The Secret Lives of Transcription Factors: In Heterochromatin Regulation 3031290275, 9783031290275

Table of contents :
Contents
Chapter 1: HP1 and Noncanonical Functions of STAT
STAT
HP1
Summary
References
Chapter 2: Biological Functions of STAT, HP1, and Heterochromatin
Heterochromatin Loss During Aging
Heterochromatin Redistribution in Cellular Senescence
Cellular Senescence Versus Cancer Development
STAT and HP1 Protect Against DNA Damage and Promote Longevity
References
Chapter 3: Other Transcription Factors with Noncanonical Functions in Heterochromatin Regulation
GAGA Factor
Rb
ATF2
NF-κB
PAX
C/EBPα
References
Chapter 4: HP1 in Liquid–Liquid Phase Separation and its Regulation by 53BP1
Liquid–Liquid Phase Separation and HP1
53BP1
53BP1 and Proteins in DNA Damage Response
Cell Cycle Phase-Dependent DNA Damage Repair
Other Pathways in DNA Damage Response
53BP1 in Heterochromatin Regulation
References
Index

Citation preview

SpringerBriefs in Biochemistry and Molecular Biology Willis X. Li · Louise Silver-Morse

The Secret Lives of Transcription Factors In Heterochromatin Regulation

SpringerBriefs in Biochemistry and Molecular Biology

SpringerBriefs in Biochemistry and Molecular Biology are a series of slim high quality publications encompassing the entire spectrum of biochemistry and molecular biology. Each individual proposal will be evaluated by hand-picked external reviewers to ensure publication of Springer Briefs of the highest standard. SpringerBriefs published in the series represent the cutting edge in biochemistry and molecular biology research in a format intersecting the traditional review and the scientific book volume.

Willis X. Li • Louise Silver-Morse

The Secret Lives of Transcription Factors In Heterochromatin Regulation

Willis X. Li Department of Medicine University of California, San Diego La Jolla, CA, USA

Louise Silver-Morse Department of Biomedical Genetics University of Rochester Medical Center Rochester, NY, USA

ISSN 2211-9353     ISSN 2211-9361 (electronic) SpringerBriefs in Biochemistry and Molecular Biology ISBN 978-3-031-29027-5    ISBN 978-3-031-29029-9 (eBook) https://doi.org/10.1007/978-3-031-29029-9 © Springer Nature Switzerland AG 2023 This work is subject to copyright. All rights are solely and exclusively licensed by the Publisher, whether the whole or part of the material is concerned, specifically the rights of translation, reprinting, reuse of illustrations, recitation, broadcasting, reproduction on microfilms or in any other physical way, and transmission or information storage and retrieval, electronic adaptation, computer software, or by similar or dissimilar methodology now known or hereafter developed. The use of general descriptive names, registered names, trademarks, service marks, etc. in this publication does not imply, even in the absence of a specific statement, that such names are exempt from the relevant protective laws and regulations and therefore free for general use. The publisher, the authors, and the editors are safe to assume that the advice and information in this book are believed to be true and accurate at the date of publication. Neither the publisher nor the authors or the editors give a warranty, expressed or implied, with respect to the material contained herein or for any errors or omissions that may have been made. The publisher remains neutral with regard to jurisdictional claims in published maps and institutional affiliations. This Springer imprint is published by the registered company Springer Nature Switzerland AG The registered company address is: Gewerbestrasse 11, 6330 Cham, Switzerland

Contents

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 HP1 and Noncanonical Functions of STAT ������������������������������������������    1 STAT����������������������������������������������������������������������������������������������������������    1 HP1������������������������������������������������������������������������������������������������������������    4 Summary����������������������������������������������������������������������������������������������������    6 References��������������������������������������������������������������������������������������������������    7

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 Biological Functions of STAT, HP1, and Heterochromatin������������������   13 Heterochromatin Loss During Aging��������������������������������������������������������   13 Heterochromatin Redistribution in Cellular Senescence ��������������������������   15 Cellular Senescence Versus Cancer Development ������������������������������������   16 STAT and HP1 Protect Against DNA Damage and Promote Longevity������������������������������������������������������������������������������   16 References��������������������������������������������������������������������������������������������������   17

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Other Transcription Factors with Noncanonical Functions in Heterochromatin Regulation��������������������������������������������������������������   19 GAGA Factor ��������������������������������������������������������������������������������������������   19 Rb��������������������������������������������������������������������������������������������������������������   21 ATF2����������������������������������������������������������������������������������������������������������   23 NF-κB��������������������������������������������������������������������������������������������������������   24 PAX������������������������������������������������������������������������������������������������������������   25 C/EBPα������������������������������������������������������������������������������������������������������   27 References��������������������������������������������������������������������������������������������������   28

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HP1 in Liquid–Liquid Phase Separation and its Regulation by 53BP1 ��������������������������������������������������������������������������������������������������   37 Liquid–Liquid Phase Separation and HP1������������������������������������������������   37 53BP1��������������������������������������������������������������������������������������������������������   38 53BP1 and Proteins in DNA Damage Response ��������������������������������������   38

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Contents

Cell Cycle Phase-Dependent DNA Damage Repair����������������������������������   39 Other Pathways in DNA Damage Response����������������������������������������������   40 53BP1 in Heterochromatin Regulation������������������������������������������������������   40 References��������������������������������������������������������������������������������������������������   42 Index������������������������������������������������������������������������������������������������������������������   45

Chapter 1

HP1 and Noncanonical Functions of STAT

Abstract  The past decades have seen an explosion of research on chromatin and epigenetic regulation and their roles in modulating gene expression. Studies on histone modifications, chromatin remodeling complexes, chromatin/chromatin interactions, chromatin interactions with the nuclear architecture, and the interaction of transcription factors with other nuclear proteins have underscored the complexity of transcriptional regulation of gene expression. Emerging evidence has shown that some conventional transcription factors, such as signal transducer and activator of transcription (STAT), also affect gene expression in unconventional ways and have functions that are not directly related to the modulation of gene expression. In particular, it has been shown that a fraction of cellular STAT proteins resides in heterochromatin. This chapter reviews the noncanonical function of STAT in heterochromatin regulation in relation to heterochromatin protein 1 (HP1), the central player in heterochromatin regulation. Keywords  HP1 · STAT · Heterochromatin · Noncanonical STAT · Unphosphorylated STAT (uSTAT) · Epigenetics

STAT The components of the canonical Janus kinase/signal transducer and activator of transcription (JAK/STAT) signaling pathway were delineated in the late 1980s and early 1990s (Darnell Jr. et  al. 1994) (Fig.  1.1). In humans, seven STAT proteins (STAT1–4, 5a, 5b, and 6) and four JAKs (JAK1–3 and tyrosine kinase 2) have been identified, while Drosophila has one JAK and one STAT. Dictyostelium appears to have four STATs (Kawata et al. 1997; Zhukovskaya et al. 2004; Gao et al. 2004) and no JAKs (Goldberg et  al. 2006). The JAK/STAT pathway has not been found in plants. In mammals and Drosophila, the signaling pathway is similar; upon binding

© The Author(s), under exclusive license to Springer Nature Switzerland AG 2023 W. X. Li, L. Silver-Morse, The Secret Lives of Transcription Factors, SpringerBriefs in Biochemistry and Molecular Biology, https://doi.org/10.1007/978-3-031-29029-9_1

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Fig. 1.1  Canonical JAK/STAT signaling. A schematic representation of the canonical JAK/STAT pathway is shown. In the canonical pathway, unphosphorylated STAT (uSTAT) is formant and resides in the cytoplasm. The pathway is triggered by extracellular ligand binding to membrane receptors, which leads to the activation of receptor-associated JAK kinase, which in turn activates STAT by phosphorylation, converting uSTAT to pSTAT. Dimerized pSTAT proteins translocate into the nucleus, where they function as transcription factors to induce the transcription of target genes

of extracellular ligands to transmembrane receptors, the receptors dimerize and activate receptor-associated JAKs, which in turn phosphorylate tyrosine residues in the cytoplasmic tails of the receptors. These phosphotyrosine residues serve as docking sites for cytoplasmic STAT proteins, which JAKs then phosphorylate on a crucial C-terminal tyrosine residue. Phosphorylated STAT (pSTAT) proteins dimerize and translocate to the nucleus, where they function as conventional transcriptional regulators (Li 2008). Though no JAKs have been found in Dictyostelium, there are data suggesting that at least one of the Dictyostelium STATs, DdSTATa, also functions as a conventional transcriptional regulator (Kawata et al. 1997; Mohanty et al. 1999; Fukuzawa and Williams 2000; Shimada et al. 2004; Ginger et al. 2000). In 2008, Shi et al. reported that unphosphorylated Drosophila STAT has additional functions. It is found in the nucleus as well as in the cytoplasm. In the nucleus, it maintains heterochromatin stability in association with HP1 (Shi et al. 2008). Shi et al. further showed that unphosphorylated STAT (uSTAT), but not pSTAT, physically interact with HP1, and that uSTAT and HP1 colocalize at the telomeres and centromeres of polytene chromosomes (Fig. 1.2). Thus, STAT is the first conventional transcription factor found to be associated with heterochromatin.

STAT

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Fig. 1.2  Non-canonical STAT function. Drosophila salivary gland cells were immunostained with antibodies to HP1, total STAT, or pSTAT. Confocal images of single cells were shown for localization of HP1 (red), STAT (green, upper panel), or pSTAT (green, lower panel). Heterochromatin is recognized as HP1-enriched regions in the nucleus. Note that total STAT is unevenly distributed in the nucleus, with the concentrated regions colocalized with HP1, as indicated by the yellow color in the merged image; and that pSTAT is not colocalized with HP1. (The figure is adapted from Shi et al. (2008))

uSTATs—STAT1, STAT3, and low levels of STAT5a—have been found in the nucleus of mammalian cells too (Meyer et al. 2002; Liu et al. 2005; Iyer and Reich 2008). Moreover, the Li lab has extended the findings of Shi et  al. to mammals, reporting that HP1α and unphosphorylated STAT5a and STAT3 are associated in mammalian cells and enhance heterochromatin stability (Hu et al. 2013; Dutta et al. 2020). Furthermore, it has been demonstrated that this enhancement of heterochromatin stability suppresses cancer formation in a murine model system (Hu et  al. 2013; Dutta et  al. 2020). Since STATs are conserved evolutionarily from slime molds to humans (Kawata et al. 1997), it would be interesting to determine whether uSTAT associates with HP1 homologs in Dictyostelium. In addition to uSTATs, it appears that JAK2 can be found in the nuclei of mammalian cells. Specifically, nuclear JAK2 has been detected in oocytes (Ito et  al. 2004) and hematopoietic cells (Dawson et al. 2009). Nuclear JAK2 has also probably been detected in hepatocytes (Lobie et al. 1996; Ram and Waxman 1997), and perhaps in alpha and beta cells of pancreatic islets (Sorenson and Stout 1995). Immunostaining of unfertilized mouse oocytes, and one-cell and two-cell mouse embryos, has shown JAK2 to be localized to chromatin during interphase, meiosis, and mitosis; by the four-cell stage, nuclear JAK2 is no longer detectable (Ito et al. 2004). In human hematopoietic cells, a function for nuclear JAK2 has been identified; it specifically phosphorylates histone H3 at amino acid Y41, preventing its association with HP1α (Dawson et al. 2009). Since unphosphorylated STAT5a is

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associated with HP1α in mammalian cells and stabilizes heterochromatin, nuclear JAK activity may destabilize heterochromatin in at least two ways: by decreasing the association of STAT5a with HP1α and by decreasing the binding of HP1α to chromatin. It is not known if there is nuclear JAK in Drosophila. Notable among a number of proteins that interact with STATs (Shuai 2000) are the protein inhibitors of activated STATs (PIAS) proteins, which bind to pSTAT dimers, preventing DNA recognition (Chung et  al. 1997; Betz et  al. 2001). Four PIAS genes have been identified in mammals (Shuai 2000); there is one in Drosophila, the Su(var)2–10, or dPIAS gene (Betz et al. 2001); and one PIAS-like gene has been found in Dictyostelium (Kawata et al. 2011). Intriguingly, mutations in Drosophila Su(var)2–10 result in chromosome condensation defects as well as aberrant segregation of chromosomes during anaphase, though the dPIAS protein does not seem to associate with mitotic chromosomes; furthermore, the polytene chromosomes of Su(var)2–10 mutants appear disorganized (Hari et  al. 2001). dPIAS is normally found around the periphery of diploid nuclei in Drosophila, and in intranuclear spots, while in polytene chromosomes it is found at the chromocenter and some telomeres (Hari et al. 2001). It is not known whether the role of dPIAS in chromosome organization involves STAT.

HP1 Heterochromatin protein 1 (HP1), as its name implies, has been associated with heterochromatin ever since the protein was first identified in Drosophila (James and Elgin 1986; Schoelz and Riddle 2022). Four more isoforms were subsequently found in flies, encoded by four additional genes; the original HP1 is now known as HP1a (Vermaak et al. 2005). The HP1 family of proteins is evolutionarily conserved with Swi6 and Chp2 characterized in fission yeast (Lorentz et al. 1994; Thon and Verhein-Hansen 2000), LHP1  in the plant Arabidopsis thaliana (Gaudin et  al. 2001), and HP1α, HP1β, and HP1γ isoforms identified in humans and mice (Singh et al. 1991). Drosophila HP1a has been a focus of research on the HP1 family of proteins. Immunostaining experiments have shown that HP1a localizes at pericentromeric heterochromatin and telomeres (James et al. 1989; Kellum et al. 1995). In line with these results, Drosophila embryos carrying loss-of-function HP1a mutations show defects in chromosome segregation, indicating that HP1a is required for normal mitoses (Kellum and Alberts 1995). Moreover, HP1a is required for telomere capping, with telomere elongation and telomeric fusions observed in the absence of HP1a (Savitsky et al. 2002; Fanti et al. 1998; Perrini et al. 2004). Transcriptional repression in telomeres requires HP1a as well; the repression is alleviated in the absence of HP1a (Perrini et al. 2004). In view of the fact that HP1a associates with heterochromatin, it is not surprising that loss-of-function HP1a mutations suppress position effect variegation (Eissenberg et al. 1992). In mammals, too, HP1 localizes to heterochromatin (Aagaard et al. 1999).

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However, despite its strong association with heterochromatin, HP1a is also associated with nearly 200 euchromatic sites on salivary gland polytene chromosomes (Piacentini et al. 2003; De Lucia et al. 2005; de Wit et al. 2007; Piacentini et al. 2009). These sites include a number of heat shock puffs, and other sites of active transcription. Extensive colocalization of HP1a and an active form of RNA polymerase II have been demonstrated at such sites. Further support for the idea that HP1a has a positive role in euchromatic gene transcription comes from the finding that transcript levels of HP1a-associated euchromatic genes decrease upon lowering of the HP1a dosage. Thus, HP1a can function as a positive regulator of transcription in a euchromatic environment. Nonetheless, HP1a is not a conventional transcriptional regulator. Recombinant HP1a does not recognize specific DNA sequences, though it binds to Drosophila genomic DNA in vitro (Zhao et al. 2001). Its positive effect on transcription is thought to result from its binding, possibly via other proteins, to nascent transcripts at the site of transcription, and its stabilization of the transcripts (Piacentini et al. 2003; Piacentini et al. 2009). Consistent with its many known functions and perhaps some unknown functions, HP1a and its homologs associate with a very diverse group of chromatinassociated proteins. There is evidence that HP1a in Drosophila interacts directly with the histone methyltransferase Su(var)3–9 (Schotta et al. 2002), the histone H3K36 demethylase dKDM4A (Lin et al. 2008), Su(var)3–7 (Delattre et al. 2000; Cléard et al. 1997), the STAT transcription factor (Shi et al. 2008), and the origin recognition complex (ORC), specifically the ORC1 subunit (Pak et al. 1997). In mammalian systems, HP1α has been shown to associate directly with the nuclear receptor-associated transcription intermediary factor TIFα (Le Douarin et  al. 1996), the KRAB domain-binding KAP-1 corepressor (also known as TIFβ) (Ryan et al. 1999; Le Douarin et al. 1996), the inner nuclear membrane lamin B receptor (Ye and Worman 1996; Ye et al. 1997), the p150 subunit of the histone H3 and H4 chaperone chromatin assembly factor 1 (CAF-1) (Murzina et al. 1999), the SP100 protein found in nuclear bodies (nuclear speckles) (Seeler et al. 1998; Lehming et al. 1998), the retinoblastoma protein (Rb) (Nielsen et al. 2001), and histone H3 via H3mK9 (Lachner et al. 2001; Bannister et al. 2001), and human STAT3, 5 (Dutta et al. 2020; Hu et al. 2013). Mammalian HP1α can also associate with the avian inner centromere protein (INCENP), a component of the mitotic chromosome scaffold, when it is transfected into mammalian cells (Ainsztein et al. 1998). Evolutionary conservation has been demonstrated for the interaction of HP1 with Orc1 and with H3K9. Xenopus HP1α associates with Xenopus Orc1 (Pak et al. 1997), and the HP1 homolog in fission yeast, Swi6, interacts with an H3K9-containing peptide (Bannister et al. 2001). The extent to which the other interactions are evolutionarily conserved is not known.

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Summary Studies have shown a noncanonical mode of JAK/STAT signaling, in which uSTAT plays a role in maintaining heterochromatin stability (Shi et al. 2008; Hu et al. 2013) (Fig. 1.3), although the molecular mechanism remains unclear. Studies by other groups have suggested the involvement of many different transcription factors in regulating heterochromatin as noncanonical functions (to be discussed in other chapters). It is hypothesized that these transcription factors can have noncanonical functions in initiating and/or maintaining heterochromatin formation. To initiate heterochromatin formation, a transcription factor such as uSTAT binds to DNA sequences and recruits HP1, thus “seeding” heterochromatin formation (Fig.  1.4a). To maintain heterochromatin stability, transcription factors such as uSTAT bind to and form a complex with HP1 proteins that are already in heterochromatin, stabilizing the HP1-nucleosome association in heterochromatin (Fig. 1.4b).

Fig. 1.3  Non-canonical JAK/STAT signaling. A schematic representation of the non-canonical mode of JAK/STAT signaling, in which a portion of the unphosphorylated STAT (uSTAT) pool is localized in the nucleus on heterochromatin in association with HP1 in the nucleus. Phosphorylation of STAT by JAK reduces the amount of uSTAT in heterochromatin, leading to HP1 dispersal and heterochromatin instability. Phosphorylated STAT (pSTAT) in turn binds to cognitive sites in euchromatin to induce the expression of target genes

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Fig. 1.4  Possible roles of protein factors binding to HP1 in heterochromatin dynamics. Schematic illustrations of heterochromatin initiation (a) and maintenance (b) in the presence of HP1 and an associating factor (e.g., uSTAT). Protein factors such as uSTAT can have two functions in heterochromatin regulation. (a) Establishing heterochromatin by binding to DNA then recruiting HP1 to “seed” the initial heterochromatin formation, and (b) maintaining heterochromatin stability by associating with HP1 to form a protein complex. In this case, reducing the affinity between the protein factor and HP1, such as by STAT phosphorylation, leads to heterochromatin instability

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Liu L, McBride KM, Reich NC (2005) STAT3 nuclear import is independent of tyrosine phosphorylation and mediated by importin-alpha3. Proc Natl Acad Sci U S A 102(23):8150–8155. http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?cmd=Retrieve&db=PubMed&dopt=Citation& list_uids=15919823 Lobie PE, Ronsin B, Silvennoinen O, Haldosén LA, Norstedt G, Morel G (1996) Constitutive nuclear localization of Janus kinases 1 and 2. Endocrinology 137(9):4037–4045. https://doi. org/10.1210/endo.137.9.8756581. https://www.ncbi.nlm.nih.gov/pubmed/8756581 Lorentz A, Ostermann K, Fleck O, Schmidt H (1994) Switching gene swi6, involved in repression of silent mating-type loci in fission yeast, encodes a homologue of chromatin-­ associated proteins from drosophila and mammals. Gene 143(1):139–143. https://doi. org/10.1016/0378-­1119(94)90619-­x. https://www.ncbi.nlm.nih.gov/pubmed/8200530 Meyer T, Gavenis K, Vinkemeier U (2002) Cell type-specific and tyrosine phosphorylation-­ independent nuclear presence of STAT1 and STAT3. Exp Cell Res 272(1):45–55. https://doi. org/10.1006/excr.2001.5405. https://www.ncbi.nlm.nih.gov/pubmed/11740864 Mohanty S, Jermyn KA, Early A, Kawata T, Aubry L, Ceccarelli A, Schaap P, Williams JG, Firtel RA (1999) Evidence that the Dictyostelium Dd-STATa protein is a repressor that regulates commitment to stalk cell differentiation and is also required for efficient chemotaxis. Development 126(15):3391–3405. https://www.ncbi.nlm.nih.gov/pubmed/10393118 Murzina N, Verreault A, Laue E, Stillman B (1999) Heterochromatin dynamics in mouse cells: interaction between chromatin assembly factor 1 and HP1 proteins. Mol Cell 4(4):529–540. https:// doi.org/10.1016/s1097-­2765(00)80204-­x. https://www.ncbi.nlm.nih.gov/pubmed/10549285 Nielsen SJ, Schneider R, Bauer UM, Bannister AJ, Morrison A, O'Carroll D, Firestein R, Cleary M, Jenuwein T, Herrera RE, Kouzarides T (2001) Rb targets histone H3 methylation and HP1 to promoters. Nature 412(6846):561–565. http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?cmd =Retrieve&db=PubMed&dopt=Citation&list_uids=11484059 Pak DT, Pflumm M, Chesnokov I, Huang DW, Kellum R, Marr J, Romanowski P, Botchan MR (1997) Association of the origin recognition complex with heterochromatin and HP1 in higher eukaryotes. Cell 91(3):311–323. S0092-8674(00)80415-8 [pii]. http://www.ncbi.nlm.nih.gov/ pubmed/9363940 Perrini B, Piacentini L, Fanti L, Altieri F, Chichiarelli S, Berloco M, Turano C, Ferraro A, Pimpinelli S (2004) HP1 controls telomere capping, telomere elongation, and telomere silencing by two different mechanisms in drosophila. Mol Cell 15(3):467–476. https://doi.org/10.1016/j.molcel.2004.06.036. S1097276504003818 [pii]. http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?c md=Retrieve&db=PubMed&dopt=Citation&list_uids=15304225 Piacentini L, Fanti L, Berloco M, Perrini B, Pimpinelli S (2003) Heterochromatin protein 1 (HP1) is associated with induced gene expression in drosophila euchromatin. J Cell Biol 161(4):707–714. http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?cmd=Retrieve&db=PubMed &dopt=Citation&list_uids=12756231 Piacentini L, Fanti L, Negri R, Del Vescovo V, Fatica A, Altieri F, Pimpinelli S (2009) Heterochromatin protein 1 (HP1a) positively regulates euchromatic gene expression through RNA transcript association and interaction with hnRNPs in drosophila. PLoS Genet 5(10):e1000670. https://doi.org/10.1371/journal.pgen.1000670. http://www.ncbi.nlm.nih.gov/ pubmed/19798443 Ram PA, Waxman DJ (1997) Interaction of growth hormone-activated STATs with SH2-­ containing phosphotyrosine phosphatase SHP-1 and nuclear JAK2 tyrosine kinase. J Biol Chem 272(28):17694–17702. http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?cmd=Retrieve& db=PubMed&dopt=Citation&list_uids=9211920 Ryan RF, Schultz DC, Ayyanathan K, Singh PB, Friedman JR, Fredericks WJ, Rauscher FJ (1999) KAP-1 corepressor protein interacts and colocalizes with heterochromatic and euchromatic HP1 proteins: a potential role for Krüppel-associated box-zinc finger proteins in heterochromatin-­mediated gene silencing. Mol Cell Biol 19(6):4366–4378. https://doi. org/10.1128/mcb.19.6.4366. https://www.ncbi.nlm.nih.gov/pubmed/10330177 Savitsky M, Kravchuk O, Melnikova L, Georgiev P (2002) Heterochromatin protein 1 is involved in control of telomere elongation in Drosophila melanogaster. Mol Cell Biol 22(9):3204–3218. https://doi.org/10.1128/mcb.22.9.3204-­3218.2002. https://www.ncbi.nlm.nih.gov/ pubmed/11940677

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Chapter 2

Biological Functions of STAT, HP1, and Heterochromatin

Abstract  A conventional transcription factor may be defined as a protein that binds to a particular DNA sequence or a set of sequences and interacts directly with the basal transcriptional machinery to control or modulate gene expression. It is also possible to define a transcription factor more broadly to include proteins that bind to a particular DNA sequence, but interact indirectly with the basal transcriptional machinery or proteins that bind to DNA indirectly or in a nonsequence-specific manner and interact either directly or indirectly with the basal transcriptional machinery. These proteins can be thought of as unconventional transcription factors. There is evidence that certain unconventional transcription factors, such as HP1, play important roles in regulating heterochromatin formation and the epigenome, thereby influencing the global transcriptome, chromosome behavior, and genome integrity, with implications for aging and cancer development. Keywords  Transcription factor · Heterochromatin · Aging · Cancer · Senescence

Heterochromatin Loss During Aging Research has suggested that during animal aging, total heterochromatin levels in cells gradually decline, and this is accompanied by the redistribution of certain heterochromatin domains (Tsurumi and Li 2012; Tsurumi and Li 2020; Villeponteau 1997; Lee et al. 2020) (Fig. 2.1). On the one hand, erosion of heterochromatin in genes normally repressed due to differentiation leads to their ectopic expression, with possible detrimental consequences to cells. It has been shown that heterochromatin formation plays an essential role in maintaining cellular identity and function including maintenance of HSC identity (Koide et al. 2016). On the other hand, aberrant heterochromatinization could occur at genes that should be expressed, leading to their abnormal repression, resulting in loss of functionality of differentiated cell types. Furthermore, heterochromatin formation plays an essential role in maintaining cellular identity and function including maintenance of HSC identity (Koide © The Author(s), under exclusive license to Springer Nature Switzerland AG 2023 W. X. Li, L. Silver-Morse, The Secret Lives of Transcription Factors, SpringerBriefs in Biochemistry and Molecular Biology, https://doi.org/10.1007/978-3-031-29029-9_2

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Fig. 2.1  Heterochromatin levels decline with aging. (a) Gut tissues from young (3-day old) and old (35-day old) female flies (Drosophila melanogaster) were dissected and stained with anti-HP1 (magenta), which correlates with heterochromatin levels. Images were scanned at identical settings with confocal microscopy. Note that HP1 forms prominent foci in the young gut (one pointed by an arrow), whereas in the old gut tissues, HP1 staining seems more diffuse and lacks the prominent foci (Larson et al. 2012). (b) Percent survival of adult female flies (Drosophila melanogaster) of the indicated genotypes at 25oC. Flies had been made coisogenic by extensive outcrossing before the aging experiment. n donates the number of flies counted. p values are from Log rank analysis. Flies carrying one copy of hsp70-HP1 (expressing more HP1) were longer lived (p = 6.31 × 10–24), and flies heterozygous for Su(var)2055 (loss-of-function allele of the HP1a gene) lived shorter (p = 2.03 × 10–86) when compared with wild-type controls (indicated as “+/+”) (Larson et al. 2012). (The panels in this figure are adapted from Larson et al. (2012))

Heterochromatin Redistribution in Cellular Senescence

15

et al. 2016). Indeed, it has been shown that the levels of Suv39h2 and/or Suv39h1 methyltransferases decrease with age in hematopoietic stem cells (HSCs) and this is associated with a loss of B cell differentiation and with hematopoietic changes characteristic of aging (Djeghloul et al. 2016; Keenan et al. 2020). Studies have suggested that heterochromatin formation may promote longevity and that aging is associated with gradual loss of heterochromatin. The human premature aging diseases, Hutchinson–Gilford progeria syndrome (HGPS) and atypical Werner syndrome, are associated with germline mutations in lamins that cause expression of a truncated mutant form of lamin A called progerin (Shumaker et al. 2006). Cells from a HGPS patient exhibit abnormal nuclear morphology with much-­ reduced heterochromatin at the nuclear periphery, accompanied by a decrease in the heterochromatin marker H3K9me3 and a delocalization of HP1α (Shumaker et al. 2006). The global loss of heterochromatin markers and changes in nuclear morphology are also seen in human cells during normal physiological aging (Scaffidi and Misteli 2006). Similar changes in nuclear architecture and peripheral heterochromatin were also found during aging of Caenorhabditis elegans (Haithcock et  al. 2005) and Drosophila (Larson et  al. 2012; Brandt et  al. 2008; Chen et  al. 2016; Yan et  al. 2011). By examining bulk genome levels of histone modifications including H3K4me3 and the repressive heterochromatin marker H3K9me3 and HP1a binding to chromatin using whole-genome chromatin immunoprecipitation at different ages in Drosophila (Wood et  al. 2010), the authors found a striking reduction in both H3K9me3 and HP1a enrichment at constitutive heterochromatin, usually found in pericentric regions and the fourth chromosome, in older flies compared to younger flies. These studies suggest that the mechanisms underlying animal aging might be evolutionarily conserved.

Heterochromatin Redistribution in Cellular Senescence Studies of cellular senescence, a state of irreversible cell cycle arrest, have focused on the formation of senescence-associated heterochromatin foci (SAHF), which are domains of facultative heterochromatin involving H3K9me2/3 and HP1α that accumulate in senescent cells (Aird and Zhang 2013). Cellular senescence marks a replicative or growth arrest initially observed in cultured cells and is believed to be a mechanism underlying organismic aging (Hayflick and Moorhead 1961; Pawlikowski et al. 2013). It has been proposed that SAHF formation plays a role in cellular senescence in part by sequestering and repressing proliferation-promoting genes, including the E2F target gene cyclin A (Narita et  al. 2003). Interestingly, removing senescent cells from aged mice delayed their aging process and improved the aging phenotypes in adipose, skeletal muscle and eye tissues (Villeponteau 1997; Baker et  al. 2011), suggesting that cellular senescence may be a driver of organismic aging. However, the precise biological functions of SAHF and the mechanisms of their formation remain incompletely understood. It has also been

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reported that SAHF formation is dispensable for cellular senescence (Kosar et al. 2011) and that cellular senescence is associated with loss of heterochromatin (Zhang et al. 2021). Thus, the increase in SAHF in senescent cells may result from heterochromatin redistribution, amid an overall decline in heterochromatin levels with age progression.

Cellular Senescence Versus Cancer Development Cancer and aging are highly correlated, and cancer has been considered an age-­ related disease. The relationship between cancer and aging is complex. One important aspect is that age-related changes in cells may contribute to microenvironments permissive for cancer development (Fane and Weeraratna 2020). For instance, mechanistic studies of cellular senescence have shown that the P16INK4a tumor suppressor and senescence marker play a central role in inducing cellular senescence and cancer development (Schmitt et al. 2002). Interestingly, it has been shown that at moderate expression levels, P16INK4a is required for tumor suppression, hence the name. However, at high expression levels, it induces cell cycle arrest, leading to cellular senescence. Senescent cells may in turn influence the surrounding tissues by secreting factors that induce chronic inflammation, contributing to organismic aging (Baker et al. 2011; Sato et al. 2015).

 TAT and HP1 Protect Against DNA Damage S and Promote Longevity One important function of heterochromatin is to protect genome integrity (Allshire and Madhani 2018; Janssen et al. 2018). Loss of heterochromatin may thus increase chromosomal aberrations and DNA damage with age, whereas strengthening or increasing heterochromatin alleviates genomic damage and delays aging (Muñoz-­ Najar and Sedivy 2011; Benayoun et al. 2015; Yan et al. 2011). In addition, studies have shown that heterochromatin formation is important for suppression of transposable elements, and therefore loss of heterochromatin may result in remobilization of transposable elements, whose reinsertion into the genome may lead to mutagenic events (Benayoun et al. 2015; Muñoz-Najar and Sedivy 2011; Larson et al. 2012). Interestingly, it has been shown that overexpressing HP1a and uSTAT in flies can counter the global loss of heterochromatin in older flies, increasing their life span and delaying their loss of muscle integrity, whereas mutant flies with reduced HP1a (and heterochromatin) exhibit decreased life span and loss of muscle integrity (Larson et al. 2012). Furthermore, HP1a and heterochromatinization have also been shown to be important for maintaining appropriate levels of ribosomal RNA (rRNA)

References

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production, demonstrating the intimate link between heterochromatin and cellular metabolism. Moreover, this study has shown that uSTAT, demonstrated previously to promote heterochromatin stability (Shi et al. 2006, 2008), behaved similarly to HP1a in delaying or reversing of the aging phenotypes.

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Larson K, Yan SJ, Tsurumi A, Liu J, Zhou J, Gaur K, Guo D, Eickbush TH, Li WX (2012) Heterochromatin formation promotes longevity and represses ribosomal RNA synthesis. PLoS Genet 8(1):e1002473. https://doi.org/10.1371/journal.pgen.1002473 Lee JH, Kim EW, Croteau DL, Bohr VA (2020) Heterochromatin: an epigenetic point of view in aging. Exp Mol Med 52(9):1466–1474. https://doi.org/10.1038/s12276-­020-­00497-­4 Muñoz-Najar U, Sedivy JM (2011) Epigenetic control of aging. Antioxid Redox Signal 14(2):241–259. https://doi.org/10.1089/ars.2010.3250 Narita M, Nunez S, Heard E, Narita M, Lin AW, Hearn SA, Spector DL, Hannon GJ, Lowe SW (2003) Rb-mediated heterochromatin formation and silencing of E2F target genes during cellular senescence. Cell 113(6):703–716 Pawlikowski JS, Adams PD, Nelson DM (2013) Senescence at a glance. J Cell Sci 126(Pt 18):4061–4067. https://doi.org/10.1242/jcs.109728 Sato S, Kawamata Y, Takahashi A, Imai Y, Hanyu A, Okuma A, Takasugi M, Yamakoshi K, Sorimachi H, Kanda H, Ishikawa Y, Sone S, Nishioka Y, Ohtani N, Hara E (2015) Ablation of the p16(INK4a) tumour suppressor reverses ageing phenotypes of klotho mice. Nat Commun 6:7035. https://doi.org/10.1038/ncomms8035 Scaffidi P, Misteli T (2006) Lamin A-dependent nuclear defects in human aging. Science 312(5776):1059–1063. doi:1127168 [pii]. https://doi.org/10.1126/science.1127168 Schmitt CA, Fridman JS, Yang M, Lee S, Baranov E, Hoffman RM, Lowe SW (2002) A senescence program controlled by p53 and p16INK4a contributes to the outcome of cancer therapy. Cell 109(3):335–346. https://doi.org/10.1016/s0092-­8674(02)00734-­1 Shi S, Calhoun HC, Xia F, Li J, Le L, Li WX (2006) JAK signaling globally counteracts heterochromatic gene silencing. Nat Genet 38(9):1071–1076 Shi S, Larson K, Guo D, Lim SJ, Dutta P, Yan SJ, Li WX (2008) Drosophila STAT is required for directly maintaining HP1 localization and heterochromatin stability. Nat Cell Biol 10(4):489–496 Shumaker DK, Dechat T, Kohlmaier A, Adam SA, Bozovsky MR, Erdos MR, Eriksson M, Goldman AE, Khuon S, Collins FS, Jenuwein T, Goldman RD (2006) Mutant nuclear Lamin a leads to progressive alterations of epigenetic control in premature aging. Proc Natl Acad Sci U S A 103(23):8703–8708. doi:0602569103 [pii]. https://doi.org/10.1073/pnas.0602569103 Tsurumi A, Li WX (2012) Global heterochromatin loss: a unifying theory of aging? Epigenetics 7(7):680–688. https://doi.org/10.4161/epi.20540 Tsurumi A, Li WX (2020) Aging mechanisms—a perspective mostly from Drosophila. Adv Genet 2(1):e10026. https://doi.org/10.1002/ggn2.10026 Villeponteau B (1997) The heterochromatin loss model of aging. Exp Gerontol 32(4–5):383–394. doi:S0531-5565(96)00155-6 [pii] Wood JG, Hillenmeyer S, Lawrence C, Chang C, Hosier S, Lightfoot W, Mukherjee E, Jiang N, Schorl C, Brodsky AS, Neretti N, Helfand SL (2010) Chromatin remodeling in the aging genome of drosophila. Aging Cell 9(6):971–978. https://doi.org/10.1111/j.1474-­9726.2010.00624.x Yan SJ, Lim SJ, Shi S, Dutta P, Li WX (2011) Unphosphorylated STAT and heterochromatin protect genome stability. FASEB J 25(1):232–241. https://doi.org/10.1096/fj.10-­169367 Zhang X, Liu X, Du Z, Wei L, Fang H, Dong Q, Niu J, Li Y, Gao J, Zhang MQ, Xie W, Wang X (2021) The loss of heterochromatin is associated with multiscale three-dimensional genome reorganization and aberrant transcription during cellular senescence. Genome Res 31:1121. https://doi.org/10.1101/gr.275235.121

Chapter 3

Other Transcription Factors with Noncanonical Functions in Heterochromatin Regulation

Abstract  Past research in the regulation of gene expression has led to the observation that many conventional and unconventional transcription factors can physically reside in heterochromatin and in other intriguing nuclear locations. What are these proteins doing in these locations? Though a definitive answer is not presently available, consideration of the question may be instructive based on what we have learned from STAT and HP1. This chapter reviews unconventional functions for transcription factors GAGA, Rb, NF-κB, ATF2, PAX, and C/EBPα involved in heterochromatin regulation, as listed in Table 3.1. The list by no means exhausts all transcription factors with potential noncanonical epigenetic functions. It is possible that noncanonical functions in heterochromatin regulation may emerge for even more transcription factors in the future. Keywords  GAGA · Rb · NF-κB · ATF2 · PAX · C/EBPα

GAGA Factor In 1988, Biggin and Tjian reported the purification of the GAGA transcription factor from Drosophila embryos (Biggin et al. 1988). GAGA factor (GAF) recognizes and binds to (CT⋅GA)n sequences in the promoter regions of the Ultrabithorax, engrailed, hsp70, hsp26, histone H3, and histone H4 genes, among others, activating gene expression (Biggin et al. 1988; Lu et al. 1993; Wilkins and Lis 1997; van Steensel et al. 2003), and was at first considered to be a conventional transcriptional activator. However, there is no evidence that GAF directly recruits the transcriptional machinery. Rather GAF is involved in creating nucleosome-free sites, and in chromatin remodeling with the help of nucleosome remodeling factor (NURF), and in these ways seems to alleviate inhibitory effects of chromatin (Croston et al. 1991; Tsukiyama et al. 1994; Tsai et al. 2016; Fuda et al. 2015; Lehmann 2004). Using an in vitro nucleosome system based on the hsp70 promoter, Tsukiyama et al. showed that GAF binding leads to nucleosome disruption, DNase hypersensitivity at the © The Author(s), under exclusive license to Springer Nature Switzerland AG 2023 W. X. Li, L. Silver-Morse, The Secret Lives of Transcription Factors, SpringerBriefs in Biochemistry and Molecular Biology, https://doi.org/10.1007/978-3-031-29029-9_3

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Table 3.1  Protein factors implicated in heterochromatin regulation Protein factor HP1 STAT GAGA Rb NF-κB ATF-2 PAX3/9 C/EBPα 53BP1

Function in heterochromatin Central component of heterochromatin Associates with HP1 Binds to DNA sequences in heterochromatin Heterochromatin maintenance Recruit G9a methyltransferase Heterochromatin localization and maintenance Associates with satellite repeats in heterochromatin Associates with and HP1α Involved in HP1 liquid-liquid phase separation

Reference Schoelz 2022 Shi, 2008 Raff 1994 Gonzalo 2005 Chen 2009 Seong et al. 2011 Bulut-Karslioglu 2012 Siegel, 2013 Zhang et al. 2022

TATA box and heat shock elements, and an energy-dependent rearrangement of adjacent nucleosomes (Tsukiyama et  al. 1994). Further support for the idea that GAF alleviates inhibitory effects of chromatin comes from the finding that loss-offunction mutations in GAF enhance position effect variegation (PEV) in Drosophila (Farkas et al. 1994). Thus, GAF is a somewhat unconventional transcription factor, which is thought to prime genes such as hsp70 for rapid induction by establishing an open chromatin architecture in the promoter region (Chetverina et  al. 2021) (Table 3.1). However, GAF has other functions in addition to establishing or maintaining an open chromatin structure in the regulatory regions of particular genes. In 1994, Raff et al. reported that GAF associates with regions of heterochromatin in Drosophila embryos, including centromeric regions, where immunostaining was observed during the entire cell cycle (Raff et al. 1994). GAF associates with centromeric regions in larvae as well, but at this developmental stage the association occurs primarily during mitosis (Platero et al. 1998). The heterochromatic regions with which GAF associates share a similar distribution of GA/CT-rich repeats. The revelation that GAF associates with heterochromatin as well as euchromatin was followed by the demonstration that GAF is required for proper nuclear division in early Drosophila embryos (Bhat et al. 1996). Early embryos depend upon GAF derived from maternal mRNA stored in the oocyte. At the syncytial blastoderm stage, embryos from mothers carrying a hypomorphic mutation in the Trithorax-like gene, which encodes GAF, showed asynchrony in nuclear cleavage cycles, failures in chromosome condensation, abnormal chromosome segregation, and chromosomal fragmentation; heterochromatin-associated GAF was not seen in these embryos. These data suggest a more global role for GAF in chromosome structure and function. GAF is found in a number of heteromeric, chromatin-associated protein complexes (Lehmann 2004). Its association with the ATP-dependent NURF, the heterodimeric FACT complex, a trxG complex, and polycomb repressive complex 1 (PRC1), presumably explains how GAF can perform diverse functions. In 2010, Matharu et al. reported the identification of GAF homologs in humans, mice, and zebra fish (Matharu et al. 2010).

Rb

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Rb The retinoblastoma (Rb) protein was identified in 1987 as the product of the human Rb gene, whose deletion or mutation is associated with Rbs and other cancers (Lee et al. 1987; Murphree 1984). Rb is a highly conserved protein, with homologs found in a number of animal and plant species, including Caenorhabditis elegans (Lu and Horvitz 1998), Drosophila (Du et  al. 1996; Stevaux et  al. 2002), Xenopus laevis (Destrée et al. 1992), chicken (Boehmelt et al. 1994), mouse (Bernards et al. 1989), maize (Grafi et al. 1996; Xie et al. 1996; Ach et al. 1997) and other plants (Durfee et  al. 2000), and the plantlike single-celled alga Chlamydomonas (Umen and Goodenough 2001). In mammalian cells, Rb has a role in regulating progression through the cell cycle, particularly the G1 phase (Goodrich, 1991). Rb in hypophosphorylated form binds to E2F transcription factors early in G1, repressing transcription of E2F target genes implicated in cell cycle progression; phosphorylation of Rb results in its dissociation from E2F, relieving the block to progression through G1 (Chellappan et al. 1991; Weintraub et al. 1992). There is evidence that Rb functions similarly in Drosophila (Du et al. 1996; Stevaux et al. 2002) and plants (Ach et al. 1997; Huntley et al. 1998; Nakagami et al. 2002; Park et al. 2005; Shimizu-Sato et al. 2008). Rb is also involved in regulating progression through the cell cycle in Chlamydomonas, where it ensures that cells have reached an appropriate size before proceeding from G1 to S phase (Umen and Goodenough 2001). Moreover, in endoreduplicating Drosophila ovarian follicle cells and tobacco leaf cells, Rb functions in the regulation of endocycles, in which cells undergo multiple rounds of DNA replication without mitoses. Suppression of Rb results in extra DNA replication (Bosco et al. 2001; Park et al. 2005), and consistently with these data, Rb undergoes increased phosphorylation as endoreduplication is initiated in maize endosperm (Grafi et al. 1996). Rb is another unconventional transcription factor in that it does not bind directly to DNA but binds indirectly by interacting with DNA-binding proteins. Rb is often associated with transcriptional repression, which it effects by binding to E2F and other conventional transcription factors, and by recruiting chromatin-modifying enzymes such as histone deacetylase (HDAC1) to target promoters (Brehm et al. 1998; Magnaghi-Jaulin et al. 1998). Rb has also been shown to recruit HP1 and H3 methylase to repress gene transcription (Nielsen et al. 2001). Rb also appears to be involved in transcriptional activation. It is required for MyoD-driven differentiation of cells in culture to a muscle phenotype (Gu et al. 1993) and for C/EBP-driven differentiation of cultured cells to an adipocyte phenotype (Chen et al. 1996), and it can upregulate transcriptional activation by c-jun (Nead et al. 1998), hBrm (Singh et al. 1995), and ATF2 (Kim et al. 1992). However, it is not clear how Rb boosts the transcription of genes that are positively regulated by these transcription factors. While Rb can bind to each of these factors, with MyoD, C/EBP, c-jun, and hBrm binding specifically to hypophosphorylated Rb, Rb does not seem to be a part of the transcription-activating protein complexes.

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More recently, Rb has been shown to have additional functions. In 2005, Gonzalo et al. reported that mouse embryonic fibroblasts deficient in RB1, RBL1, and RBL2 exhibited aneuploidy, and aberrant, abnormally enlarged centromeres and elongated telomeres, as well as a reduction in H4K20 methylation at pericentromeric and telomeric heterochromatin (Gonzalo et al. 2005). DNA methylation at major satellite DNA was also decreased in the triple knockout cells. Thus, the loss of these Rb proteins resulted in global defects in constitutive heterochromatin. Both the heterochromatic defects and the reduction in H4K20 methylation occurred independently of the E2F family of transcriptional regulators, as determined by viral transduction of a dominant-negative DP, an E2F cofactor. In 2008, Longworth et al. reported similar findings in Drosophila (Longworth et al. 2008). Fused or broken chromosomes could be observed in rbf1 mutant cells, with anaphase bridges in which chromosomes failed to separate normally; and in rbf1 mutant larvae, chromosomes often showed regions of hypocondensation during early prophase or prometaphase. This rbf1 phenotype in Drosophila was also independent of dDP. As this phenotype would predict, overexpression of RBF1 in Drosophila wing imaginal discs resulted in chromatin hypercondensation. Interestingly, the hypercondensation phenotype was suppressed by heterozygous mutation of genes encoding particular subunits of the condensin II complex: dCap­D3, dCap-H2, or dCap-G. Rbf1 and dCAP-D3 were shown to colocalize at a number of loci on polytene chromosomes, though being largely excluded from constitutively heterochromatic loci. While the exclusion from polytene constitutive heterochromatin is unexpected given that in mouse fibroblasts the loss of Rb family proteins resulted in defects in constitutive heterochromatin, it should be noted that localization of Rbs to constitutive heterochromatin in mouse cells has not actually been demonstrated (Gonzalo et al. 2005). Longworth et al. extended their observations to human cells, where they showed that RB promoted localization of CAP-D3 to chromatin. Manning et  al. subsequently demonstrated that when nontransformed human cells were depleted of RB, the levels of centromere-associated CAP-D3 and the cohesin protein Rad21 decreased (Manning et al. 2010). These cells exhibited an increase in intercentromeric distance, a deformation of centromeric structure, defects in chromosome alignment at the metaphase plate, reduced sister chromatin cohesion, and an increased aneuploidy. Cells from Drosophila Rb mutant larvae were also examined, and showed sister chromatid cohesion defects and increased aneuploidy. Similarly, Coschi et  al. reported that decreased levels of chromatin-­ associated CAP-D3 in primary embryonic fibroblasts from Rb mutant mice were correlated with a looser chromosome alignment at the metaphase plate, and increased aneuploidy (Coschi et al. 2010). Furthermore, Chen et al. found meiotic defects in Arabidopsis plants carrying a mutation in RBR, the single Arabidopsis Rb-related gene (Chen et al. 2011). Chiasma formation was reduced, with subsequent errors in chromosome disjunction, leading to aneuploidy and reduced fertility. RBR mutant chromosomes also appeared to be longer than wild type, suggesting a defect in condensation.

ATF2

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Rb has been reported to interact with more than one hundred different proteins, though a direct association has been demonstrated only for some of them (Morris and Dyson 2001). Of particular interest with regard to possible Rb functions in addition to transcriptional regulation, is a direct interaction with the human origin recognition complex subunit Orc1, and the Drosophila Orc complex (Mendoza-Maldonado et al. 2010; Bosco et al. 2001). Notably, the Orc complex in Drosophila has also been shown to interact with HP1 (Pak et al. 1997). Rb may also bind directly to human DNA replication initiation factor MCM7 (Sterner et  al. 1998), primate nuclear matrix-associated protein p84 (Durfee et al. 2000), primate lamin A/C (Mancini et  al. 1994), 1994], human lamin-associated polypeptide 2α (Markiewicz et  al. 2002), and human and Drosophila CAP-D3. (Longworth et al. 2008).

ATF2 Activating transcription factor 2 (ATF2), also called cyclic AMP response element (CRE)-binding protein 2 (CREB2), was first identified in 1989 in a yeast screen for CRE-binding proteins (Maekawa et al. 1989). ATF2 has been known for its important functions in mediating response to stress and other extracellular stimuli. The human ATF2 gene, located on chromosome 2q32, is expressed in almost all cell types and is likely required for normal organismal development. In a mouse model, total loss of the ATF2 gene results in postnatal lethality; ATF2 partial loss-­ of-­function mutations lead to developmental abnormalities including neurological defect and early mortality (Maekawa et al. 1999). Several isoforms of ATF2 exist due to alternative splicing, and these alternative splice isoforms display diverse functions. One truncated isoform, ATF2SV5, which lacks amino acids from 210 to 505 (full length is 505aa), was originally expected to be transcriptionally inactive. However, it is expressed and has oncogenic effects in cooperation with metastasis-­ related genes such as CCL4, CCR7, S100A8, and BrafV600E(Claps et al. 2016). A small ATF2 isoform, ATF2-sm, which consists of only four of the necessary exons and lacks the basic leucine zipper domain, is differentially expressed and has particular effects on genes involved in pregnancy and labor (Bailey et al. 2002). ATF2 is regulated at multiple levels and has various cellular functions. ATF2 transcripts can be regulated by RNA-binding proteins and miRNAs. For example, miR-622, miR-451, miR26b, and miR204 have been shown to bind to the 3’UTR of ATF2 mRNA for its degradation under different physiological situations (Arora et al. 2011; Song et al. 2016; Zhang et al. 2015). Conversely, ELAV-like protein 1, an RNA-binding protein, can bind to the 3’ UTR of ATF2 mRNA to stabilize it (Arora et al. 2011). ATF2 is posttranslationally modified for function, and multiple extracellular signals, including UV radiation, cytokines, and growth factors, can lead to ATF2 modification and activation (Lopez-Bergami et al. 2010). It has been shown that ATF2 is activated by stress-activated kinases via phosphorylation at T69 and T71 (Gupta et al. 1995). Phosphorylation of these two threonine residues

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enables ATF2 to interact with other AP1 proteins or to enter the nucleus (Hayakawa et al. 2004). ATF2 is phosphorylated by the protein kinase ATM on serines 490 and 498 following ionizing radiation (Bhoumik et  al. 2005). ATF2 can also be phosphorylated by PKCε at T52, allowing accumulation of ATF2 at the outer mitochondrial membrane, resulting in breach in membrane integrity and eventually cell death (Lau et al. 2012). The ATF2 pool at the mitochondrial membrane interferes with multiple ion channels including VDAC1 and hexokinase-1. In addition, acetylation can also affect the transcriptional activity of ATF2. For instance, the histone acetyltransferase p300 HAT can acetylase ATF2 at the basic leucine zipper domain at Lys 357 and 374, enhancing the activity of ATF2 (Karanam et al. 2007). Thus, while the abundance of ATF2 is regulated at the mRNA and protein level, multiple posttranslational modifications enhance or alter ATF2 function by impacting the structure, localization, and stability of the protein. As a stress-response factor, ATF2 has been shown to be required for heterochromatin regulation in response to cellular stresses. In fission yeast, the ATF2 homolog binds to the silent mating-type locus and recruits histone deacetylases and histone methyltransferase to initiate heterochromatin formation in response to stresses (Jia et al. 2004; Kim et al. 2004). In Drosophila, the human ATF2 homolog, dATF-2, is also required for heterochromatin assembly, with extensive colocalization with HP1. However, heat shock or osmotic stress induces MAPK-mediated phosphorylation of dATF-2 and its release from heterochromatin, resulting in heterochromatin disruption (Seong et al. 2011). The effect of phosphorylation on ATF2 heterochromatin regulation is reminiscent of its effect on STAT (Li 2008; Shi et  al. 2008), although further studies are needed to understand the molecular mechanisms underlying phosphorylation effects on noncanonical functions of transcription factors in heterochromatin regulation.

NF-κB Nuclear factor-kappa B (NF-κB) is a family of transcription factors involved in regulating inflammatory and immune responses, including both innate and adaptive immune functions, as well as cell proliferation and survival (Oeckinghaus and Ghosh 2009; Liu et al. 2017; Taniguchi and Karin 2018). NF-κB family transcription factors include p50 (NF-κB1), p52, p65, RelA, RelB, and c-Rel that regulate immune or inflammatory responses via transcriptionally activating expression of a number of cytokines and chemokines. In the absence of extracellular stimuli, members of the NF-κB group are sequestered in the cytoplasm by inhibitor of κB (IκB) proteins, which contain multiple ankyrin repeats that bind to and block the nuclear localization signal of NF-κB proteins (Jacobs and Harrison 1998; Liu et al. 2017). Upon stimulation and release from IκB, NF-κB proteins enter the nucleus, where they regulate target gene transcription by binding to the κB site present in the enhancer regions of many genes involved in innate and adaptive immune responses (Sun et al. 2013).

PAX

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NF-κB activation is mediated through two different signaling pathways: canonical and noncanonical (Sun 2017). The canonical pathway is involved in diverse immune regulations, including innate and inflammatory immune responses, as well as lymphocyte activation and differentiation. Activation of the canonical NF-κB pathway is initiated at cytokine receptors, pattern-recognition receptors, T/B cell receptors, and TNF receptors. Following receptor activation, the IKK (IκB kinase) complex phosphorylates IκBα at two N-terminal serines, resulting in its degradation and the subsequent nuclear translocation of NF-κB, such as p50/p65 dimers (Hayden and Ghosh 2008). The noncanonical NF-κB pathway, on the other hand, is only activated in response to certain stimuli, such as ligands of the TNFR family, including RANK, BAFFR, CD40, and LTBR, and it is mainly involved in the development and maturation of immune cells (Sun 2017). Unlike the canonical pathway, the noncanonical pathway does not involve IκBα degradation and only participates in adaptive immune responses and act as a supplementary signaling pathway to the canonical one. The noncanonical NF-κB pathway relies on processing of p100, an NF-κB2 precursor protein. It is initiated by the NF-κB-inducing kinase activating IKK complex and inducing p100 phosphorylation, which results in p100 ubiquitination and processing. P100 processing degrades its C-terminal IκB-like structure, leading to NF-κB2 p52 generation and translocalization of the p52/RelB dimer into the nucleus (Sun 2017). NF-κB activation has been associated with many types of cancers and has therefore been targeted for cancer therapy (Taniguchi and Karin 2018). Although whether and how NF-κB activation causes cancer remains unresolved, studies have implicated epigenetic changes linked to NF-κB activation. For instance, a study of NF-κB inhibition with a parthenolide analog has found extensive changes in epigenetic modifications (Nakshatri et al. 2015). Parthenolide is an NF-κB inhibitor and has potent anticancer as well as anti-inflammatory activities (Ghantous et al. 2013). In particular, the authors found that inhibition of NF-κB using inhibitor DMAPT (a parthenolide analog) resulted in increased histone H3K36 trimethylation and H4K20 trimethylation. On the other hand, Rel-B of the NF-κB family was found to directly associate with the H3 lysine 9 methyltransferase G9a and induce facultative heterochromatin formation thereby resulting in silencing of several proinflammatory genes (Chen et al. 2009). The formation of facultative heterochromatin at the interleukin-1beta promoter required occupancy of G9a, which further recruits and complexes with HP1 for transcriptional silencing. Thus, while NF-κB plays critical roles in inducing proinflammatory genes in acute immune response, it may also induce heterochromatin formation to limit severe systemic inflammation in humans by silencing proinflammatory genes.

PAX The paired box (PAX) family of genes encode evolutionarily conserved tissue-­ specific transcription factors with important functions in tissue specification and organogenesis during early animal development (Thompson et  al. 2021;

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Paixão-­Côrtes et al. 2015; Monsoro-Burq 2015; Blake and Ziman 2014; Stuart et al. 1994). First identified in Drosophila as a family of homologs to the paired gene essential for segmentation and embryonic development (Bopp et al. 1986), members of the PAX family were found across phylogenies, each containing a PAX domain, which is a DNA-binding domain of approximately 130 a.a. sequence that binds to DNA with the TCACGC/G motif (Paixão-Côrtes et  al. 2015; Jun and Desplan 1996). Human and mouse each contains 9 PAX family protein transcription factors—PAX1 to PAX9. These proteins are often referred to as “master regulators” because they play essential roles in determination and maintenance of tissue and cell types. Mutations in PAX genes are associated with many human diseases, such as acute lymphoblastic leukemia (ALL), hypothyroidism, microphthalmia, to name a few (Thompson et al. 2021). Studies in human genetics and mouse models have highlighted the essential roles of PAX genes in the development of multiple organs, including the developing nervous system and musculature (Blake and Ziman 2014). PAX3 is well characterized in for its function and expression in early development, including skeletal muscle and neural crest development, migration, and diseases related to these cell lineages (Boudjadi et al. 2018; Buckingham and Relaix 2007). Mutant alleles in the mouse Pax3 gene, with different strengths, and the use of transgenic animals facilitated characterization of Pax3 function in neural crest development. Mouse Pax3 expression is detected early during the migration of neural crest cells (NCCs), and also in somitic cells associated with the migratory NCCs, and its expression extinguishes as these NCCs differentiate (Serbedzija and McMahon 1997). Germline mutations in the Pax3 gene are associated with splotch phenotypes in mice (Tremblay et  al. 1995; Epstein et  al. 2000) and with an autosomal-­ dominant combination of auditory and pigment defects in human Waardenburg syndrome (Baldwin et al. 1992; Read and Newton 1997). On the other hand, persistent Pax3 expression in NCCs is associated with developmental defects and lethality. One of the Pax3 target genes is Sostdc1, a soluble inhibitor of bone morphogenetic protein (BMP), whose upregulation inhibits BMP-induced osteogenesis, thereby resulting in cleft palate in the mouse (Wu et  al.  2008). Thus, in addition to specifying neural crest cell types, Pax3 also functions to maintain an undifferentiated state in the cells by counteracting differentiation signals. PAX9 is best known for its essential function in craniofacial and tooth development (Bonczek et al. 2017). A recent study using cultured human cells has shown that PAX9 regulates small subunit ribosome biogenesis (Farley-Barnes et al. 2020). Studies in the mouse have found that Pax9, like Pax3, is essential for controlling neural crest development, and is expressed early in many mesenchymal cells derived from NCCs (Kist et al. 2007). The expression of Pax9 in cranial mesenchymal cells, including those in the nose, palate, and teeth, is consistent with its essential functions in craniofacial and tooth development. Indeed, Pax9 homozygous-null mice die at birth, while deletion of Pax9 specifically in the neural crest results in mice with cleft palate and defects in tooth development (Kist, Greally, and Peters). Studies employing mouse genetics have shown that Pax9 is required for the development of a wide range of organs, including the thymus, parathyroid glands, ultimobranchial bodies, teeth, and skeletal elements (Peters et al. 1998).

C/EBPα

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Interestingly, Pax3 and Pax9 have been found to be redundant regulators of mouse heterochromatin (Bulut-Karslioglu et al. 2012). Heterochromatin formation plays key roles in maintaining genome integrity by repressing transcription from repetitive sequences, including major satellite repeats (Allshire and Madhani 2018). Mouse Pax3 and Pax9 have both been found to associate with DNA within major satellite repeats in pericentric heterochromatin, thereby repressing transcription from these sequences (Bulut-Karslioglu et  al.  2012). Consistent with such roles, PAX-binding sites are found in major satellite repeat sequences and are associated with the enrichment of the heterochromatin mark, H3K9me3. Simultaneous depletion of Pax3 and Pax9 resulted in the impairment of heterochromatin structure and in chromosome segregation defects (Bulut-Karslioglu et  al.  2012). The observed loss of heterochromatin marks and the derepression of RNA transcripts from major satellite repeats are reminiscent of STAT3 knockdown results (Dutta et al. 2020). Thus, PAX transcription factors also seem to have noncanonical functions in heterochromatin regulation.

C/EBPα The CCAAT/enhancer-binding protein α (C/EBPα) is an essential transcription factor of the leucine zipper family, which is involved in various systems (Wang et al. 2019). The CEBPA gene coding for C/EBPα is intronless and resides on chromosome 19.q13.1 (Schmidt et al. 2020). There are two isoforms of C/EBPα, a shorter isoform (p30) with only one transactivation element and a longer one (p42) with three transactivation elements (Muruganandan et al. 2020). Both p30 and p42 contain the basic-region leucine zipper (bZIP) and could dimerize with other proteins in the C/EBP family (Muruganandan et al. 2020; Wang et al. 2019). As the name suggests, C/EBPα would bind to the CCAAT box motif that is present in multiple promoters. Furthermore, the bZIP region of C/EBPα is important for specific DNA binding, which could be regulated by dimerization because C/EBPα could bind to DNA as either homodimer or heterodimer (Wang et  al. 2019). Previous research suggests that C/EBPα is mainly involved in biochemical activities in blood cells and could act as an adipogenic transcription factor, a hematopoietic transcription factor, as well as an osteoclastogenic transcription factor (Muruganandan et al. 2020). The role of C/EBPα in osteoclastogenic signaling is to some extent similar to that of PPARγ (Muruganandan et  al. 2020; Jules et  al. 2018). Generally, C/EBPα is involved in maintaining regular function, lineage commitment, as well as differentiation of osteoclastogenic cells. C/EBPα could bind to DNA and activate the transcription of several transcription factors and proteins functionally required for osteoblasts, including a vacuolar proton pump (Chen et  al.  2013). The signaling pathway is triggered by a paracrine mechanism. When the motif IVVY of RANK ligand is present, C/EBPα is activated and then it induces transcription of downstream genes including NF-κB, NFATc1, and c-fos (Chen et al. 2013). When RANK ligands are not present, C/EBPα is sufficient to induce lineage commitment.

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However, the differentiation of osteoblasts would be largely impaired in the absence of RANK ligand. Moreover, C/EBPα participated in the function of osteoblasts since it could contribute to the extracellular acidification process (Muruganandan et al. 2020). Interestingly, previous studies have shown that C/EBPα could colocalize with pericentromeric heterochromatin (Schaufele et al. 2003). At transcriptionally inactive state, the master regulator C/EBPα is concentrated at pericentromeric heterochromatin regions in the mouse cell nucleus (Schaufele et al. 2003). Imaging studies on living cells suggest that C/EBPα assumes a dimer structure, as deduced from fluorescence resonance energy transfer (FRET) between the C/EBPα proteins tagged with different fluorescent moieties (Schaufele et al. 2003). Using the same FRET technique, the authors have further shown that the bZIP DNA-binding domains of C/EBPα dimers were further apart when they are at pericentromeric heterochromatin than when they are at the euchromatic regions of the nucleus. Thus, it was concluded that the conformation of C/EBPα varies with intranuclear location and with cellular environment. Furthermore, the authors concluded that the accumulation of C/EBPα in pericentromeric heterochromatin is functionally important and can be regulated, and was observed in diverse cell lineage (Schaufele et  al. 2003). Noticeably, C/EBPα can bind to highly repetitive satellite DNA sequences, and these sequences can reduce the interaction between C/EBPα and its cognate DNA motif (Liu et al. 2007). These studies suggest that C/EBPα forms dimers in living cells, and its conformation varies with subnuclear location with the cellular environment. More recent studies have shown that C/EBPα and HP1α may directly interact (Sun et al. 2010; Demarco et al. 2006; Sun et al. 2011). Further, a study using combination of deletion, mutagenesis, and FRET has demonstrated that the BZip domain of C/EBPα is responsible for the interaction with HP1α in heterochromatin (Siegel et al. 2013). Thus, heterochromatin dynamics, which plays important roles in epigenetic regulation, is controlled by a network of protein interactions mediated by the heterochromatin protein 1 (HP1) (Table 3.1).

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Chapter 4

HP1 in Liquid–Liquid Phase Separation and its Regulation by 53BP1

Abstract  Recent research has suggested a novel role of HP1 in heterochromatin formation. It has been shown that HP1 proteins can oligomerize to form liquid–liquid phase separation (LLPS), resulting in the formation of membraneless liquid droplets of proteins. It has been proposed that LLPS be a mechanism of organizing functional centers in the nucleus. LLPS forms when the concentration of certain proteins reaches a threshold, separating from the surrounding solution by phase separation, like oil in water. Importantly, the LLPS process of HP1 can be regulated by interacting with other proteins. Tumor suppressor p53-binding protein 1 (53BP1) has been shown recently to be among protein factors involved in regulating LLPS formation by HP1. This chapter summarizes the current understanding of LLPS in HP1-mediated heterochromatin formation, and the regulation of this process, using association with tumor suppressor 53BP1 as an example. Keywords  liquid–liquid phase separation (LLPS) · Heterochromatin Protein 1 (HP1) · Tumor Suppressor p53-Binding Protein 1 (53BP1)

Liquid–Liquid Phase Separation and HP1 Recent research has demonstrated that HP1 can form condensates via liquid–liquid phase separation (LLPS), and it has been proposed that this property of HP1 is important for its role in heterochromatin formation (Strom et al. 2017). Experiments in vitro have shown that both Drosophila HP1a and human HP1α can form higher-­ order oligomerization and phase-separated droplets (Larson et al. 2017). The presence of DNA and other proteins can promote or reverse the phase-separation process (Larson et al. 2017). In addition, it has been shown that phase-separated droplets compartmentalize heterochromatin and dynamically expose buried nucleosomal regions (Sanulli et al. 2019). Hypothetically, the reshape of the nucleosome core is correlated with histone modification and could increase multivalent interactions between nucleosomes, promoting phase separation (Sanulli et al. 2019). Furthermore, © The Author(s), under exclusive license to Springer Nature Switzerland AG 2023 W. X. Li, L. Silver-Morse, The Secret Lives of Transcription Factors, SpringerBriefs in Biochemistry and Molecular Biology, https://doi.org/10.1007/978-3-031-29029-9_4

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4  HP1 in Liquid–Liquid Phase Separation and its Regulation by 53BP1

using in vitro DNA curtain assay, it has been shown that phase-separated droplets form in three steps (Keenen et al. 2021). Before DNA condensation, HP1α would bind to distal points of DNA strands simultaneously. The bound HP1α would then capture lateral DNA fluctuations to initiate DNA compaction. The process would proceed by trapping more noncompacted DNA through HP1a-DNA and HP1a– HP1a interactions. Further experiments indicated that the stability of compaction is more susceptible to fluctuations in HP1 concentration than in DNA level (Keenen et  al. 2021). However, how HP1 compact DNA in  vivo is not completely understood. Binding with other proteins could affect the critical concentration of HP1 for condensate formation (Keenen et  al. 2021). One such protein studied in detail is tumor suppressor p53-binding protein 1 (53BP1).

53BP1 53BP1 is a large 1972 amino acid protein encoded by the gene TP53BP1, which is located on chromosome 15 in humans. The protein was first identified in 1994 in a yeast two-hybrid screen as an interactor of p53, hence the name (Iwabuchi et  al. 1994). As a multi-domain protein, 53BP1 acts as an adaptor protein in DNA damage response and plays an important role in determining which double-strand break (DSB) repair pathway should be activated (Zimmermann and de Lange 2014). This is due to the ability of 53BP1 to bind to multiple DSB-responsive proteins. In doing so, 53BP1 inhibits breast cancer type 1 (BRCA1) susceptibility protein-mediated homolog-directed DNA repair (HDR) and promotes nonhomologous end-joining (NHEJ)mediated DSB repair (Mirza-Aghazadeh-Attari et al. 2019; Chapman et al. 2012). Loss of 53BP1 has been associated with genome instability and cell cycle arrest.

53BP1 and Proteins in DNA Damage Response DNA damage response (DDR) is carried out by different mechanisms involving multiple proteins. The main player of DDR is p53, which orchestrates multiple downstream pathways and determines the fate of the cell, depending on the severity of the damage (Williams and Schumacher 2016). p53 is rapidly recruited to DNA damage sites, where it organizes DDR and directs early repair pathway selection independent of its transcription function (Wang et al. 2022). The rapid recruitment of p53 depends on its modification by poly (ADP-ribose) polymerase (PARP), downstream from signaling cascades emerging from the MRE11-RAD50-NBS1 (MRN) complex, a DNA damage sensor (Bian et al. 2019). Depending on the severity of the damage, apoptosis, autophagy, cell cycle arrest, NHEJ, or HDR will take place. For instance, recruitment of the downstream effector 53BP1 by p53 early during DDR promotes NHEJ over the more error-prone alternative NHEJ response, microhomology-mediated end joining (MMEJ) (Wang et al. 2022; Seol et al. 2018).

Cell Cycle Phase-Dependent DNA Damage Repair

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BRCA1 is essential for DSB repair by HDR. Mutations in the BRCA1 increase an individual’s susceptibility to breast and ovarian cancer and are associated with defects in homologous recombination-mediated DSB repair, leading to instability and tumorigenesis (Mirza-Aghazadeh-Attari et al. 2019; Chapman et al. 2012). A study has shown that cells expressing a mutant BRCA1 protein lacking the N-terminal RING domain, associated with exon 2 deletional mutation, have increased genomic instability but can carry out BRCA1-dependent DNA repair (Li et al. 2016). However, removing 53BP1 rescues the defects due to loss of BRCA1 RING domain. Thus, mice with RING-less mutant BRCA1 and loss of 53BP1 do not have increased tumor susceptibility, suggesting that the function of BRCA1 in genomic integrity requires 53BP1 (Li et al. 2016). PARP is a group of proteins involved in apoptosis, DNA repair, and genomic stability. PARP1 is essential for base excision repair and DNA single-stranded breaks (SSBs) repair. PARP inhibition leads to unrepaired SSBs, which can be converted to become DSBs, which are normally repaired by the HDR pathway involving BRCA proteins. However, DNA damage is lethal to cells with inadequate homologous recombination, which is most common in mammary tumors with loss of BRCA1 or BRCA2. Thus, PARP inhibition causes synthetic lethality to cancer cells with BRCA1 or BRCA2 mutations. A study has shown, however, that loss of 53BP1 can increase homologous recombination in mutant cells, therefore reducing the effectiveness of PARP inhibition in cancer therapy (Jaspers et al. 2013).

Cell Cycle Phase-Dependent DNA Damage Repair Several DNA damage response (DDR) proteins, including 53BP1 and BRCA1, form ionizing-radiation-induced foci (IRIF) upon irradiation. By observing the distribution of 53BP1 and BRCA1 proteins within IRIF using super-resolution microscopy, a study has found that during the G0 and G1 phases of the cell cycle, 53BP1 is enriched in IRIF, consistent with its role in repairing DSBs by NHEJ; during the S phase, 53BP1 was pushed to the periphery while BRCA1 appeared at the core of IRIF, consistent with DSB repair by HDR (Chapman et al. 2012). The researchers propose that there is a temporal switch in DNA repair caused by BRCA1 inhibiting 53BP1-dependent DNA repair, and that a failure to do so, as in BRCA1-deficient cancer cells, results in genome instability (Chapman et al. 2012). Indeed, repairing DSBs can occur in two different pathways depending on the cell cycle phase (Zimmermann and de Lange 2014). During the S phase, DSBs are repaired by HDR, and during the G1 phase, they are repaired by classical NHEJ. DSBs are first detected by the MRE11-RAD50-NBS1 (MRN) DNA damage sensor, which activates ATM or ATRs, which signal 53BP1 to form foci near DNA lesions using its tandem Tudor domains and dysfunctional telomeres forming telomere-­dysfunction-induced foci (TIFs). These foci do not immediately induce classical NHEJ. Instead, they cooperate with RIF1 to prevent any resection of DSBs during G1, which favored classical NHEJ repair over homologous recombination.

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4  HP1 in Liquid–Liquid Phase Separation and its Regulation by 53BP1

53BP1 along with RIF1 blocks 5′ end resection, preventing HDR initiation. It has been shown that silencing 53BP1 in cells with damaged chromatin results in DSB repair performed by RAD51, which converts a gene to mutagenic single-strand annealing by RAD52 (Ochs et  al. 2016). RAD52-mediated HDR is believed to increase genome instability. Thus, 53BP1 coordinates different proteins and fosters fidelity of HDR (Ochs et al. 2016). It is thus speculated that 53BP1 does not determine DSB repair pathway choice, but instead acts as a DSB escort preventing potentially tumorigenic recombination (Mirman and de Lange 2020).

Other Pathways in DNA Damage Response 53BP1 also plays an important role in regulating the nuclear factor kappa-light-­ chain-enhancer of activated B cells (NF-κB) and serine/threonine kinase AKT signaling pathways during DDR. NF-κB signaling promotes tumorigenesis by inducing cellular proliferation, angiogenesis, epithelial–mesenchymal transition, metastasis, and alterations in the tumor microenvironment. NF-κB is partly activated by DNA damage, which promotes survival of cancer cells, leading to resistance to chemotherapy. 53BP1 functions as tumor suppressor partly by inhibiting the NF-κB pathway, decreasing metastasis and tumor progression (Li et  al. 2012; Mirza-Aghazadeh-Attari et al. 2019). On the other hand, studies have shown that 53BP1 can regulate AKT signaling to suppress tumor growth and increase cancer cell apoptosis (Hong et al. 2012). AKT signaling is another pathway that generally promotes tumorigenesis. Upon activation, AKT mediates phosphorylation of the X-ray repair cross-complementing 4 (XRCC4)-like factors (XLF), causing its dissociation from and disruption of the XRCC4/DNA ligase IV complex, leading to impaired function of 53BP1 in NHEJ (Liu et al. 2015). The other major pathway in DDR is HDR, which is important as programmable nucleases, such as CRISPR-Cas9, can be passed through HDR for precise genome editing. However, HDR has constraints, such as competition from the NHEJ pathway. 53BP1 plays a role in determining pathway choice in DDR and favors NHEJ over HDR (Ochs et  al. 2016). Indeed, inhibiting 53BP1 can increase HDR efficiency. Using ubiquitin variants to target the 53BP1 ubiquitin-dependent recruitment (UDR) domain involved in ubiquitylated histone recognition, it has been shown that one such variant, i53, blocked 53BP1 accumulation at DSBs, increasing the efficiency of HDR-based precise genome editing (Canny et al. 2018).

53BP1 in Heterochromatin Regulation Recently, a previously unknown or noncanonical function of 53BP1 in protecting genome integrity has been identified, which is regulating HP1 and heterochromatin through a process called LLPS (Zhang et al. 2022). HP1 by itself can undergo LLPS

53BP1 in Heterochromatin Regulation

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(Strom et al. 2017; Larson et al. 2017). In experiments in vitro, the critical concentration required by HP1 to form condensates is much higher than physiological concentration of HP1 in vivo (Keenen et al. 2021), raising the possibility that other proteins may cooperate with HP1, lowering its critical concentration for condensate formation. Aside from its role in DNA DSB repair, 53BP1 can associate with HP1 to regulate LLPS and heterochromatin formation (Zhang et  al. 2022), which is believed to play a role in protecting cells from DNA damage to ensure genome stability. The eukaryotic cell consists of numerous compartments or organelles, such as the nucleus, ribosomes, and mitochondria, to name a few, that are delimited by membranes. Cells also contain membraneless compartments, such as nucleoli, Cajal bodies, promyelocytic leukemia protein (PML) bodies, and processing bodies (P bodies), that are formed by phase separation under normal or stressed conditions, such as in cancer cells (Jiang et al. 2020). LLPS is the formation of membraneless liquid droplets by proteins or macromolecules. When their concentration reaches a threshold, they can separate from the surrounding solution, achieving phase separation. Previous studies have shown that core heterochromatin proteins like HP1 undergo LLPS (Strom et al. 2017; Larson et al. 2017). A recent study investigated how LLPS is involved in heterochromatin formation (Zhang et al. 2022). The researchers first investigated 53BP1 puncta formation in the nucleus. These 53BP1 puncta were distinct from DSB foci but were localized at heterochromatin centers and colocalized with HP1α. Deletion of 53BP1 led to a reduction in heterochromatin centers and derepression of transcription of heterochromatic sequences, suggesting a role of 53BP1 in heterochromatin regulation (Zhang et al. 2022). Then, the researchers investigated the biological function of 53BP1 puncta in the nucleus. It was found that 53BP1 puncta are characteristic of LLPS, and that 53BP1 undergoes LLPS with HP1α in a mutually dependent manner. The researchers further studied LLPS by 53BP1 and HP1α in vitro with purified proteins. They found that HP1α by itself did not form liquid droplets in the absence of crowding agents, and that 53BP1 by itself formed very few droplets. However, when combined together, 53BP1 and HP1α readily undergo LLPS and a large amount of protein would precipitate upon centrifugation. They further narrowed down the domain of 53BP1 important for LLPS and identified the OD-Tudor-UDR-NLS region as being required for puncta formation and LLPS. They further found that the C-terminal BRCTs and the N-terminus, which are vital to DSB repair, did not affect puncta formation, suggesting that 53BP1 puncta formation is a function completely distinct from DSB repair (Zhang et al. 2022). Based on in  vitro LLPS assays and in  vivo microscopic observations, the researchers concluded that 53BP1 and HP1ɑ undergo LLPS in a mutually dependent manner, and that they together are required for maintaining heterochromatin stability. Thus, in addition to regulating DSB repair, 53BP1 is important for maintaining heterochromatin and genome stability through LLPS (Zhang et al. 2022). It remains to be investigated the roles of other proteins in modulating LLPS and other properties of HP1, the central component of heterochromatin.

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Index

A Aging, 13, 15–17 ATF-2, 21, 23, 24

N Non-canonical STAT, 3, 6 Nuclear factor-kappa B (NF-κB), 24

C Cancer, 3, 16, 21, 25, 39–41 C/EBPα, 27, 28

P Paired box (PAX), 25–27

E Epigenetics, 25, 28 G GAGA, 19–20 H Heterochromatin, 2–6, 13–17, 20, 22, 24, 25, 27, 28, 37, 40, 41 Heterochromatin Protein 1 (HP1), 4, 28 L Liquid-liquid phase separation (LLPS), 37, 40, 41

R Retinoblastoma protein (Rb), 5, 22 S Senescence, 15, 16 Signal transducer and activator of transcription (STAT), 1–6, 13–17, 24 T Transcription factors, 2, 5, 6, 19–21, 23–27 Tumor suppressor p53-binding protein 1 (53BP1), 38–41 U Unphosphorylated STAT (uSTAT), 2, 3, 6, 16, 17

© Springer Nature Switzerland AG 2023 W. X. Li, L. Silver-Morse, The Secret Lives of Transcription Factors, SpringerBriefs in Biochemistry and Molecular Biology, https://doi.org/10.1007/978-3-031-29029-9

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