM [44]. Remarkably, a new investigation has shown that heterochromatic proteins, particularly heterochromatin protein 1 α and β (HP1 α and HP1β), ensure the structural integrity of nucleoli [60]. This agrees that HP1 α triggers LLPS-driven heterochromatin formation [61,62]. A novel superresolution microscopy (SRM) analysis revealed that nucleolar protein 10 (NOL10) constitutes the “skeletal structure” of the nucleolar GC and that this protein is indispensable to maintaining the structure of the nucleolus [63]. Recent advancements in techniques such as SRM and correlative light and electron microscopy (CLEM) have enabled the imaging of individual proteins within nucleolar sub-compartments [63]. Furthermore, the use of high-speed atomic force microscopy (HS-AFM) has allowed the dynamic imaging of the globular regions and the unstructured tails of nucleolar intrinsically disordered proteins (IDPs) [63]. In the future, the application of high-resolution techniques is expected to further improve our understanding of the sub-structure of the nucleolus. Besides the participation of phase-separated proteins in nucleolar structure, it has been recently pointed out that certain rRNAs contribute to this process, suggesting that nucleolar architecture is coordinated with rRNA processing and maturation [64]. A specific rRNA intermediate, 27SA 2 , is crucial in maintaining the spherical shape of nucleoli in C. elegans [64]. 2.2. Ribosome synthesis: the canonical nucleolar role Although ribosome synthesis is preserved across all domains of life, the presence of a nucleolus has been only documented in Eukaryotes and, most recently, in Archaea [65]. In the nucleolus, POL-I transcribes the rDNA genes encoding the small subunit S-rRNA (18S) and the large subunit 5.8S and 28S L-rRNAs as a single polycistronic precursor (47S pre-rRNA). Each rDNA repeat unit at NORs consists of a pre-rRNA coding region and an intergenic spacer (IGS). The pre-rRNA coding region generates the 47S pre-rRNA, which is further processed into three mature rRNA molecules: 18S, 5.8S, and 28S (Fig. 1). The rDNA coding sequences are highly conserved across species. Still, the IGS region is less conserved and contains regulatory elements, such as two promoters and several repetitive enhancer elements [66] (Fig. 1). At the IGS, the gene promoter controls the polycistronic 47S pre-rRNA transcription. In contrast, the spacer promoter encodes non-coding RNAs likely involved in gene silencing [66]. Notably, long non-coding RNAs (lncRNAs) are produced from the IGSs of rDNA genes, and they seem to have regulatory roles in responding to environmental stressors, as described below. Thus, these nucleolar lncRNAs allow cellular adaptation to stress by reducing the intense transcription of rDNA genes and causing protein sequestration within the nucleolus [67]. POL-I transcription requires the formation of a preinitiation complex on the pre-rRNA promoter, including binding the nucleolar transcription factor UBF and SL1, among others [13]. The transcription initiation factor IA (TIF-IA; a.k.a. RRN3) interacts with POL-I, enabling the POL-I binding to SL1 [68]. SL1 complex contains the TATA-binding protein (TBP) and five TBP-associated factors: TAF1A, TAF1B, TAF1C, TAF1D and TAF12 [69]. Another vital transcription factor is UBF, which displaces histone H1 and maintains an open chromatin configuration susceptible to binding of additional factors [70]. Several other factors are associated with the preinitiation complex to initiate transcription, as is the case of TOPO I and II, which remove DNA supercoils at the rDNA promoter [71,72], and Treacle that enables the recruitment of POL-I and UBF to the pre-initiation complex [73]. Transcription termination factor 1 (TTF1) controls transcription termination and stops elongation by POL-I. Genes encoding the 5S rRNA are also present in tandem repeats, but in a separate genomic location, on the distal end of chromosome 1 [66]. These genes are transcribed by RNA polymerase-III (POL-III) outside of the nucleolus [66]. Incorporation of the 5S rRNA into the preribosome is poorly understood. A recent study has proposed that nascent 5S rRNA binds to RPL5, RPL11, and other proteins within the nucleoplasm [74]. The 5S ribonucleoprotein complex is then recruited by nucleolar factors (ribosome production factor 2 or RPF2, and ribosome biogenesis regulatory protein homolog or RRS1) to form a hexameric complex that is ultimately incorporated into the pre-60S subunit [74]. In addition to removing the transcribed spacer regions through endonucleolytic and exonucleolytic cleavages, rRNAs are submitted to further modifications in the nucleolus (methylation and pseudouridylation) by snoRNPs [75]. The nascent pre-rRNAs associate cotranscriptionally with some ribosomal proteins and snoRNPs to form large particles in which pre-rRNAs are folded and modified to yield the mature rRNAs [76]. In the nucleoplasm, RNA polymerase-II (POL-II) transcribes the genes encoding ribosomal proteins (RPs) and assembly factors, which are translated into the cytoplasm and reimported to the nucleus for assembly. Most RPs are transported to the nucleus by multiple karyopherins or importins, which recognize the nuclear localization signals of their cargo proteins and carry them through the nuclear pore complex [77]. Additionally, NCL and NPM assist in targeting RPs to the nucleolus [78,79]. POL-II also transcribes about 200 snoRNAs in the nucleoplasm, participating in pre-RNA processing [80]. SnoRNAs associate with nucleolar proteins 56 and 58 (NOP56 and NOP58), FBL, among others, which prompts their nucleolar targeting [81]. Interestingly, chromosome region maintenance 1 (CRM1) promotes the nucleolar targeting of snoRNAs via the exposition of the nucleolar localization sequence (NoLS) of NOP58 [81]. Later, the 80 RPs that form the 60S large and 40S small ribosomal subunits, 47 and 33 RPs, respectively [49], enter the nucleolus and associate with the mature rRNAs to generate the precursors of the K. Gonz´ alez-Arzola BBA - Gene Regulatory Mechanisms 1867 (2024) 195029 6 ribosomal subunits [82]. These are then exported to the cytoplasm through the nuclear pore complex, forming a mature ribosome to initiate protein synthesis (Fig. 3). Intriguingly, a latest report stated that a nucleolar pool of active POL-II binds to the IGS regions of rDNA genes to drive their expression [83]. Nucleoli are wholly disassembled by the end of prophase to prepare cell division, segregating the daughter cells’ cellular components. Nucleolar disassembly initiates when rDNA transcription is abolished through cyclin-dependent kinase 1 (CDK1)-mediated POL-I inactivation [84]. The nucleolus formation occurs at the end of mitosis (in telophase) when pre-nucleolar bodies containing FBL, NPM, and ribosomal RNA processing protein 1 homolog A (RRP1A a.k.a. NOP52) are recruited to NORs [85]. The latter arises when rDNA transcription resumes due to decreased CDK1 activity, leading to further re-activation of POL-I [65]. During interphase, multiple nucleoli fuse to form larger nucleoli that contain several NORs and are surrounded by heterochromatin [52]. 3. Nucleolar stress 3.1. The nucleolus as a central hub for cellular stress response coordination In recent decades, various investigations have demonstrated that the nucleolus is an organelle with multiple functions. Although ribosome biogenesis is the traditional primary function of the nucleolus, only ~30 % of nucleolar proteins are associated with ribogenesis [7]. Various proteomic studies have identified nucleolar factors involved in many other mammalian processes, i.e., cell cycle progression, proliferation, apoptosis, and the response to different insults [86–88]. The varied functions of over 4500 nucleolus-associated proteins, according to the latest update of the Nucleolar Proteome Database (NOPdb3.0), are consistent with non-canonical additional roles for the nucleolus [86]. The nucleolus senses several types of cellular stress, which turns this organelle into a central hub for coordinating the subsequent response [7]. Due to the absence of a delimiting membrane, the nucleoli composition is highly variable. Agreeing to such a dynamic nature, the nucleolus is an ideal location for rapid cellular homeostasis regulation through the transient sequestration of critical regulators of the cellular stress response, which often results in nucleolar changes in size, number, and protein composition on demand [11]. Importantly, given that ribosome biogenesis is directly coupled with cell proliferation, it is not surprising that nucleoli play a crucial role in orchestrating primary cellular responses to stress. Thus, in accordance with their fluid nature, nucleolar components exhibit continuous flux within the nucleolus and are exchanged with the surrounding nucleoplasm under various stress situations. The link between cellular stress and nucleoli comes from the notion that these organelles directly regulate the stress-responsive factor par excellence, p53 [89,90]. Expectedly, a common feature in many p53- mediated cellular responses is nucleolar structure disorganization [10,90]. p53 is a master transcription factor predominantly regulated by proteasomal degradation mediated by its ubiquitin ligase murine double minute 2 (MDM2, HDM2 in humans) [91]. Several kinds of cellular injuries induce redistribution to the nucleoplasm of nucleolar proteins to stabilize p53, which further induces cell cycle arrest or apoptosis by controlling dozens of target genes [92,93]. These nucleolar proteins include NPM [93], RPL5 [94], RPL6 [95], RPL11 [96], RPL23 [97], ribosomal protein small 3 (RPS3) [98], RPS14 [99], RPS27-like [100] and glioma tumor-suppressor candidate region gene 2 (GLTSCR2) [101]. The above-mentioned nucleolar proteins are released from the nucleoli to inhibit MDM2, thus leading to p53 stabilization [93–101]. All this reinforces the idea that nucleolar proteins activate p53 and that the nucleolus is a sensor for cellular stress. Despite this, growing evidence suggests that p53 is not the only stress-responsive factor activated by the nucleolus [102]. This is particularly important because many cancer cells contain mutant p53 [103]. Such p53-independent mechanisms also involve ribosomal proteins and other nucleolar factors such as NPM and ARF [102]. When released from nucleoli, RPL5, RPL11, and RPS14 inhibit the activity of the transcriptional factor c-Myc, thus preventing the c-Myc-mediated activation of genes involved in cell grown and proliferation [104,105]. Moreover, NPM and ARF translocate from the nucleolus and bypass p53 to directly regulate apoptosis. Following DNA injuries, NPM relocates to the cytoplasm to sequester BCL2-associated x protein (BAX), thus avoiding the pro-apoptotic effect of BAX [106]. In response to genotoxic stress, ARF activates ataxia telangiectasia mutated (ATM)/ataxia telangiectasia and Rad3-related protein (ATR) signaling pathways, thus inducing p53-independent cell cycle arrest [107]. On the other hand, other transcription factors, such as the plant-specific family called NAM/ATAF/CUC (NAC), respond to nucleolar stress caused by perturbations of ribosome biogenesis in plants, which lack p53 [108]. Remarkably, some biomolecular condensates are formed within the nucleolar space and play a crucial role in protein sequestration during stressful conditions [109]. Some are aggregates similar to amyloid plaques, a.k.a. amyloid bodies or A-bodies, originating from lncRNAs transcribed from the ribosomal IGSs [110]. A-bodies are induced in response to various environmental stimuli such as heat shock, acidosis, and other stressful situations [109,111]. During stress, clusters of lncRNAs with dinucleotide repeats (CU or AG) accumulate in the nucleolar area. The low-complexity sequences of lncRNAs facilitate charge-based interactions with cationic proteins to produce nucleolar liquid-like foci which sequester these proteins away from their downstream effectors [110]. Heat shock triggers the formation of lncRNAs located 16 kb (IGS16RNA) and 22 kb (IGS22RNA) downstream of the rDNA transcription start site (TSS) [112]. In contrast, acidosis causes the accumulation of a transcript located 28 kb (IGS28RNA) downstream of the TSS [112]. When the proteasome activity in mammalian cells is hampered, it causes the accumulation of polyubiquitylated proteins. This leads to the formation of RNA-protein aggregates in the nucleolar space that are known as nucleolar aggresomes [113,114]. These subnucleolar structures contain polyadenylated RNA, ubiquitin, and several nucleoplasmic proteasome target proteins. The latter includes key factors in oncogenesis, cell cycle-regulating cyclins and cyclindependent kinases (CDKs), and stress response kinases [114]. There is another class of sub-nucleolar structures called intranucleolar bodies (INBs). INBs are found in abundance in unstressed nucleoli and are located near rDNA with heterochromatic features [115]. They appear to regulate the transcription and maintenance of rDNA genes. INB formation is enhanced by specific DNA-damaging conditions like infrared irradiation and etoposide or camptothecin treatment [115]. However, these nucleolar aggregates do not disrupt the structural integrity of the nucleolus. INBs contain proteins that are involved in DNA repair and replication, protein turnover, RNA processing, chromatin organization, as well small ubiquitin-like modifier 1 (SUMO 1) and SUMO 2/3 [115]. The close relationship between the nucleolus and stress was described in the middle of the last century when nucleoli were selectively irradiated with ultraviolet light, and cell cycle arrest immediately occurred [116]. Since, at that time, the only known function of the nucleolus was the synthesis of ribosomes, this led to the hypothesis that deficiency in ribosomes or the ribosomal functioning is the cause of cell cycle arrest. Consequently, the term “nucleolar stress” was traditionally used to refer to “ribotoxic stress” or “ribosome biogenesis stress.” Nucleolar stress has just emerged as a novel concept, and it is now referred to as diverse cellular insults that induce abnormalities in nucleolar structure and function [6,10]. As pointed out before, many stressors display a nucleolar response leading to the initiation of the p53-dependent cell cycle arrest or apoptosis, depending on the severity of the injuries. Nevertheless, a report suggests that p53-deficient U2OS osteosarcoma and HCT-116 colon cancer cells are also sensitive to the nucleolar stress induced by rDNA inhibition [117]. In such cases, cell cycle arrest relies on the degradation of E2F transcription factor 1 (E2F1) [117]. In addition, p53- K. Gonz´ alez-Arzola BBA - Gene Regulatory Mechanisms 1867 (2024) 195029 7 knockout erythroid cells undergoing defects in ribosome synthesis were able to block cell cycle progression due to the stabilization of the cell cycle inhibitor p27 [118]. In contrast, cells lacking p53 protein expression submitted to disruption of ribosome biogenesis displayed G1/S arrest or apoptosis due to p53-independent activation of p21 expression [119]. In addition to activating p21, nucleolar stress triggers additional mechanisms that lead to cell cycle arrest. One such mechanism involves the translocation of ribosome protein large 3 (RPL3, a.k.a. uL3) into the nucleoplasm following a genotoxic treatment [120]. There, RPL3 binds to poly ADP-ribose polymerase 1 (PARP-1), thus preventing the binding of the latter to the E2F1 promoter and avoiding the transcription factor activation. As E2F1 is responsible for activating the transcription of genes required for entry into the late G1/S phase, this leads to cell cycle arrest [120]. Moreover, RPL3 negatively regulates cyclin D1 by reducing its mRNA and protein levels, thus inhibiting the progression through the G1 phase [120]. Strikingly, RPL3 expression is downregulated in colon cancer cells [121], which makes some chemotherapeutic agents ineffective [122]. Moreover, RPL3 acts as an inhibitor of the autophagic process in colon tumor tissues [122]. Autophagy has been linked to both tumorigenesis and chemotherapeutic resistance [123]. Therefore, RPL3 improves the activity of many anticancer drugs thanks to its pro-apoptotic and anti-autophagic activities in p53 deficient cancer cells [122]. These reports suggest that p53-independent mechanisms trigger cell cycle progression blocking or cell death under nucleolar alterations. The p53-dependent and -independent responses to nucleolar alterations are extensively reviewed in [5,124]. Hence, a more precise description of nucleolar stress can be annotated as stressful cellular events that impact nucleolar morphology and functions, ultimately activating p53 or other stress signaling pathways to induce cell cycle arrest or cell death. According to Yang et al., the factors inducing nucleolar stress can be classified as canonical and non-canonical [10]. The formers impair or alter any step of the ribosome biogenesis; thus, canonical nucleolar stress inducers are often POL-I inhibitors [125] or factors inducing aberrant expression of nucleolar proteins [126]. On the other hand, the non-canonical signals include various general cellular insults that affect non-ribosomal functions, e.g., UV light, heat shock, hypoxia, free radicals, nutrient deprivation, and many others [10]. In any case, nucleolar stress is often reflected in drastic changes in the structure and composition of the nucleolus (Table 1). 3.2. Canonical nucleolar stress Cellular stressors that alter any of the multiple steps of ribosome biogenesis may lead to canonical nucleolar stress, reviewed below (Fig. 4). 3.2.1. rDNA transcription inhibition A well-described example of canonical nucleolar stress is the nucleolar segregation induced by actinomycin D. This drug intercalates with both double and single-stranded DNA, mainly at GC-rich regions which are present at a high frequency in rDNA genes, thus pausing POL-I and blocking rDNA transcription elongation [131]. Noteworthy is that actinomycin D selectively inhibits POL-I and induces nucleolar stress at a low concentration (5 nM). In contrast, high doses (>30 nM) may also produce DNA damage and nuclear transcription inhibition [161]. Other intercalating agents affecting nucleolar structure are doxorubicin and mitoxantrone. These compounds establish covalent complexes between DNA and type II topoisomerases, inhibiting the latter and preventing the rDNA transcription initiation by POL-I [129,130]. Mitomycin C is another intercalating drug that inhibits rDNA transcription by an unknown mechanism [131]. Methotrexate is a folate analog that inhibits the synthesis of purine and pyrimidine, thereby impeding the transcription carried out by POL-I [131,132]. The platinum compounds oxaliplatin and cisplatin are extensively used cancer chemotherapeutic agents, and they have been proposed to Table 1 Effects of canonical and non-canonical signals triggering nucleolar stress. Stress type Cause Effect on nucleolar structure Ref. Inhibition of rDNA transcription Actinomycin D (low dose) Nucleolar caps, translocation of nucleolar proteins to nucleoplasm [127,128] Doxorubicin Nucleolar caps, translocation of nucleolar proteins to nucleoplasm [129,130] Mitoxantrone Nucleolar caps [129,130] Mitomycin C Nucleolar caps [131] Methotrexate Nucleolar caps, translocation of nucleolar proteins to nucleoplasm [131,132] Oxaliplatin Nucleolar caps, translocation of nucleolar proteins to nucleoplasm [132] Cisplatin (high dose) Nucleolar caps, translocation of nucleolar proteins to the nucleoplasm [132,133] BMH-21 Nucleolar caps, translocation of nucleolar proteins to nucleoplasm [134,135] Metarrestin Nucleolar disruption [136] CX-3543 (quarfloxin) Not determined [137] CX-5461 (pidnarulex) Not determined [138] Ellipticines Not determined [139] PMR-116 Not determined [131,140] Sodium cerivastatin Not determined [141] Aminoacridines Nucleolar disruption [142] Gold nanoparticles Redistribution of nucleolar proteins [143,144] Inhibition of early rRNA processing Camptothecin and derivatives Intranucleolar bodies [115] Nucleolar caps, translocation of nucleolar proteins to nucleoplasm [132] Flavopiridol Nucleolar necklace, translocation of nucleolar proteins to nucleoplasm [132] Roscovitine Nucleolar necklace, translocation of nucleolar proteins to nucleoplasm [132] DRB Nucleolar necklace, translocation of nucleolar proteins to nucleoplasm [132,145] TBA Redistribution of nucleolar proteins [146] Inhibition of late rRNA processing Etoposide Intranucleolar bodies [115] Nucleolar spots, translocation of nucleolar proteins to nucleoplasm [132] MG-132 Nucleolar aggresomes [114] None [132,147] Homoharringtonine None [132] Cycloheximide None [132] 5-fluorouracil None [132] Inhibition of subunit assembly or maturation Diazaborine None [148] Ribozinoindole-1 None [149] Nuclear export inhibition Leptomycin B Changes in the nucleolar composition [150] (continued on next page) K. Gonz´ alez-Arzola BBA - Gene Regulatory Mechanisms 1867 (2024) 195029 8 trigger nucleolar stress by rDNA transcription inhibition [132]. However, recent studies have highlighted that, at clinically relevant doses, oxaliplatin causes cell death via ribosome biogenesis stress, whereas cisplatin induces cytotoxicity via the DNA damage response [162–164]. At high doses (100 μ M), cisplatin induces the formation of adducts with DNA and UBF, thus hijacking UBF and repressing rDNA transcription [11,133]. However, at a low concentration (6.25 μ M), cisplatin neither inhibits rDNA transcription nor induces nucleolar stress [164]. This different drug mechanism for cell death triggering may elucidate the distinct clinical implementation of platinum analogs: cisplatin is mainly employed in breast and lung tumors, while oxaliplatin is used almost solely in colorectal and other gastrointestinal cancers [162]. The mechanism by which oxaliplatin induces ribosome biogenesis stress is not fully understood to date, but it is thought that it also involves UBF sequestration [11]. A high-throughput screening of compounds with antitumor activity led to the identification of the drug BMH-21 as a potent activator of p53 without triggering the cellular DNA damage response [134]. This intercalating compound binds GC-rich sequences of rDNA, thus hindering POL-I transcription. In addition, BMH-21 [135], aminoacridines [165], hernandonine [166], and sempervirine [167] promote the proteasome-mediated degradation of one of the POL-I catalytic subunits (RNA polymerase I subunit A or RPA194). Metarrestin is another anticancer drug that effectively suppresses metastasis by hindering rDNA transcription and disrupting the nucleolar structure [136]. The anticancer drug CX-3543 disrupts complexes between NCL and rDNA G-quadruplex, leading to the inhibition of elongation by POL-I [137]. A molecule called CX-5461, another fluoroquinolone derivative with antitumoral potential, interferes with the interaction between the SL1 complex and POL-I, thus blocking the pre-initiation complex formation [138]. Significantly, CX-5461 has been recently approved by the American Food and Drug Administration (FDA) for the fast-track program to treat breast cancer [11]. However, its antitumoral effect is thought to be related to its ability to inhibit topoisomerase II rather than a POL-I blocking effect [168]. Ellipticines are plant alkaloids used as antitumoral agents that have been found to inhibit POL-I-mediated transcription since they cause SL1 dissociation from the rDNA promoter [139]. The newly developed drug PMR-116 by Pimera Therapeutics has high tumor activity and great potential as a POL-I inhibitor; however, its inhibitory mechanism remains unclear [131,140]. Also recently, an already FDA-approved compound, sodium cerivastatin, has been tested as a POL-I inhibitor due to its ability to interfere with the polymerase’s interaction with TIF-IA, thus giving rise to rDNA transcription blockage [141]. In addition, rapamycin, a well-known inhibitor of the mammalian target of the rapamycin (mTOR) pathway, impedes phosphorylation of TIF-IA, thus interfering with the POL-I transcription machinery [169]. POL-III transcribes the 5S RNA, making it a feasible selective target for ribosome synthesis inhibition [170]. New investigations showed that the natural lactone triptolide disrupts the formation of the transcription factor-IIIB (TFIIIB) complex (the main POL-III transcription factor) at 5S rDNA promoters, thus inhibiting 5S RNA transcription [170]. One common effect of rDNA transcription inhibition is a nucleolar disorganization characterized by the condensation and separation of the nucleolus into bipartite structures composed by the FC and GC, forming structures around the nucleolar remnant, the so-called nucleolar caps [127] (Fig. 5). Nucleolar caps are dynamic structures in which the FC and DFC are inverted and surrounded by the GC. They are hallmarks of the transcriptional shutdown and often contain nucleolar components, such as rDNA, FBL, UBF, and nucleoplasmic RBPs [127]. Gold nanoparticles have been found to interfere with the transcription of rDNA and compromise rRNA processing [144]. These nanoparticles are inert and biocompatible, making them promising tools for cancer therapy. Due to their ability to induce photothermal killing preferentially in tumoral cells, gold nanoparticles could reduce the severe side effects of other anticancer treatments [171]. Indeed, a study has shown that incorporating gold nanospheres combined with mild heat stress can reduce the viability and proliferation of breast cancer cells [143]. 3.2.2. Alterations in rRNA processing, ribosomal assembly, or ribonucleoprotein transport Processing the 47S rRNA polycistronic precursor requires several steps to obtain the mature rRNAs. Camptothecin, topotecan, and irinotecan are type-I topoisomerase inhibitors that mainly disturb early stages of the rRNA processing, measured as a defective generation of the 32S rRNA intermediate, while barely affecting rDNA transcription [132]. Acridine derivatives also abolish the early processing of rRNAs [142]. Other compounds also affecting early rRNA processing are the CDK inhibitors flavopiridol and roscovitine [132]. The exact mechanism of these inhibitors is not yet fully understood. It was thought that it relied on downregulation of the U8 snoRNA, essential for 3 ′ rRNA processing, upon CDK9 inhibition [172]. However, new research underlined the involvement of nucleolar POL-II and CDK9 in improving the POL-I- mediated transcription by counteracting the adverse effects of sense intergenic non-coding RNAs (sincRNAs) produced by POL-I [83], which expands the role of CDK inhibitors beyond the processing of rRNA. The nucleotide analog 5,6-dichloro-1-β-D-ribofuranosylbenzimidazole (DRB) is a POL-II inhibitor whose activity depends on casein kinase II (CKII) [173]. DRB has been found to interfere with the early processing of rRNA, thus producing nucleolar stress [132,145]. When early rRNA processing is impaired, but the rDNA transcription remains active, a different nucleolar re-organization appears, the so-called nucleolar necklace [145] (Fig. 5). Each necklace bead corresponds to the FC-DFC modules. Besides rDNA and rRNAs, it contains POL-I, UBF, FBL, and TOPOI [145]. Among the drugs suppressing the late steps of the rRNA processing Table 1 (continued) Stress type Cause Effect on nucleolar structure Ref. Ser/Thr protein phosphatase inhibition Okadaic acid Nucleolar accumulation of p80 coilin and splicing small nuclear ribonucleoproteins (snRNPs) [151] Viral infection Adenovirus, coronavirus, poliovirus, etc. Changes in nucleolar morphology and composition [152,153] Nutrient deprivation Serum removal Downregulation of rDNA transcription and reduced ribosome production [154] Hypoxia Reduction of the cellular concentration of O 2 Nucleolar disruption, downregulation of rDNA transcription [155] Oxidative stress H 2 O 2 Translocation of nucleolar proteins (NPM) to nucleoplasm, downregulation of rDNA transcription [156] Osmotic stress Hypoosmotic conditions Not determined [157] Osmotic stress Hyperosmotic conditions Loss of nucleolar FC, translocation of nucleolar proteins (NCL) to nucleoplasm [158] Thermal stress Heat shock Accumulation of misfolded proteins in the GC [159] Formation of A-bodies [109,110] Cold-warm shock Export of the RNA exosome complex from the nucleoli to the nucleoplasm [160] Acidosis pH decrease Formation of A-bodies [109,110] K. Gonz´ alez-Arzola BBA - Gene Regulatory Mechanisms 1867 (2024) 195029 9 are molecules classically related to DNA metabolism, but they also affect RNA metabolism. This is the case of the type II topoisomerase poison etoposide and the DNA/RNA synthesis inhibitor 5-fluorouracil [132]. The proteasome inhibitors MG-132 and bortezomib, besides the translation inhibitors homoharringtonine and cycloheximide, also block late rRNA processing [132] by undetermined mechanisms. Remarkably, compounds inhibiting late rRNA processing apparently did not induce alterations in nucleolar structure [132]. A derivative of a thrombin-binding aptamer (TBA), which is a short DNA oligonucleotide that adopts G-quadruplex structures, has been reported to induce nucleolar stress by impairing rRNA processing, leading to the accumulation of pre-rRNAs. This causes cell cycle arrest or apoptosis in p53-deficient colon cancer cells [146]. Remarkably, the cytotoxic activity of the TBA derivative is mediated by the ribosomal protein RPL3, which is translocated from the nucleolus to the nucleoplasm to positively regulate p21 expression in a p53-independent manner [146]. Owing to its antiproliferative activity, this aptamer can be used as a therapeutic tool against tumor development due to its Fig. 4. Schematic representation of action mechanisms of compounds targeting ribosome biogenesis. Overview of the effects of diverse anticancer drugs on rDNA transcription, early or late rRNA processing, ribosomal subunit assembly and maturation, and nuclear export of the ribonucleoprotein complexes. Processing the 47S rRNA precursor into mature 28S, 5.8S, and 18S rRNAs is a multi-step process that produces different rRNA intermediates (e.g., 32S rRNA). Actinomycin D (low concentration), mitomycin C, methotrexate, BMH-21, metarrestin, and PMR-116 block transcription by RNA polymerase-I (POL-I). CX-5461 disrupts the interaction between promoter selectivity factor complex (SL1) and POL-I, whereas ellipticines elicit SL1 dissociation from the rDNA promoter. Sodium cerivastatin and rapamycin interfere with the transcription initiation factor-IA (TIF-IA)-POL-I interaction. Doxorubicin and mitoxantrone abolish rDNA transcription by inhibiting type-II topoisomerases (TOPO II). Oxaliplatin and cisplatin (at high doses) form adducts with DNA and upstream binding factor (UBF), leading to UBF sequestration. BMH-21, aminoacridines, hernandonine, and sempervirine promote the ubiquitin-mediated proteasome degradation of one catalytic subunit of POL-I (RPA194). CX-3543 interrupts the binding of rDNA G-quadruplex to nucleolin (NCL). Triptolide disrupts the formation of the transcription factor-IIIB (TFIIIB) complex at the 5S rDNA promoter, thus downregulating RNA polymerase III (POL-III). Camptothecin and its derivatives are type-I topoisomerase inhibitors that mainly affect early processing of the rRNA. Aminoacridines also interfere with this early process. Flavopiridol and roscovitine are cyclin-dependent kinase-9 (CDK9) inhibitors that impede phosphorylation and activation of RNA polymerase-II (POL-II), thus avoiding the POL-II-mediated transcription of the small nucleolar RNA (snoRNA) U8, essential for early rRNA processing. CDK9 inhibitors also promote the POL-I-interposed rDNA transcription by hampering the POL-II-mediated transcription of anti-sense intergenic non-coding RNAs (asincRNAs) that counteract the adverse effects of POL-I-transcribed sense intergenic non-coding RNAs (sincRNAs). 5,6-dichloro-1-beta-D-ribofuranosylbenzimidazole (DRB) is a casein kinase II (CKII) inhibitor that prevents activation of POL-II, which, in turn, interferes with the early processing of rRNA by snoRNA U8. Etoposide, MG-132, bortezomib, homoharringtonine, cycloheximide, and 5-fluo- rouracil hinder the late steps of the rRNA processing process by undetermined mechanisms: diazaborine and ribozinoindole-1 block assembly and maturation of the large ribosomal subunit, respectively. Leptomycin B hinders the export receptor chromosome region maintenance 1 (CRM1), thus inhibiting the nuclear export of ribosomal subunits. P: a phosphorylated amino-acid residue. K. Gonz´ alez-Arzola BBA - Gene Regulatory Mechanisms 1867 (2024) 195029 16 Acknowledgements The author would like to express gratitude to Dr. Jose Carlos Reyes for providing constructive criticism on the review. References [1] T. Hirose, K. Ninomiya, S. Nakagawa, T. Yamazaki, A guide to membraneless organelles and their various roles in gene regulation, Nat. Rev. Mol. Cell Biol. 24 (2023) 288–304. [2] H. Zhang, W. Qin, H. Romero, H. Leonhardt, M.C. Cardoso, Heterochromatin organization and phase separation, Nucleus 14 (2023) 2159142. [3] H.B. Schmidt, D. G¨ orlich, Transport selectivity of nuclear pores, phase separation, and membraneless organelles, Trends Biochem. Sci. 41 (2016) 46–61. [4] A.I. Lamond, T. Ly, S. Hutten, A. Nicolas, The nucleolus, in: R.A. Bradshaw, P. D. Stahl (Eds.), Encyclopedia of Cell Biology, Academic Press, Place Published, 2016, pp. 254–269. [5] D. Sakthivel, A. Brown-Suedel, L. Bouchier-Hayes, The role of the nucleolus in regulating the cell cycle and the DNA damage response, Adv. Protein Chem. Struct. Biol. 135 (2023) 203–241. [6] M.C. Lafita-Navarro, M. Conacci-Sorrell, Nucleolar stress: from development to cancer, Semin. Cell Dev. Biol. 136 (2023) 64–74. [7] S. Boulon, B.J. Westman, S. Hutten, F.M. Boisvert, A.I. Lamond, The nucleolus under stress, Mol. Cell 40 (2010) 216–227. [8] L.M. Ogawa, S.J. Baserga, Crosstalk between the nucleolus and the DNA damage response, Mol. BioSyst. 13 (2017) 443–455. [9] D. Hernandez-Verdun, Assembly and disassembly of the nucleolus during the cell cycle, Nucleus 2 (2011) 189–194. [10] K. Yang, J. Yang, J. Yi, Nucleolar stress: hallmarks, sensing mechanism and diseases, Cell Stress 2 (2018) 125–140. [11] A. Corman, O. Sirozh, V. Lafarga, O. Fernandez-Capetillo, Targeting the nucleolus as a therapeutic strategy in human disease, Trends Biochem. Sci. 48 (2023) 274–287. [12] J.L. Woolford Jr., S.J. Baserga, Ribosome biogenesis in the yeast Saccharomyces cerevisiae, Genetics 195 (2013) 643–681. [13] B. McStay, I. Grummt, The epigenetics of rRNA genes: from molecular to chromosome biology, Annu. Rev. Cell Dev. Biol. 24 (2008) 131–157. [14] T. Kobayashi, A new role of the rDNA and nucleolus in the nucleus—rDNA instability maintains genome integrity, BioEssays 30 (2008) 267–272. [15] M. Tsekrekou, K. Stratigi, G. Chatzinikolaou, The nucleolus: in genome maintenance and repair, Int. J. Mol. Sci. 18 (2017). [16] M. van Sluis, B. McStay, A localized nucleolar DNA damage response facilitates recruitment of the homology-directed repair machinery independent of cell cycle stage, Genes Dev. 29 (2015) 1151–1163. [17] R. Santoro, I. Grummt, Molecular mechanisms mediating methylation-dependent silencing of ribosomal gene transcription, Mol. Cell 8 (2001) 719–725. [18] Y. Hori, C. Engel, T. Kobayashi, Regulation of ribosomal RNA gene copy number, transcription and nucleolus organization in eukaryotes, Nat. Rev. Mol. Cell Biol. 24 (2023) 414–429. [19] C. Herdman, J.C. Mars, V.Y. Stefanovsky, M.G. Tremblay, M. Sabourin-Felix, H. Lindsay, M.D. Robinson, T. Moss, A unique enhancer boundary complex on the mouse ribosomal RNA genes persists after loss of Rrn3 or UBF and the inactivation of RNA polymerase I transcription, PLoS Genet. 13 (2017) e1006899. [20] B. McClintock, The relation of a particular chromosomal element to the development of the nucleoli in Zea mays, Z. Zellforsch. Mikrosk. Anat. 21 (1934) 294–326. [21] A.S. Henderson, D. Warburton, K.C. Atwood, Location of ribosomal DNA in the human chromosome complement, Proc. Natl. Acad. Sci. U. S. A. 69 (1972) 3394–3398. [22] F.-M. Boisvert, S. van Koningsbruggen, J. Navascu´ es, A.I. Lamond, The multifunctional nucleolus, Nat. Rev. Mol. Cell Biol. 8 (2007) 574–585. [23] I. Floutsakou, S. Agrawal, T.T. Nguyen, C. Seoighe, A.R.D. Ganley, B. McStay, The shared genomic architecture of human nucleolar organizer regions, Genome Res. 23 (2013) 2003–2012. [24] C. Mais, J.E. Wright, J.L. Prieto, S.L. Raggett, B. McStay, UBF-binding site arrays form pseudo-NORs and sequester the RNA polymerase I transcription machinery, Genes Dev. 19 (2005) 50–64. [25] M. van Sluis, C. van Vuuren, H. Mangan, B. McStay, NORs on human acrocentric chromosome p-arms are active by default and can associate with nucleoli independently of rDNA, Proc. Natl. Acad. Sci. U. S. A. 117 (2020) 10368–10377. [26] C.P. Brangwynne, T.J. Mitchison, A.A. Hyman, Active liquid-like behavior of nucleoli determines their size and shape in Xenopus laevis oocytes, Proc. Natl. Acad. Sci. 108 (2011) 4334–4339. [27] M. Feric, N. Vaidya, T.S. Harmon, D.M. Mitrea, L. Zhu, T.M. Richardson, R. W. Kriwacki, R.V. Pappu, C.P. Brangwynne, Coexisting liquid phases underlie nucleolar subcompartments, Cell 165 (2016) 1686–1697. [28] M. Anastassova-Kristeva, The nucleolar cycle in man, J. Cell Sci. 25 (1977) 103–110. [29] T.C. Hsu, S.E. Spirito, M.L. Pardue, Distribution of 18+28S ribosomal genes in mammalian genomes, Chromosoma 53 (1975) 25–36. [30] J.A. Riback, J.M. Eeftens, D.S.W. Lee, S.A. Quinodoz, A. Donlic, N. Orlovsky, L. Wiesner, L. Beckers, L.A. Becker, A.R. Strom, U. Rana, M. Tolbert, B.W. Purse, R. Kleiner, R. Kriwacki, C.P. Brangwynne, Viscoelasticity and advective flow of RNA underlies nucleolar form and function, Mol. Cell 83 (2023), 3095-3107. e3099. [31] D.L.J. Lafontaine, J.A. Riback, R. Bascetin, C.P. Brangwynne, The nucleolus as a multiphase liquid condensate, Nat. Rev. Mol. Cell Biol. 22 (2021) 165–182. [32] R.W. Yao, G. Xu, Y. Wang, L. Shan, P.F. Luan, Y. Wang, M. Wu, L.Z. Yang, Y. H. Xing, L. Yang, L.L. Chen, Nascent pre-rRNA sorting via phase separation drives the assembly of dense fibrillar components in the human nucleolus, Mol. Cell 76 (2019), 767-783.e711. [33] F. Lamaye, S. Galliot, L. Alibardi, D.L. Lafontaine, M. Thiry, Nucleolar structure across evolution: the transition between bi- and tri-compartmentalized nucleoli lies within the class Reptilia, J. Struct. Biol. 174 (2011) 352–359. [34] M. Thiry, D.L.J. Lafontaine, Birth of a nucleolus: the evolution of nucleolar compartments, Trends Cell Biol. 15 (2005) 194–199. [35] N. Jaberi-Lashkari, B. Lee, F. Aryan, E. Calo, An evolutionarily nascent architecture underlying the formation and emergence of biomolecular condensates, Cell Rep. 42 (2023) 112955. [36] D.L.J. Lafontaine, When two became three: shaping the nucleolus with treacle, Cell Rep. 42 (2023) 113060. [37] D. Hernandez-Verdun, P. Roussel, M. Thiry, V. Sirri, D.L.J. Lafontaine, The nucleolus: structure/function relationship in RNA metabolism, WIREs RNA 1 (2010) 415–431. [38] K. Murano, M. Okuwaki, M. Hisaoka, K. Nagata, Transcription regulation of the rRNA gene by a multifunctional nucleolar protein, B23/nucleophosmin, through its histone chaperone activity, Mol. Cell. Biol. 28 (2008) 3114–3126. [39] L.B. Maggi Jr., M. Kuchenruether, D.Y. Dadey, R.M. Schwope, S. Grisendi, R. R. Townsend, P.P. Pandolfi, J.D. Weber, Nucleophosmin serves as a rate-limiting nuclear export chaperone for the mammalian ribosome, Mol. Cell. Biol. 28 (2008) 7050–7065. [40] I. Ugolini, S. Bilokapic, M. Ferrolino, J. Teague, Y. Yan, X. Zhou, A. Deshmukh, M. White, R.W. Kriwacki, M. Halic, Chromatin localization of nucleophosmin organizes ribosome biogenesis, Mol. Cell 82 (2022), 4443-4457.e4449. [41] J.K. Box, N. Paquet, M.N. Adams, D. Boucher, E. Bolderson, K.J. O’Byrne, D. J. Richard, Nucleophosmin: from structure and function to disease development, BMC Mol. Biol. 17 (2016) 1–12. [42] C. Korgaonkar, J. Hagen, V. Tompkins, A.A. Frazier, C. Allamargot, F.W. Quelle, D.E. Quelle, Nucleophosmin (B23) targets ARF to nucleoli and inhibits its function, Mol. Cell. Biol. 25 (2005) 1258–1271. [43] K. Gonz´ alez-Arzola, A. Díaz-Quintana, N. Bernardo-García, J. Martínez-F´ abregas, F. Rivero-Rodríguez, M.´ A. Casado-Combreras, C.A. Elena-Real, A. Vel´ azquez- Cruz, S. Gil-Caballero, A. Vel´ azquez-Campoy, E. Szulc, M.P. Gavil´ an, I. Ayala, R. Arranz, R.M. Ríos, X. Salvatella, J.M. Valpuesta, J.A. Hermoso, M.A. De la Rosa, I. Díaz-Moreno, Nucleus-translocated mitochondrial cytochrome c liberates nucleophosmin-sequestered ARF tumor suppressor by changing nucleolar liquid–liquid phase separation, Nat. Struct. Mol. Biol. 29 (2022) 1024–1036. [44] D.M. Mitrea, J.A. Cika, C.S. Guy, D. Ban, P.R. Banerjee, C.B. Stanley, A. Nourse, A. A. Deniz, R.W. Kriwacki, Nucleophosmin integrates within the nucleolus via multi-modal interactions with proteins displaying R-rich linear motifs and rRNA, eLife 5 (2016) e13571. [45] D.M. Mitrea, J.A. Cika, C.B. Stanley, A. Nourse, P.L. Onuchic, P.R. Banerjee, A. H. Phillips, C.-G. Park, A.A. Deniz, R.W. Kriwacki, Self-interaction of NPM1 modulates multiple mechanisms of liquid–liquid phase separation, Nat. Commun. 9 (2018) 842. [46] R. Cong, S. Das, I. Ugrinova, S. Kumar, F. Mongelard, J. Wong, P. Bouvet, Interaction of nucleolin with ribosomal RNA genes and its role in RNA polymerase I transcription, Nucleic Acids Res. 40 (2012) 9441–9454. [47] H. Ginisty, H. Sicard, B. Roger, P. Bouvet, Structure and functions of nucleolin, J. Cell Sci. 112 (1999) 761–772. [48] E. Doron-Mandel, I. Koppel, O. Abraham, I. Rishal, T.P. Smith, C.N. Buchanan, P. K. Sahoo, J. Kadlec, J.A. Oses-Prieto, R. Kawaguchi, S. Alber, E.E. Zahavi, P. Di Matteo, A. Di Pizio, D.-A. Song, N. Okladnikov, D. Gordon, S. Ben-Dor, R. Haffner- Krausz, G. Coppola, A.L. Burlingame, P. Jungwirth, J.L. Twiss, M. Fainzilber, The glycine arginine-rich domain of the RNA-binding protein nucleolin regulates its subcellular localization, EMBO J. 40 (2021) e107158. [49] A. N´ emeth, A. Conesa, J. Santoyo-Lopez, I. Medina, D. Montaner, B. P´ eterfia, I. Solovei, T. Cremer, J. Dopazo, G. L¨ angst, Initial genomics of the human nucleolus, PLoS Genet. 6 (2010) e1000889. [50] C. Bersaglieri, R. Santoro, Methods for mapping 3D-chromosome architecture around nucleoli, Curr. Opin. Cell Biol. 81 (2023) 102171. [51] M. van Sluis, B. McStay, Nucleolar DNA double-Strand break responses underpinning rDNA genomic stability, Trends Genet. 35 (2019) 743–753. [52] B. McStay, Nucleolar organizer regions: genomic ‘dark matter’ requiring illumination, Genes Dev. 30 (2016) 1598–1610. [53] M. van Sluis, B. McStay, Nucleolar reorganization in response to rDNA damage, Curr. Opin. Cell Biol. 46 (2017) 81–86. [54] L. Stenstr¨ om, D. Mahdessian, C. Gnann, A.J. Cesnik, W. Ouyang, M.D. Leonetti, M. Uhl´ en, S. Cuylen-Haering, P.J. Thul, E. Lundberg, Mapping the nucleolar proteome reveals a spatiotemporal organization related to intrinsic protein disorder, Mol. Syst. Biol. 16 (2020) e9469. [55] L. Shan, G. Xu, R.W. Yao, P.F. Luan, Y. Huang, P.H. Zhang, Y.H. Pan, L. Zhang, X. Gao, Y. Li, S.M. Cao, S.X. Gao, Z.H. Yang, S. Li, L.Z. Yang, Y. Wang, C.C. L. Wong, L. Yu, J. Li, L. Yang, L.L. Chen, Nucleolar URB1 ensures 3’ ETS rRNA removal to prevent exosome surveillance, Nature 615 (2023) 526–534. [56] V. Stamatopoulou, P. Parisot, C. De Vleeschouwer, D.L.J. Lafontaine, Use of the iNo score to discriminate normal from altered nucleolar morphology, with applications in basic cell biology and potential in human disease diagnostics, Nat. Protoc. 13 (2018) 2387–2406. K. Gonz´ alez-Arzola BBA - Gene Regulatory Mechanisms 1867 (2024) 195029 17 [57] E. Nicolas, P. Parisot, C. Pinto-Monteiro, R. de Walque, C. De Vleeschouwer, D.L. J. Lafontaine, Involvement of human ribosomal proteins in nucleolar structure and p53-dependent nucleolar stress, Nat. Commun. 7 (2016) 11390. [58] M. Haruka, W. Kenji, T. Hiroaki, F. Tomoko, I. Yuma, T. Makio, S.-S. Kumiko, O. Hiroko, H. Tokuko, A. Akinori, O. Hiroshi, S. Yuka, O. Koji, T. Tsutomu, I. Etsuro, G.G. Ilya, T. Kazuaki, N. Mitsuyoshi, S. Noriko, Ribosomal protein L5 facilitates rDNA-bundled condensate and nucleolar assembly, Life Sci. Alliance 5 (2022) e202101045. [59] S. Fumagalli, V.V. Ivanenkov, T. Teng, G. Thomas, Suprainduction of p53 by disruption of 40S and 60S ribosome biogenesis leads to the activation of a novel G2/M checkpoint, Genes Dev. 26 (2012) 1028–1040. [60] D. Ballmer, M. Tardat, R. Ortiz, A. Graff-Meyer, Evgeniy A. Ozonov, C. Genoud, Antoine H. Peters, G. Fanourgakis, HP1 proteins regulate nucleolar structure and function by secluding pericentromeric constitutive heterochromatin, Nucleic Acids Res. 51 (2023) 117–143. [61] A.G. Larson, D. Elnatan, M.M. Keenen, M.J. Trnka, J.B. Johnston, A. L. Burlingame, D.A. Agard, S. Redding, G.J. Narlikar, Liquid droplet formation by HP1 α suggests a role for phase separation in heterochromatin, Nature 547 (2017) 236–240. [62] S. Sanulli, M.J. Trnka, V. Dharmarajan, R.W. Tibble, B.D. Pascal, A.L. Burlingame, P.R. Griffin, J.D. Gross, G.J. Narlikar, HP1 reshapes nucleosome core to promote phase separation of heterochromatin, Nature 575 (2019) 390–394. [63] X. Jin, H. Tanaka, M. Jin, K. Fujita, H. Homma, M. Inotsume, H. Yong, K. Umeda, N. Kodera, T. Ando, H. Okazawa, PQBP5/NOL10 maintains and anchors the nucleolus under physiological and osmotic stress conditions, Nat. Commun. 14 (2023) 9. [64] D. Xu, X. Chen, Y. Kuang, M. Hong, T. Xu, K. Wang, X. Huang, C. Fu, K. Ruan, C. Zhu, X. Feng, S. Guang, rRNA intermediates coordinate the formation of nucleolar vacuoles in C. elegans, Cell Rep. 42 (2023) 112915. [65] P.F. Islas-Morales, A. C´ ardenas, M.J. Mosqueira, L.F. Jim´ enez-García, C. R. Voolstra, Ultrastructural and proteomic evidence for the presence of a putative nucleolus in an Archaeon, Front. Microbiol. 14 (2023) 1075071. [66] T.A. Potapova, J.L. Gerton, Ribosomal DNA and the nucleolus in the context of genome organization, Chromosom. Res. 27 (2019) 109–127. [67] S.A. Pirogov, V.A. Gvozdev, M.S. Klenov, Long noncoding RNAs and stress response in the nucleolus, Cells 8 (2019) 668. [68] L. Comai, N. Tanese, R. Tjian, The TATA-binding protein and associated factors are integral components of the RNA polymerase I transcription factor, SL1, Cell 68 (1992) 965–976. [69] S.J. Goodfellow, J.C. Zomerdijk, Basic mechanisms in RNA polymerase I transcription of the ribosomal RNA genes, Subcell. Biochem. 61 (2013) 211–236. [70] M. Kermekchiev, J.L. Workman, C.S. Pikaard, Nucleosome binding by the polymerase I transactivator upstream binding factor displaces linker histone H1, Mol. Cell. Biol. 17 (1997) 5833–5842. [71] S. Ray, T. Panova, G. Miller, A. Volkov, A.C. Porter, J. Russell, K.I. Panov, J. C. Zomerdijk, Topoisomerase II α promotes activation of RNA polymerase I transcription by facilitating pre-initiation complex formation, Nat. Commun. 4 (2013) 1598. [72] K.M. Rose, J. Szopa, F.S. Han, Y.C. Cheng, A. Richter, U. Scheer, Association of DNA topoisomerase I and RNA polymerase I: a possible role for topoisomerase I in ribosomal gene transcription, Chromosoma 96 (1988) 411–416. [73] Z. G´ al, B. Nieto, S. Boukoura, A.V. Rasmussen, D.H. Larsen, Treacle sticks the nucleolar responses to DNA damage together, Front. Cell Dev. Biol. 10 (2022). [74] N.M. Castillo Duque de Estrada, M. Thoms, D. Flemming, H.M. Hammaren, R. Buschauer, M. Ameismeier, J. Baßler, M. Beck, R. Beckmann, E. Hurt, Structure of nascent 5S RNPs at the crossroad between ribosome assembly and MDM2–p53 pathways, Nat. Struct. Mol. Biol. 30 (2023) 1119–1131. [75] S. Tisdale, L. Pellizzoni, Chapter 7 - RNA-processing dysfunction in spinal muscular atrophy, in: C.J. Sumner, S. Paushkin, C.-P. Ko (Eds.), Spinal Muscular Atrophy, Academic Press, Place Published, 2017, pp. 113–131. [76] A.K. Henras, C. Plisson-Chastang, M.F. O’Donohue, A. Chakraborty, P.E. Gleizes, An overview of pre-ribosomal RNA processing in eukaryotes, Wiley Interdiscip. Rev. RNA 6 (2015) 225–242. [77] S.M. Plafker, I.G. Macara, Ribosomal protein L12 uses a distinct nuclear import pathway mediated by importin 11, Mol. Cell. Biol. 22 (2002) 1266–1275. [78] P. Bouvet, J.-J. Diaz, K. Kindbeiter, J.-J. Madjar, F. Amalric, Nucleolin interacts with several ribosomal proteins through its RGG domain*, J. Biol. Chem. 273 (1998) 19025–19029. [79] M.S. Lindstr¨ om, Elucidation of motifs in ribosomal protein S9 that mediate its nucleolar localization and binding to NPM1/nucleophosmin, PLoS One 7 (2012) e52476. [80] D.L.J. Lafontaine, Noncoding RNAs in eukaryotic ribosome biogenesis and function, Nat. Struct. Mol. Biol. 22 (2015) 11–19. [81] C. Verheggen, E. Bertrand, CRM1 plays a nuclear role in transporting snoRNPs to nucleoli in higher eukaryotes, Nucleus 3 (2012) 132–137. [82] P. Nerurkar, M. Altvater, S. Gerhardy, S. Schütz, U. Fischer, C. Weirich, V. G. Panse, Eukaryotic ribosome assembly and nuclear export, Int. Rev. Cell Mol. Biol. 319 (2015) 107–140. [83] K.J. Abraham, N. Khosraviani, J.N.Y. Chan, A. Gorthi, A. Samman, D.Y. Zhao, M. Wang, M. Bokros, E. Vidya, L.A. Ostrowski, R. Oshidari, V. Pietrobon, P. S. Patel, A. Algouneh, R. Singhania, Y. Liu, V.T. Yerlici, D.D. De Carvalho, M. Ohh, B.C. Dickson, R. Hakem, J.F. Greenblatt, S. Lee, A.J.R. Bishop, K. Mekhail, Nucleolar RNA polymerase II drives ribosome biogenesis, Nature 585 (2020) 298–302. [84] J. Heix, A. Vente, R. Voit, A. Budde, T.M. Michaelidis, I. Grummt, Mitotic silencing of human rRNA synthesis: inactivation of the promoter selectivity factor SL1 by cdc2/cyclin B-mediated phosphorylation, EMBO J. 17 (1998) 7373–7381. [85] N. Angelier, M. Tramier, E. Louvet, M. Coppey-Moisan, T.M. Savino, J.R. De Mey, D. Hernandez-Verdun, Tracking the interactions of rRNA processing proteins during nucleolar assembly in living cells, Mol. Biol. Cell 16 (2005) 2862–2871. [86] Y. Ahmad, F.M. Boisvert, P. Gregor, A. Cobley, A.I. Lamond, NOPdb: nucleolar proteome database–2008 update, Nucleic Acids Res. 37 (2009) D181–D184. [87] J.S. Andersen, Y.W. Lam, A.K.L. Leung, S.-E. Ong, C.E. Lyon, A.I. Lamond, M. Mann, Nucleolar proteome dynamics, Nature 433 (2005) 77–83. [88] M.A. Jarboui, K. Wynne, G. Elia, W.W. Hall, V.W. Gautier, Proteomic profiling of the human T-cell nucleolus, Mol. Immunol. 49 (2011) 441–452. [89] C.J. Sherr, J.D. Weber, The ARF/p53 pathway, Curr. Opin. Genet. Dev. 10 (2000) 94–99. [90] C.P. Rubbi, J. Milner, Disruption of the nucleolus mediates stabilization of p53 in response to DNA damage and other stresses, EMBO J. 22 (2003) 6068–6077. [91] J.D. Oliner, J.A. Pietenpol, S. Thiagalingam, J. Gyuris, K.W. Kinzler, B. Vogelstein, Oncoprotein MDM2 conceals the activation domain of tumour suppressor p53, Nature 362 (1993) 857–860. [92] D. Bertwistle, M. Sugimoto, C.J. Sherr, Physical and functional interactions of the Arf tumor suppressor protein with nucleophosmin/B23, Mol. Cell. Biol. 24 (2004) 985–996. [93] S. Kurki, K. Peltonen, L. Latonen, T.M. Kiviharju, P.M. Ojala, D. Meek, M. Laiho, Nucleolar protein NPM interacts with HDM2 and protects tumor suppressor protein p53 from HDM2-mediated degradation, Cancer Cell 5 (2004) 465–475. [94] M.S. Dai, H. Lu, Inhibition of MDM2-mediated p53 ubiquitination and degradation by ribosomal protein L5, J. Biol. Chem. 279 (2004) 44475–44482. [95] D. Bai, J. Zhang, W. Xiao, X. Zheng, Regulation of the HDM2-p53 pathway by ribosomal protein L6 in response to ribosomal stress, Nucleic Acids Res. 42 (2014) 1799–1811. [96] M.A. Lohrum, R.L. Ludwig, M.H. Kubbutat, M. Hanlon, K.H. Vousden, Regulation of HDM2 activity by the ribosomal protein L11, Cancer Cell 3 (2003) 577–587. [97] M.S. Dai, S.X. Zeng, Y. Jin, X.X. Sun, L. David, H. Lu, Ribosomal protein L23 activates p53 by inhibiting MDM2 function in response to ribosomal perturbation but not to translation inhibition, Mol. Cell. Biol. 24 (2004) 7654–7668. [98] S. Yadavilli, L.D. Mayo, M. Higgins, S. Lain, V. Hegde, W.A. Deutsch, Ribosomal protein S3: a multi-functional protein that interacts with both p53 and MDM2 through its KH domain, DNA Repair 8 (2009) 1215–1224. [99] X. Zhou, Q. Hao, J. Liao, Q. Zhang, H. Lu, Ribosomal protein S14 unties the MDM2–p53 loop upon ribosomal stress, Oncogene 32 (2013) 388–396. [100] X. Xiong, Y. Zhao, F. Tang, D. Wei, D. Thomas, X. Wang, Y. Liu, P. Zheng, Y. Sun, Ribosomal protein S27-like is a physiological regulator of p53 that suppresses genomic instability and tumorigenesis, eLife 3 (2014) e02236. [101] S. Lee, J.Y. Kim, Y.J. Kim, K.O. Seok, J.H. Kim, Y.J. Chang, H.Y. Kang, J.H. Park, Nucleolar protein GLTSCR2 stabilizes p53 in response to ribosomal stresses, Cell Death Differ. 19 (2012) 1613–1622. [102] A. Russo, G. Russo, Ribosomal proteins control or bypass p53 during nucleolar stress, Int. J. Mol. Sci. 18 (2017). [103] X. Chen, T. Zhang, W. Su, Z. Dou, D. Zhao, X. Jin, H. Lei, J. Wang, X. Xie, B. Cheng, Q. Li, H. Zhang, C. Di, Mutant p53 in cancer: from molecular mechanism to therapeutic modulation, Cell Death Dis. 13 (2022) 974. [104] M.-S. Dai, R. Sears, H. Lu, Feedback regulation of c-Myc by ribosomal protein L11, Cell Cycle (Georgetown, Tex.) 6 (2007) 2735–2741. [105] X. Zhou, Q. Hao, J.M. Liao, P. Liao, H. Lu, Ribosomal protein S14 negatively regulates c-Myc activity, J. Biol. Chem. 288 (2013) 21793–21801. [106] S.J. Lo, L.C. Fan, Y.F. Tsai, K.Y. Lin, H.L. Huang, T.H. Wang, H. Liu, T.C. Chen, S. F. Huang, C.J. Chang, Y.J. Lin, B.Y. Yung, S.Y. Hsieh, A novel interaction of nucleophosmin with BCL2-associated X protein regulating death evasion and drug sensitivity in human hepatoma cells, Hepatology (Baltimore, Md.) 57 (2013) 1893–1905. [107] B. Eymin, P. Claverie, C. Salon, C. Leduc, E. Col, E. Brambilla, S. Khochbin, S. Gazzeri, p14ARF activates a Tip60-dependent and p53-independent ATM/ ATR/CHK pathway in response to genotoxic stress, Mol. Cell. Biol. 26 (2006) 4339–4350. [108] I. Ohbayashi, M. Sugiyama, Plant nucleolar stress response, a new face in the NAC-dependent cellular stress responses, Front. Plant Sci. 8 (2018). [109] D. Marijan, R. Tse, K. Elliott, S. Chandhok, M. Luo, E. Lacroix, T.E. Audas, Stressspecific aggregation of proteins in the amyloid bodies, FEBS Lett. 593 (2019) 3162–3172. [110] M. Wang, X. Tao, M.D. Jacob, C.A. Bennett, J.J.D. Ho, M.L. Gonzalgo, T.E. Audas, S. Lee, Stress-induced low complexity RNA activates physiological amyloidogenesis, Cell Rep. 24 (2018), 1713-1721.e1714. [111] Timothy E. Audas, Mathieu D. Jacob, S. Lee, Immobilization of proteins in the nucleolus by ribosomal intergenic spacer noncoding RNA, Mol. Cell 45 (2012) 147–157. [112] M.D. Jacob, T.E. Audas, J. Uniacke, L. Trinkle-Mulcahy, S. Lee, Environmental cues induce a long noncoding RNA–dependent remodeling of the nucleolus, Mol. Biol. Cell 24 (2013) 2943–2953. [113] L. Latonen, Nucleolar aggresomes as counterparts of cytoplasmic aggresomes in proteotoxic stress. Proteasome inhibitors induce nuclear ribonucleoprotein inclusions that accumulate several key factors of neurodegenerative diseases and cancer, Bioessays 33 (2011) 386–395. [114] L. Latonen, H.M. Moore, B. Bai, S. J¨ a¨ amaa, M. Laiho, Proteasome inhibitors induce nucleolar aggregation of proteasome target proteins and polyadenylated RNA by altering ubiquitin availability, Oncogene 30 (2011) 790–805. K. Gonz´ alez-Arzola BBA - Gene Regulatory Mechanisms 1867 (2024) 195029 18 [115] S. Hutten, A. Prescott, J. James, S. Riesenberg, S. Boulon, Y.W. Lam, A.I. Lamond, An intranucleolar body associated with rDNA, Chromosoma 120 (2011) 481–499. [116] M.E. Gaulden, R.P. Perry, Influence of the nucleolus on mitosis as revealed by ultraviolet microbeam irradiation, Proc. Natl. Acad. Sci. 44 (1958) 553–559. [117] G. Donati, E. Brighenti, M. Vici, G. Mazzini, D. Trer´ e, L. Montanaro, M. Derenzini, Selective inhibition of rRNA transcription downregulates E2F-1: a new p53- independent mechanism linking cell growth to cell proliferation, J. Cell Sci. 124 (2011) 3017–3028. [118] V. Iadevaia, S. Caldarola, L. Biondini, A. Gismondi, S. Karlsson, I. Dianzani, F. Loreni, PIM1 kinase is destabilized by ribosomal stress causing inhibition of cell cycle progression, Oncogene 29 (2010) 5490–5499. [119] A. Russo, D. Esposito, M. Catillo, C. Pietropaolo, E. Crescenzi, G. Russo, Human rpL3 induces G₁/S arrest or apoptosis by modulating p21 (waf1/cip1) levels in a p53-independent manner, Cell Cycle (Georgetown, Tex.) 12 (2013) 76–87. [120] A. Pecoraro, P. Carotenuto, G. Russo, A. Russo, Ribosomal protein uL3 targets E2F1 and Cyclin D1 in cancer cell response to nucleolar stress, Sci. Rep. 9 (2019) 15431. [121] A. Russo, S. Maiolino, V. Pagliara, F. Ungaro, F. Tatangelo, A. Leone, G. Scalia, A. Budillon, F. Quaglia, G. Russo, Enhancement of 5-FU sensitivity by the proapoptotic rpL3 gene in p53 null colon cancer cells through combined polymer nanoparticles, Oncotarget 7 (2016) 79670–79687. [122] A. Pecoraro, P. Carotenuto, B. Franco, R. De Cegli, G. Russo, A. Russo, Role of uL3 in the crosstalk between nucleolar stress and autophagy in colon cancer cells, Int. J. Mol. Sci. 21 (2020). [123] T. Andrew, H.T. Douglas, L.G. Daniel, Autophagy and cancer therapy, Mol. Pharmacol. 85 (2014) 830. [124] A. James, Y. Wang, H. Raje, R. Rosby, P. DiMario, Nucleolar stress with and without p53, Nucleus 5 (2014) 402–426. [125] B.Y. Yung, R.K. Busch, H. Busch, A.B. Mauger, P.K. Chan, Effects of actinomycin D analogs on nucleolar phosphoprotein B23 (37,000 daltons/pI 5.1), Biochem. Pharmacol. 34 (1985) 4059–4063. [126] D.G. Pestov, Z. Strezoska, L.F. Lau, Evidence of p53-dependent cross-talk between ribosome biogenesis and the cell cycle: effects of nucleolar protein Bop1 on G(1)/ S transition, Mol. Cell. Biol. 21 (2001) 4246–4255. [127] Y. Shav-Tal, J. Blechman, X. Darzacq, C. Montagna, B.T. Dye, J.G. Patton, R. H. Singer, D. Zipori, Dynamic sorting of nuclear components into distinct nucleolar caps during transcriptional inhibition, Mol. Biol. Cell 16 (2005) 2395–2413. [128] R.P. Perry, D.E. Kelley, Inhibition of RNA synthesis by actinomycin D: characteristic dose-response of different RNA species, J. Cell. Physiol. 76 (1970) 127–139. [129] Y. Pommier, Y. Sun, S.N. Huang, J.L. Nitiss, Roles of eukaryotic topoisomerases in transcription, replication and genomic stability, Nat. Rev. Mol. Cell Biol. 17 (2016) 703–721. [130] J.L. Nitiss, Targeting DNA topoisomerase II in cancer chemotherapy, Nat. Rev. Cancer 9 (2009) 338–350. [131] R. Ferreira, J.S. Schneekloth, K.I. Panov, K.M. Hannan, R.D. Hannan, Targeting the RNA polymerase I transcription for cancer therapy comes of age, Cells 9 (2020) 266. [132] K. Burger, B. Mühl, T. Harasim, M. Rohrmoser, A. Malamoussi, M. Orban, M. Kellner, A. Gruber-Eber, E. Kremmer, M. H¨ olzel, D. Eick, Chemotherapeutic drugs inhibit ribosome biogenesis at various levels*, J. Biol. Chem. 285 (2010) 12416–12425. [133] X. Zhai, H. Beckmann, H.M. Jantzen, J.M. Essigmann, Cisplatin-DNA adducts inhibit ribosomal RNA synthesis by hijacking the transcription factor human upstream binding factor, Biochemistry 37 (1998) 16307–16315. [134] K. Peltonen, L. Colis, H. Liu, S. J¨ a¨ amaa, H.M. Moore, J. Enb¨ ack, P. Laakkonen, A. Vaahtokari, R.J. Jones, T.M. af H¨ allstr¨ om, M. Laiho, Identification of novel p53 pathway activating small-molecule compounds reveals unexpected similarities with known therapeutic agents, PLoS One 5 (2010). [135] K. Peltonen, L. Colis, H. Liu, R. Trivedi, Michael S. Moubarek, Henna M. Moore, B. Bai, Michelle A. Rudek, Charles J. Bieberich, M. Laiho, A targeting modality for destruction of RNA polymerase I that possesses anticancer activity, Cancer Cell 25 (2014) 77–90. [136] K.J. Frankowski, C. Wang, S. Patnaik, F.J. Schoenen, N. Southall, D. Li, Y. Teper, W. Sun, I. Kandela, D. Hu, C. Dextras, Z. Knotts, Y. Bian, J. Norton, S. Titus, M. A. Lewandowska, Y. Wen, K.I. Farley, L.M. Griner, J. Sultan, Z. Meng, M. Zhou, T. Vilimas, A.S. Powers, S. Kozlov, K. Nagashima, H.S. Quadri, M. Fang, C. Long, O. Khanolkar, W. Chen, J. Kang, H. Huang, E. Chow, E. Goldberg, C. Feldman, R. Xi, H.R. Kim, G. Sahagian, S.J. Baserga, A. Mazar, M. Ferrer, W. Zheng, A. Shilatifard, J. Aub´ e, U. Rudloff, J.J. Marugan, S. Huang, Metarrestin, a perinucleolar compartment inhibitor, effectively suppresses metastasis, Sci. Transl. Med. 10 (2018). [137] D. Drygin, A. Siddiqui-Jain, S. O’Brien, M. Schwaebe, A. Lin, J. Bliesath, C.B. Ho, C. Proffitt, K. Trent, J.P. Whitten, J.K. Lim, D. Von Hoff, K. Anderes, W.G. Rice, Anticancer activity of CX-3543: a direct inhibitor of rRNA biogenesis, Cancer Res. 69 (2009) 7653–7661. [138] D. Drygin, A. Lin, J. Bliesath, C.B. Ho, S.E. O’Brien, C. Proffitt, M. Omori, M. Haddach, M.K. Schwaebe, A. Siddiqui-Jain, N. Streiner, J.E. Quin, E. Sanij, M. J. Bywater, R.D. Hannan, D. Ryckman, K. Anderes, W.G. Rice, Targeting RNA polymerase I with an oral small molecule CX-5461 inhibits ribosomal RNA synthesis and solid tumor growth, Cancer Res. 71 (2011) 1418–1430. [139] W.J. Andrews, T. Panova, C. Normand, O. Gadal, I.G. Tikhonova, K.I. Panov, Old drug, new target: ellipticines selectively inhibit RNA polymerase I Transcription* *this work was supported by Medical Research Council New Investigator Award Grant 89365 (to K. P.). This article contains supplemental sequences, Tables S1–S6, and Figs. S1–S3, J. Biol. Chem. 288 (2013) 4567–4582. [140] K. Panov, S. Alsahafi, Investigation of molecular mechanisms of polymerase I (Pol-l) inhibitor PMR-116 using Isothermal Titration Calorimetry (ITC), Access Microbiol. 1 (2019). [141] X. Tan, S.G. Awuah, A cell-based screening system for RNA polymerase I inhibitors, MedChemComm 10 (2019) 1765–1774. [142] L. Anikin, D.G. Pestov, 9-Aminoacridine inhibits ribosome biogenesis by targeting both transcription and processing of ribosomal RNA, Int. J. Mol. Sci. 23 (2022). [143] M. Kodiha, E. Hutter, S. Boridy, M. Juhas, D. Maysinger, U. Stochaj, Gold nanoparticles induce nuclear damage in breast cancer cells, which is further amplified by hyperthermia, Cell. Mol. Life Sci. 71 (2014) 4259–4273. [144] M. Kodiha, H. Mahboubi, D. Maysinger, U. Stochaj, Gold nanoparticles impinge on nucleoli and the stress response in MCF7 breast cancer cells, Nanobiomedicine 3 (2016) 3. [145] E. Louvet, H.R. Jun´ era, S. Le Panse, D. Hernandez-Verdun, Dynamics and compartmentation of the nucleolar processing machinery, Exp. Cell Res. 304 (2005) 457–470. [146] A. Pecoraro, A. Virgilio, V. Esposito, A. Galeone, G. Russo, A. Russo, uL3 mediated nucleolar stress pathway as a new mechanism of action of antiproliferative G- quadruplex TBA derivatives in colon cancer cells, Biomolecules 10 (2020). [147] S.A. Klibanov, H.M. O’Hagan, M. Ljungman, Accumulation of soluble and nucleolar-associated p53 proteins following cellular stress, J. Cell Sci. 114 (2001) 1867–1873. [148] M. Loibl, I. Klein, M. Prattes, C. Schmidt, L. Kappel, G. Zisser, A. Gungl, E. Krieger, B. Pertschy, H. Bergler, The drug diazaborine blocks ribosome biogenesis by inhibiting the AAA-ATPase Drg1*, J. Biol. Chem. 289 (2014) 3913–3922. [149] S.A. Kawashima, Z. Chen, Y. Aoi, A. Patgiri, Y. Kobayashi, P. Nurse, T.M. Kapoor, Potent, reversible, and specific chemical inhibitors of eukaryotic ribosome biogenesis, Cell 167 (2016), 512-524.e514. [150] F. Thomas, U. Kutay, Biogenesis and nuclear export of ribosomal subunits in higher eukaryotes depend on the CRM1 export pathway, J. Cell Sci. 116 (2003) 2409–2419. [151] C.E. Lyon, K. Bohmann, J. Sleeman, A.I. Lamond, Inhibition of protein dephosphorylation results in the accumulation of splicing snRNPs and coiled bodies within the nucleolus, Exp. Cell Res. 230 (1997) 84–93. [152] A. Greco, Involvement of the nucleolus in replication of human viruses, Rev. Med. Virol. 19 (2009) 201–214. [153] A. Salvetti, A. Greco, Viruses and the nucleolus: the fatal attraction, Biochim. Biophys. Acta 1842 (2014) 840–847. [154] I. Grummt, R. Voit, Linking rDNA transcription to the cellular energy supply, Cell Cycle (Georgetown, Tex.) 9 (2010) 225–226. [155] K. Mekhail, L. Rivero-Lopez, M. Khacho, S. Lee, Restriction of rRNA synthesis by VHL maintains energy equilibrium under hypoxia, Cell Cycle (Georgetown, Tex.) 5 (2006) 2401–2413. [156] K. Yang, M. Wang, Y. Zhao, X. Sun, Y. Yang, X. Li, A. Zhou, H. Chu, H. Zhou, J. Xu, M. Wu, J. Yang, J. Yi, A redox mechanism underlying nucleolar stress sensing by nucleophosmin, Nat. Commun. 7 (2016) 13599. [157] A.K. Velichko, N.V. Petrova, A.V. Luzhin, O.S. Strelkova, N. Ovsyannikova, N. V. Kireev II, S.V. Petrova, O.L. Kantidze Razin, Hypoosmotic stress induces R loop formation in nucleoli and ATR/ATM-dependent silencing of nucleolar transcription, Nucleic Acids Res. 47 (2019) 6811–6825. [158] L. Yang, J.M. Reece, J. Cho, C.D. Bortner, S.B. Shears, The nucleolus exhibits an osmotically regulated gatekeeping activity that controls the spatial dynamics and functions of nucleolin, J. Biol. Chem. 283 (2008) 11823–11831. [159] F. Frottin, F. Schueder, S. Tiwary, R. Gupta, R. K¨ orner, T. Schlichthaerle, J. Cox, R. Jungmann, F.U. Hartl, M.S. Hipp, The nucleolus functions as a phase-separated protein quality control compartment, Science (New York, N.Y.) 365 (2019) 342–347. [160] T. Xu, S. Liao, M. Huang, C. Zhu, X. Huang, Q. Jin, D. Xu, C. Fu, X. Chen, X. Feng, S. Guang, A ZTF-7/RPS-2 complex mediates the cold-warm response in C. elegans, PLoS Genet. 19 (2023) e1010628. [161] Y. Zhang, G.W. Wolf, K. Bhat, A. Jin, T. Allio, W.A. Burkhart, Y. Xiong, Ribosomal protein L11 negatively regulates oncoprotein MDM2 and mediates a p53- dependent ribosomal-stress checkpoint pathway, Mol. Cell. Biol. 23 (2003) 8902–8912. [162] P.M. Bruno, Y. Liu, G.Y. Park, J. Murai, C.E. Koch, T.J. Eisen, J.R. Pritchard, Y. Pommier, S.J. Lippard, M.T. Hemann, A subset of platinum-containing chemotherapeutic agents kills cells by inducing ribosome biogenesis stress, Nat. Med. 23 (2017) 461–471. [163] E.C. Sutton, C.E. McDevitt, J.Y. Prochnau, M.V. Yglesias, A.M. Mroz, M.C. Yang, R.M. Cunningham, C.H. Hendon, V.J. DeRose, Nucleolar stress induction by oxaliplatin and derivatives, J. Am. Chem. Soc. 141 (2019) 18411–18415. [164] E.C. Sutton, V.J. DeRose, Early nucleolar responses differentiate mechanisms of cell death induced by oxaliplatin and cisplatin, J. Biol. Chem. 296 (2021) 100633. [165] L. Morgado-Palacin, S. Llanos, M. Urbano-Cuadrado, C. Blanco-Aparicio, D. Megias, J. Pastor, M. Serrano, Non-genotoxic activation of p53 through the RPL11-dependent ribosomal stress pathway, Carcinogenesis 35 (2014) 2822–2830. [166] Y.T. Chen, J.J. Chen, H.T. Wang, Targeting RNA polymerase I with hernandonine inhibits ribosomal RNA synthesis and tumor cell growth, Mol. Cancer Res. 17 (2019) 2294–2305. [167] C. Caggiano, E. Guida, F. Todaro, P. Bielli, M. Mori, F. Ghirga, D. Quaglio, B. Botta, F. Moretti, P. Grimaldi, P. Rossi, E.A. Jannini, M. Barchi, S. Dolci, Sempervirine inhibits RNA polymerase I transcription independently from p53 in tumor cells, Cell Death Dis. 6 (2020) 111. K. Gonz´ alez-Arzola BBA - Gene Regulatory Mechanisms 1867 (2024) 195029 19 [168] P.M. Bruno, M. Lu, K.A. Dennis, H. Inam, C.J. Moore, J. Sheehe, S.J. Elledge, M. T. Hemann, J.R. Pritchard, The primary mechanism of cytotoxicity of the chemotherapeutic agent CX-5461 is topoisomerase II poisoning, Proc. Natl. Acad. Sci. U. S. A. 117 (2020) 4053–4060. [169] V. Iadevaia, Y. Huo, Z. Zhang, L.J. Foster, C.G. Proud, Roles of the mammalian target of rapamycin, mTOR, in controlling ribosome biogenesis and protein synthesis, Biochem. Soc. Trans. 40 (2012) 168–172. [170] X. Liang, R. Xie, J. Su, B. Ye, S. Wei, Z. Liang, R. Bai, Z. Chen, Z. Li, X. Gao, Inhibition of RNA polymerase III transcription by Triptolide attenuates colorectal tumorigenesis, J. Exp. Clin. Cancer Res. 38 (2019) 217. [171] J.B. Vines, J.-H. Yoon, N.-E. Ryu, D.-J. Lim, H. Park, Gold nanoparticles for photothermal cancer therapy, Front. Chem. 7 (2019). [172] K. Burger, B. Mühl, M. Rohrmoser, B. Coordes, M. Heidemann, M. Kellner, A. Gruber-Eber, V. Heissmeyer, K. Str¨ asser, D. Eick, Cyclin-dependent kinase 9 links RNA polymerase II transcription to processing of ribosomal RNA, J. Biol. Chem. 288 (2013) 21173–21183. [173] R. Zandomeni, M.C. Zandomeni, D. Shugar, R. Weinmann, Casein kinase type II is involved in the inhibition by 5,6-dichloro-1-beta-D-ribofuranosylbenzimidazole of specific RNA polymerase II transcription, J. Biol. Chem. 261 (1986) 3414–3419. [174] R.M. Martin, G. Ter-Avetisyan, H.D. Herce, A.K. Ludwig, G. L¨ attig-Tünnemann, M.C. Cardoso, Principles of protein targeting to the nucleolus, Nucleus 6 (2015) 314–325. [175] S. Yuan, Z. Liu, Z. Xu, J. Liu, J. Zhang, High mobility group box 1 (HMGB1): a pivotal regulator of hematopoietic malignancies, J. Hematol. Oncol. 13 (2020) 91. [176] M.A. Mensah, H. Niskanen, A.P. Magalhaes, S. Basu, M. Kircher, H.L. Sczakiel, A. M.V. Reiter, J. Elsner, P. Meinecke, S. Biskup, B.H.Y. Chung, G. Dombrowsky, C. Eckmann-Scholz, M.P. Hitz, A. Hoischen, P.-M. Holterhus, W. Hülsemann, K. Kahrizi, V.M. Kalscheuer, A. Kan, M. Krumbiegel, I. Kurth, J. Leubner, A. C. Longardt, J.D. Moritz, H. Najmabadi, K. Skipalova, L. Snijders Blok, A. Tzschach, E. Wiedersberg, M. Zenker, C. Garcia-Cabau, R. Buschow, X. Salvatella, M.L. Kraushar, S. Mundlos, A. Caliebe, M. Spielmann, D. Horn, D. Hnisz, Aberrant phase separation and nucleolar dysfunction in rare genetic diseases, Nature 614 (2023) 564–571. [177] N. Maharjan, S. Saxena, It takes two to tango: DPRs in ALS and SCA36, Neuron 107 (2020) 202–204. [178] O. Sirozh, B. Jung, L. Sanchez-Burgos, I. Ventoso, V. Lafarga, O. Fern´ andez- Capetillo, Nucleolar stress caused by arginine-rich peptides triggers a ribosomopathy and accelerates ageing in mice, bioRxiv (2023), 2023.2008.2010.552792. [179] H.M. Odeh, J. Shorter, Arginine-rich dipeptide-repeat proteins as phase disruptors in C9-ALS/FTD, Emerg. Top. Life Sci. 4 (2020) 293–305. [180] C.J. Lord, A. Ashworth, The DNA damage response and cancer therapy, Nature 481 (2012) 287–294. [181] A. Trenner, A.A. Sartori, Harnessing DNA double-strand break repair for cancer treatment, Front. Oncol. 9 (2019). [182] N. Chatterjee, G.C. Walker, Mechanisms of DNA damage, repair, and mutagenesis, Environ. Mol. Mutagen. 58 (2017) 235–263. [183] A.C. Vítor, P. Huertas, G. Legube, S.F. de Almeida, Studying DNA double-strand break repair: an ever-growing toolbox, Front. Mol. Biosci. 7 (2020). [184] N.A. Tchurikov, D.V. Yudkin, M.A. Gorbacheva, A.I. Kulemzina, I.V. Grischenko, D.M. Fedoseeva, D.V. Sosin, Y.V. Kravatsky, O.V. Kretova, Hot spots of DNA double-strand breaks in human rDNA units are produced in vivo, Sci. Rep. 6 (2016) 25866. [185] M.P. Crossley, M. Bocek, K.A. Cimprich, R-loops as cellular regulators and genomic threats, Mol. Cell 73 (2019) 398–411. [186] H.M. Moore, B. Bai, F.-M. Boisvert, L. Latonen, V. Rantanen, J.C. Simpson, R. Pepperkok, A.I. Lamond, M. Laiho, Quantitative proteomics and dynamic imaging of the nucleolus reveal distinct responses to UV and ionizing radiation*, Mol. Cell. Proteomics 10 (2011) (M111.009241). [187] L.M. Korsholm, Z. G´ al, L. Lin, O. Quevedo, D.A. Ahmad, E. Dulina, Y. Luo, J. Bartek, D.H. Larsen, Double-strand breaks in ribosomal RNA genes activate a distinct signaling and chromatin response to facilitate nucleolar restructuring and repair, Nucleic Acids Res. 47 (2019) 8019–8035. [188] L.M. Korsholm, Z. G´ al, B. Nieto, O. Quevedo, S. Boukoura, C.C. Lund, D.H. Larsen, Recent advances in the nucleolar responses to DNA double-strand breaks, Nucleic Acids Res. 48 (2020) 9449–9461. [189] Shane M. Harding, Jonathan A. Boiarsky, Roger A. Greenberg, ATM dependent silencing links nucleolar chromatin reorganization to DNA damage recognition, Cell Rep. 13 (2015) 251–259. [190] D.H. Larsen, M. Stucki, Nucleolar responses to DNA double-strand breaks, Nucleic Acids Res. 44 (2016) 538–544. [191] R. Scully, A. Panday, R. Elango, N.A. Willis, DNA double-strand break repairpathway choice in somatic mammalian cells, Nat. Rev. Mol. Cell Biol. 20 (2019) 698–714. [192] A. Marnef, A.-L. Finoux, C. Arnould, E. Guillou, V. Daburon, V. Rocher, T. Mangeat, P.E. Mangeot, E.P. Ricci, G. Legube, A cohesin/HUSH- and LINC- dependent pathway controls ribosomal DNA double-strand break repair, Genes Dev. 33 (2019) 1175–1190. [193] D.H. Larsen, F. Hari, J.A. Clapperton, M. Gwerder, K. Gutsche, M. Altmeyer, S. Jungmichel, L.I. Toledo, D. Fink, M.B. Rask, M. Grøfte, C. Lukas, M.L. Nielsen, S.J. Smerdon, J. Lukas, M. Stucki, The NBS1-treacle complex controls ribosomal RNA transcription in response to DNA damage, Nat. Cell Biol. 16 (2014) 792–803. [194] J. Fages, C. Chailleux, J. Humbert, S.M. Jang, J. Loehr, J.P. Lambert, J. Cˆ ot´ e, D. Trouche, Y. Canitrot, JMJD6 participates in the maintenance of ribosomal DNA integrity in response to DNA damage, PLoS Genet. 16 (2020) e1008511. [195] M. Kruhlak, E.E. Crouch, M. Orlov, C. Monta˜ no, S.A. Gorski, A. Nussenzweig, T. Misteli, R.D. Phair, R. Casellas, The ATM repair pathway inhibits RNA polymeraseI transcription in response to chromosome breaks, Nature 447 (2007) 730–734. [196] D.E. Pefani, M.L. Tognoli, D. Pirincci Ercan, V. Gorgoulis, E. O’Neill, MST2 kinase suppresses rDNA transcription in response to DNA damage by phosphorylating nucleolar histone H2B, EMBO J. 37 (2018). [197] C. Mooser, I.E. Symeonidou, P.A. Leimbacher, A. Ribeiro, A.K. Shorrocks, S. Jungmichel, S.C. Larsen, K. Knechtle, A. Jasrotia, D. Zurbriggen, A. Jeanrenaud, C. Leikauf, D. Fink, M.L. Nielsen, A.N. Blackford, M. Stucki, Treacle controls the nucleolar response to rDNA breaks via TOPBP1 recruitment and ATR activation, Nat. Commun. 11 (2020) 123. [198] M. Sun, T. Nishino, J.F. Marko, The SMC1-SMC3 cohesin heterodimer structures DNA through supercoiling-dependent loop formation, Nucleic Acids Res. 41 (2013) 6149–6160. [199] C.T. Carson, R.A. Schwartz, T.H. Stracker, C.E. Lilley, D.V. Lee, M.D. Weitzman, The Mre11 complex is required for ATM activation and the G2/M checkpoint, EMBO J. 22 (2003) 6610–6620. [200] S. Panier, S.J. Boulton, Double-strand break repair: 53BP1 comes into focus, Nat. Rev. Mol. Cell Biol. 15 (2014) 7–18. [201] F. Bonath, J. Domingo-Prim, M. Tarbier, M.R. Friedl¨ ander, N. Visa, Nextgeneration sequencing reveals two populations of damage-induced small RNAs at endogenous DNA double-strand breaks, Nucleic Acids Res. 46 (2018) 11869–11882. [202] M. Uhl´ en, E. Bj¨ orling, C. Agaton, C.A. Szigyarto, B. Amini, E. Andersen, A. C. Andersson, P. Angelidou, A. Asplund, C. Asplund, L. Berglund, K. Bergstr¨ om, H. Brumer, D. Cerjan, M. Ekstr¨ om, A. Elobeid, C. Eriksson, L. Fagerberg, R. Falk, J. Fall, M. Forsberg, M.G. Bj¨ orklund, K. Gumbel, A. Halimi, I. Hallin, C. Hamsten, M. Hansson, M. Hedhammar, G. Hercules, C. Kampf, K. Larsson, M. Lindskog, W. Lodewyckx, J. Lund, J. Lundeberg, K. Magnusson, E. Malm, P. Nilsson, J. Odling, P. Oksvold, I. Olsson, E. Oster, J. Ottosson, L. Paavilainen, A. Persson, R. Rimini, J. Rockberg, M. Runeson, A. Sivertsson, A. Sk¨ ollermo, J. Steen, M. Stenvall, F. Sterky, S. Str¨ omberg, M. Sundberg, H. Tegel, S. Tourle, E. Wahlund, A. Wald´ en, J. Wan, H. Wern´ erus, J. Westberg, K. Wester, U. Wrethagen, L.L. Xu, S. Hober, F. Pont´ en, A human protein atlas for normal and cancer tissues based on antibody proteomics, Mol. Cell. Proteomics 4 (2005) 1920–1932. [203] G. Antoniali, L. Lirussi, M. Poletto, G. Tell, Emerging roles of the nucleolus in regulating the DNA damage response: the noncanonical DNA repair enzyme APE1/Ref-1 as a paradigmatical example, Antioxid. Redox Signal. 20 (2014) 621–639. [204] L. Lirussi, G. Antoniali, C. Vascotto, C. D’Ambrosio, M. Poletto, M. Romanello, D. Marasco, M. Leone, F. Quadrifoglio, K.K. Bhakat, A. Scaloni, G. Tell, Nucleolar accumulation of APE1 depends on charged lysine residues that undergo acetylation upon genotoxic stress and modulate its BER activity in cells, Mol. Biol. Cell 23 (2012) 4079–4096. [205] S.Y. Lee, H. Lee, E.S. Kim, S. Park, J. Lee, B. Ahn, WRN translocation from nucleolus to nucleoplasm is regulated by SIRT1 and required for DNA repair and the development of chemoresistance, Mutat. Res. 774 (2015) 40–48. [206] Y. Mao, I.R. Mehl, M.T. Muller, Subnuclear distribution of topoisomerase I is linked to ongoing transcription and p53 status, Proc. Natl. Acad. Sci. 99 (2002) 1235–1240. [207] J.P. Laine, P.L. Opresko, F.E. Indig, J.A. Harrigan, C. von Kobbe, V.A. Bohr, Werner protein stimulates topoisomerase I DNA relaxation activity, Cancer Res. 63 (2003) 7136–7146. [208] M. Guerra-Rebollo, F. Mateo, K. Franke, M.S. Huen, F. Lopitz-Otsoa, M. S. Rodríguez, V. Plans, T.M. Thomson, Nucleolar exit of RNF8 and BRCA1 in response to DNA damage, Exp. Cell Res. 318 (2012) 2365–2376. [209] M.S.Y. Huen, R. Grant, I. Manke, K. Minn, X. Yu, M.B. Yaffe, J. Chen, RNF8 transduces the DNA-damage signal via histone ubiquitylation and checkpoint protein assembly, Cell 131 (2007) 901–914. [210] R. Roy, J. Chun, S.N. Powell, BRCA1 and BRCA2: different roles in a common pathway of genome protection, Nat. Rev. Cancer 12 (2011) 68–78. [211] M. Engbrecht, A. Mangerich, The nucleolus and PARP1 in cancer biology, Cancers (Basel) 12 (2020). [212] A. Ray Chaudhuri, A. Nussenzweig, The multifaceted roles of PARP1 in DNA repair and chromatin remodelling, Nat. Rev. Mol. Cell Biol. 18 (2017) 610–621. [213] A. Koike, H. Nishikawa, W. Wu, Y. Okada, A.R. Venkitaraman, T. Ohta, Recruitment of phosphorylated NPM1 to sites of DNA damage through RNF8- dependent ubiquitin conjugates, Cancer Res. 70 (2010) 6746. [214] J. Kobayashi, H. Fujimoto, J. Sato, I. Hayashi, S. Burma, S. Matsuura, D.J. Chen, K. Komatsu, Nucleolin participates in DNA double-strand break-induced damage response through MDC1-dependent pathway, PLoS One 7 (2012) e49245. [215] D.D. Scott, M. Oeffinger, Nucleolin and nucleophosmin: nucleolar proteins with multiple functions in DNA repair, Biochem. Cell Biol. 94 (2016) 419–432. [216] M. Poletto, L. Lirussi, D.M. Wilson, G. Tell, Nucleophosmin modulates stability, activity, and nucleolar accumulation of base excision repair proteins, Mol. Biol. Cell 25 (2014) 1641–1652. [217] C. Yang, M.S. Kim, D. Chakravarty, F.E. Indig, F. Carrier, Nucleolin binds to the proliferating cell nuclear antigen and inhibits nucleotide excision repair, Mol. Cell. Pharmacol. 1 (2009) 130–137. K. Gonz´ alez-Arzola BBA - Gene Regulatory Mechanisms 1867 (2024) 195029 20 [218] X. Duan, S.G. Kelsen, A.B. Clarkson Jr., R. Ji, S. Merali, SILAC analysis of oxidative stress-mediated proteins in human pneumocytes: new role for treacle, Proteomics 10 (2010) 2165–2174. [219] Z. Zhao, M.A. Dammert, S. Hoppe, H. Bierhoff, I. Grummt, Heat shock represses rRNA synthesis by inactivation of TIF-IA and lncRNA-dependent changes in nucleosome positioning, Nucleic Acids Res. 44 (2016) 8144–8152. [220] Z. Zhao, Marcel A. Dammert, I. Grummt, H. Bierhoff, lncRNA-induced nucleosome repositioning reinforces transcriptional repression of rRNA genes upon hypotonic stress, Cell Rep. 14 (2016) 1876–1882. [221] Z. Zhao, N. Sentürk, C. Song, I. Grummt, lncRNA PAPAS tethered to the rDNA enhancer recruits hypophosphorylated CHD4/NuRD to repress rRNA synthesis at elevated temperatures, Genes Dev. 32 (2018) 836–848. [222] S. Feng, A. Desotell, A. Ross, M. Jovanovic, J.L. Manley, A nucleolar long “noncoding” RNA encodes a novel protein that functions in response to stress, Proc. Natl. Acad. Sci. U. S. A. 120 (2023) e2221109120. [223] Y. Zhou, K.M. Schmitz, C. Mayer, X. Yuan, A. Akhtar, I. Grummt, Reversible acetylation of the chromatin remodelling complex NoRC is required for noncoding RNA-dependent silencing, Nat. Cell Biol. 11 (2009) 1010–1016. [224] K.-M. Schmitz, C. Mayer, A. Postepska, I. Grummt, Interaction of noncoding RNA with the rDNA promoter mediates recruitment of DNMT3b and silencing of rRNA genes, Genes Dev. 24 (2010) 2264–2269. [225] C. Mayer, K.M. Schmitz, J. Li, I. Grummt, R. Santoro, Intergenic transcripts regulate the epigenetic state of rRNA genes, Mol. Cell 22 (2006) 351–361. [226] M.S. Lindstr¨ om, D. Jurada, S. Bursac, I. Orsolic, J. Bartek, S. Volarevic, Nucleolus as an emerging hub in maintenance of genome stability and cancer pathogenesis, Oncogene 37 (2018) 2351–2366. [227] N.J. Yoo, Y.R. Kim, S.H. Lee, Somatic mutation of PHF6 gene in T-cell acute lymphoblatic leukemia, acute myelogenous leukemia and hepatocellular carcinoma, Acta Oncol. (Stockholm, Sweden) 51 (2012) 107–111. [228] T. Kobayashi, Regulation of ribosomal RNA gene copy number and its role in modulating genome integrity and evolutionary adaptability in yeast, Cell. Mol. Life Sci. 68 (2011) 1395–1403. [229] P.A. Defossez, R. Prusty, M. Kaeberlein, S.J. Lin, P. Ferrigno, P.A. Silver, R.L. Keil, L. Guarente, Elimination of replication block protein Fob1 extends the life span of yeast mother cells, Mol. Cell 3 (1999) 447–455. [230] D.A. Sinclair, K. Mills, L. Guarente, Accelerated aging and nucleolar fragmentation in yeast sgs1 mutants, Science (New York, N.Y.) 277 (1997) 1313–1316. [231] A. Buchwalter, M.W. Hetzer, Nucleolar expansion and elevated protein translation in premature aging, Nat. Commun. 8 (2017) 328. [232] J. Hallgren, M. Pietrzak, G. Rempala, P.T. Nelson, M. Hetman, Neurodegeneration-associated instability of ribosomal DNA, Biochim. Biophys. Acta 1842 (2014) 860–868. [233] V. Tiku, A. Antebi, Nucleolar function in lifespan regulation, Trends Cell Biol. 28 (2018) 662–672. [234] R.A. MacLeod, D. Spitzer, I. Bar-Am, J.E. Sylvester, M. Kaufmann, A. Wernich, H. G. Drexler, Karyotypic dissection of Hodgkin’s disease cell lines reveals ectopic subtelomeres and ribosomal DNA at sites of multiple jumping translocations and genomic amplification, Leukemia 14 (2000) 1803–1814. [235] C. Guetg, P. Lienemann, V. Sirri, I. Grummt, D. Hernandez-Verdun, M.O. Hottiger, M. Fussenegger, R. Santoro, The NoRC complex mediates the heterochromatin formation and stability of silent rRNA genes and centromeric repeats, EMBO J. 29 (2010) 2135–2146. [236] D.O. Warmerdam, J. van den Berg, R.H. Medema, Breaks in the 45S rDNA lead to recombination-mediated loss of repeats, Cell Rep. 14 (2016) 2519–2527. K. Gonz´ alez-Arzola