DEPOD - human DEPhOsphorylation Database

Basic Information
Phosphatases
Pathway
Interacting Proteins
Phosphorylating Kinases

Gene Name	HIST1H2BM (QuickGO)	Interactive visualization of HIST1H2BM structures (A quick tutorial to explore the interctive visulaization) Representative structure: NA
Synonyms	HIST1H2BM, H2BFE
Protein Name	HIST1H2BM
Alternative Name(s)	Histone H2B type 1-M;Histone H2B.e;H2B/e;
Protein Family	Belongs to the histone H2B family
EntrezGene ID	8342 (Comparitive Toxicogenomics)
UniProt AC (Human)	Q99879 (protein sequence)
Enzyme Class	N/A
Molecular Weight	13989 Dalton
Protein Length	126 amino acids (AA)
Genome Browsers	NCBI \| ENSG00000273703 (Ensembl) \| UCSC
Crosslinking annotations	Query our ID-mapping table
Orthologues	Quest For Orthologues (QFO) \| GeneTree \| eggNOG - KOG1744 \| eggNOG - ENOG4111NV5
Phosphorylation Network	Visualize

Domain organization, Expression, Diseases

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Localization, Function, Catalytic activity and Sequence

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Motif information from Eukaryotic Linear Motif atlas (ELM)

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Gene Ontology (P: Process; F: Function and C: Component terms)

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KEGG Pathway
Reactome Pathway

Pathway ID	Pathway Name	Pathway Description (KEGG)
hsa05034	Alcoholism	Alcoholism, also called dependence on alcohol (ethanol), is a chronic relapsing disorder that is progressive and has serious detrimental health outcomes. As one of the primary mediators of the rewarding effects of alcohol, dopaminergic ventral tegmental area (VTA) projections to the nucleus accumbens (NAc) have been identified. Acute exposure to alcohol stimulates dopamine release into the NAc, which activates D1 receptors, stimulating PKA signaling and subsequent CREB-mediated gene expression, whereas chronic alcohol exposure leads to an adaptive downregulation of this pathway, in particular of CREB function. The decreased CREB function in the NAc may promote the intake of drugs of abuse to achieve an increase in reward and thus may be involved in the regulation of positive affective states of addiction. PKA signaling also affects NMDA receptor activity and may play an important role in neuroadaptation in response to chronic alcohol exposure.
hsa05203	Viral carcinogenesis	There is a strong association between viruses and the development of human malignancies. We now know that at least six human viruses, Epstein-Barr virus (EBV), hepatitis B virus (HBV), hepatitis C virus (HCV), human papilloma virus (HPV), human T-cell lymphotropic virus (HTLV-1) and Kaposi's associated sarcoma virus (KSHV) contribute to 10-15% of the cancers worldwide. Via expression of many potent oncoproteins, these tumor viruses promote an aberrant cell-proliferation via modulating cellular cell-signaling pathways and escape from cellular defense system such as blocking apoptosis. Human tumor virus oncoproteins can also disrupt pathways that are necessary for the maintenance of the integrity of host cellular genome. Viruses that encode such activities can contribute to initiation as well as progression of human cancers.
hsa05322	Systemic lupus erythematosus	Systemic lupus erythematosus (SLE) is a prototypic autoimmune disease characterised by the production of IgG autoantibodies that are specific for self-antigens, such as DNA, nuclear proteins and certain cytoplasmic components, in association with a diverse array of clinical manifestations. The primary pathological findings in patients with SLE are those of inflammation, vasculitis, immune complex deposition, and vasculopathy. Immune complexes comprising autoantibody and self-antigen is deposited particulary in the renal glomeruli and mediate a systemic inflammatory response by activating complement or via Fc{gamma}R-mediated neutrophil and macrophage activation. Activation of complement (C5) leads to injury both through formation of the membrane attack complex (C5b-9) or by generation of the anaphylatoxin and cell activator C5a. Neutrophils and macrophages cause tissue injury by the release of oxidants and proteases.

Pathway ID	Pathway Name	Pathway Description (KEGG)
R-HSA-1221632	Meiotic synapsis.	Meiotic synapsis is the stable physical pairing of homologous chromosomes that begins in leptonema of prophase I and lasts until anaphase of prophase I. First, short segments of axial elements form along chromosomes. Telomeres then cluster at a region of the inner nuclear membrane and axial elements extend and fuse along the length of the chromosomes. Subsequent to the initiation of recombination transverse filaments of SYCP1 link axial/lateral elements to a central element containing SYCE1 and SYCE2, thus forming the synaptonemal complex (reviewed in Yang and Wang 2009).Unsynapsed regions are silenced during pachynema by recruitment of BRCA1 and ATR, which phosphorylates histone H2AX (reviewed in Inagaki et al. 2010)
R-HSA-171306	Packaging Of Telomere Ends.	Multiple steps, including C-strand resection, telomerase-mediated elongation, and C-strand synthesis are involved in processing and maintaining the telomere. Though this module posits a linear transit for the steps, in humans it is not well understood how these steps are coordinated and what other events may be involved.Telomeric DNA can form higher order structures. Electron microscopy of telomeric DNA isolated from human cells provided evidence for lariat-type structures termed telomeric loops, or t-loops (Griffith et al., 1999). t-loops are proposed to result from the invasion of the 3' G-rich single strand overhang into the double stranded telomeric TTAGGG repeat tract. The function of the t-loop is presumed to be the masking of the 3' telomeric overhang. Multiple protein factors can bind telomeric DNA and likely contribute to dynamic, higher order structures
R-HSA-1912408	Pre-NOTCH Transcription and Translation.	In humans, the NOTCH protein family has four members: NOTCH1, NOTCH2, NOTCH3 and NOTCH4. NOTCH1 protein was identified first, as the product of a chromosome 9 gene translocated in T-cell acute lymphoblastic leukemia that was homologous to Drosophila Notch (Ellisen et al. 1991). At the same time, rat Notch1 was cloned (Weinmaster et al. 1991), followed by cloning of mouse Notch1, named Motch (Del Amo et al. 1992). NOTCH2 protein is the product of a gene on chromosome 1 (Larsson et al. 1994). NOTCH2 expression is differentially regulated during B-cell development (Bertrand et al. 2000). NOTCH2 mutations are a rare cause of Alagille syndrome (McDaniell et al. 2006). NOTCH3 is the product of a gene on chromosome 19. NOTCH3 mutations are the underlying cause of CADASIL, cerebral arteriopathy with subcortical infarcts and leukoencephalopathy (Joutel et al. 1996). NOTCH4, the last NOTCH protein discovered, is the product of a gene on chromosome 6 (Li et al. 1998). MicroRNAs play an important negative role in translation and/or stability of NOTCH mRNAs. MicroRNAs miR-34 (miR-34A, miR-34B and mi-R34C), whose transcription is directly induced by the tumor suppressor protein p53 (Chang et al. 2007, Raver-Shapira et al. 2007, He et al. 2007, Corney et al. 2007) bind and negatively regulate translation of NOTCH1 mRNA (Li et al. 2009, Pang et al. 2010, Ji et al. 2009) and NOTCH2 mRNA (Li et al. 2009). NOTCH1 mRNA translation is also negatively regulated by microRNAs miR-200B and miR-200C (Kong et al. 2010), as well as miR-449A, miR-449B and miR-449C (Marcet et al. 2011). Translation of NOTCH3 mRNA is negatively regulated by microRNAs miR-150 (Ghisi et al. 2011) and miR-206 (Song et al. 2009). Translation of NOTCH4 mRNA is negatively regulated by microRNAs miR-181C (Hashimoto et al. 2010) and miR-302A (Costa et al. 2009). Nascent NOTCH peptides are co-translationally targeted to the endoplasmic reticulum for further processing, followed by modification in the Golgi apparatus, before trafficking to the plasma membrane. Endoplasmic reticulum calcium ATPases, positively regulate NOTCH trafficking, possibly by contributing to accurate folding of NOTCH precursors (Periz et al. 1999)
R-HSA-201722	Formation of the beta-catenin:TCF transactivating complex.	Once in the nucleus, beta-catenin is recruited to WNT target genes through interaction with TCF/LEF transcription factors. This family, which consists of TCF7 (also known as TCF1), TCF7L1 (also known as TCF3), TCF7L2 (also known as TCF4) and TCF7L3 (also known as LEF1), are HMG-containing transcription factors that bind to the WNT responsive elements in target gene promoters (reviewed in Brantjes et al, 2002). In the absence of WNT signal, TCF/LEF proteins recruit Groucho/TLE repressors to inhibit transcription; upon WNT stimulation, beta-catenin can displace Groucho/TLE from TCF/LEF proteins to initiate transcriptional activation (reviewed in Chen and Courey, 2000). Although this model for WNT-dependent activation of target genes is widely accepted, it is important to note that TCF/LEF proteins are not redundant and can contribute to WNT target gene expression in a number of different ways (reviewed in Brantjes et al, 2002; MacDonald et al, 2009). In particular, TCF7L1 (TCF3) is thought to have a more pronounced repressor function than other TCF/LEF family members. A couple of recent studies in Xenopus and mammalian cells show that WNT- and beta-catenin-dependent phosphorylation of TCF7L1(TCF3) promotes its dissociation from the promoter of target genes and allows gene expression through relief of this repression activity (Hikasa et al, 2010; Hikasa et al, 2011).The role of beta-catenin at WNT promoters hinges upon its ability to act as a scaffold for the recruitment of other proteins. The structure of beta-catenin consists of 12 imperfect ARM repeats (R1-12) flanked by an N-terminal and C-terminal extension (NTD and CTD respectively), with a conserved Helix C located between R12 and the CTD. Nuclear beta-catenin interacts with TCF/LEF at WNT target genes through ARM domains 3-9 (Graham et al, 2000; Poy et al, 2001; Xing et al, 2008). The N and the C terminal regions are important for the recruitment of transcriptional activator and repressors that contribute to WNT target gene expression (reviewed in Mosimann et al, 2009; Valenta et al, 2012). The N-terminal ARM domains 1-4 recruit the WNT-pathway specific activators BCL9:PYGO while the C-terminal region (R11-CTD) interacts with a wide range of general transcriptional activators that are involved in chromatin remodelling and transcription initiation. These include HATs such as P300, CBP and TIP60, histone methyltransferases such as MLL1 and 2, SWI/SNF factors BRG1 and ISWI and components of the PAF complex (reviewed in Mosimann et al, 2009; Valenta et al, 2012). Although many binding partners have been identified for the C-terminal region of beta-catenin, in many cases the timing and relationship of these interactions and indeed, the exact complex composition remains to be elucidated. Moreover, because many of the interacting partners appear to bind to overlapping regions of beta-catenin, it is unlikely that they all bind simultaneously. For simplicity, the interactions have been depicted as though they occur independently of one another; more accurately they are likely to cycle successively on and off beta-catenin to promote an active chromatin structure (reviewed in Willert and Jones, 2006; Valenta et al, 2012)
R-HSA-212300	PRC2 methylates histones and DNA.	Polycomb group proteins are responsible for the heritable repression of genes during development (Lee et al. 2006, Ku et al. 2008, reviewed in Simon and Kingston 2009, Margueron and Reinberg 2011, Di Croce and Helin 2013). Two major families of Polycomb complexes exist: Polycomb Repressive Complex 1 (PRC1) and Polycomb Repressive Complex 2 (PRC2). PRC1 and PRC2 each appear to comprise sets of distinct complexes that contain common core subunits and distinct accessory subunits (reviewed in Nayak et al. 2011). PRC2, through its component EZH2 or, in some complexes, EZH1 produces the initial molecular mark of repression, the trimethylation of lysine-27 of histone H3 (H3K27me3). How PRC2 is initially recruited to a locus remains unknown, however cytosine-guanine (CpG) motifs and transcripts have been suggested. Different mechanisms may be used at different loci. The trimethylated H3K27 produced by PRC2 is bound by the Polycomb subunit of PRC1. PRC1 ubiquitinates histone H2A and maintains repression
R-HSA-2299718	Condensation of Prophase Chromosomes.	In mitotic prophase, the action of the condensin II complex enables initial chromosome condensation.The condensin II complex subunit NCAPD3 binds monomethylated histone H4 (H4K20me1), thereby associating with chromatin (Liu et al. 2010). Binding of the condensin II complex to chromatin is partially controlled by the presence of RB1 (Longworth et al. 2008). Two mechanisms contribute to the accumulation of H4K20me1 at mitotic entry. First, the activity of SETD8 histone methyltransferase peaks at G2/M transition (Nishioka et al. 2002, Rice et al. 2002, Wu et al. 2010). Second, the complex of CDK1 and cyclin B1 (CDK1:CCNB1) phosphorylates PHF8 histone demethylase at the start of mitosis, removing it from chromatin (Liu et al. 2010).Condensin II complex needs to be phosphorylated by the CDK1:CCNB1 complex, and then phosphorylated by PLK1, in order to efficiently condense prophase chromosomes (Abe et al. 2011)
R-HSA-2559580	Oxidative Stress Induced Senescence.	Oxidative stress, caused by increased concentration of reactive oxygen species (ROS) in the cell, can happen as a consequence of mitochondrial dysfunction induced by the oncogenic RAS (Moiseeva et al. 2009) or independent of oncogenic signaling. Prolonged exposure to interferon-beta (IFNB, IFN-beta) also results in ROS increase (Moiseeva et al. 2006). ROS oxidize thioredoxin (TXN), which causes TXN to dissociate from the N-terminus of MAP3K5 (ASK1), enabling MAP3K5 to become catalytically active (Saitoh et al. 1998). ROS also stimulate expression of Ste20 family kinases MINK1 (MINK) and TNIK through an unknown mechanism, and MINK1 and TNIK positively regulate MAP3K5 activation (Nicke et al. 2005). MAP3K5 phosphorylates and activates MAP2K3 (MKK3) and MAP2K6 (MKK6) (Ichijo et al. 1997, Takekawa et al. 2005), which act as p38 MAPK kinases, as well as MAP2K4 (SEK1) (Ichijo et al. 1997, Matsuura et al. 2002), which, together with MAP2K7 (MKK7), acts as a JNK kinase. MKK3 and MKK6 phosphorylate and activate p38 MAPK alpha (MAPK14) and beta (MAPK11) (Raingeaud et al. 1996), enabling p38 MAPKs to phosphorylate and activate MAPKAPK2 (MK2) and MAPKAPK3 (MK3) (Ben-Levy et al. 1995, Clifton et al. 1996, McLaughlin et al. 1996, Sithanandam et al. 1996, Meng et al. 2002, Lukas et al. 2004, White et al. 2007), as well as MAPKAPK5 (PRAK) (New et al. 1998 and 2003, Sun et al. 2007). Phosphorylation of JNKs (MAPK8, MAPK9 and MAPK10) by MAP3K5-activated MAP2K4 (Deacon and Blank 1997, Fleming et al. 2000) allows JNKs to migrate to the nucleus (Mizukami et al. 1997) where they phosphorylate JUN. Phosphorylated JUN binds FOS phosphorylated by ERK1 or ERK2, downstream of activated RAS (Okazaki and Sagata 1995, Murphy et al. 2002), forming the activated protein 1 (AP-1) complex (FOS:JUN heterodimer) (Glover and Harrison 1995, Ainbinder et al. 1997). Activation of p38 MAPKs and JNKs downstream of MAP3K5 (ASK1) ultimately converges on transcriptional regulation of CDKN2A locus. In dividing cells, nucleosomes bound to the CDKN2A locus are trimethylated on lysine residue 28 of histone H3 (HIST1H3A) by the Polycomb repressor complex 2 (PRC2), creating the H3K27Me3 (Me3K-28-HIST1H3A) mark (Bracken et al. 2007, Kotake et al. 2007). The expression of Polycomb constituents of PRC2 (Kuzmichev et al. 2002) - EZH2, EED and SUZ12 - and thereby formation of the PRC2, is positively regulated in growing cells by E2F1, E2F2 and E2F3 (Weinmann et al. 2001, Bracken et al. 2003). H3K27Me3 mark serves as a docking site for the Polycomb repressor complex 1 (PRC1) that contains BMI1 (PCGF4) and is therefore named PRC1.4, leading to the repression of transcription of p16-INK4A and p14-ARF from the CDKN2A locus, where PCR1.4 mediated repression of p14-ARF transcription in humans may be context dependent (Voncken et al. 2005, Dietrich et al. 2007, Agherbi et al. 2009, Gao et al. 2012). MAPKAPK2 and MAPKAPK3, activated downstream of the MAP3K5-p38 MAPK cascade, phosphorylate BMI1 of the PRC1.4 complex, leading to dissociation of PRC1.4 complex from the CDKN2A locus and upregulation of p14-ARF transcription (Voncken et al. 2005). AP-1 transcription factor, formed as a result of MAP3K5-JNK signaling, as well as RAS signaling, binds the promoter of KDM6B (JMJD3) gene and stimulates KDM6B expression. KDM6B is a histone demethylase that removes H3K27Me3 mark i.e. demethylates lysine K28 of HIST1H3A, thereby preventing PRC1.4 binding to the CDKN2A locus and allowing transcription of p16-INK4A (Agger et al. 2009, Barradas et al. 2009, Lin et al. 2012). p16-INK4A inhibits phosphorylation-mediated inactivation of RB family members by CDK4 and CDK6, leading to cell cycle arrest (Serrano et al. 1993). p14-ARF inhibits MDM2-mediated degradation of TP53 (p53) (Zhang et al. 1998), which also contributes to cell cycle arrest in cells undergoing oxidative stress. In addition, phosphorylation of TP53 by MAPKAPK5 (PRAK) activated downstream of MAP3K5-p38 MAPK signaling, activates TP53 and contributes to cellular senescence (Sun et al. 2007)
R-HSA-2559582	Senescence-Associated Secretory Phenotype (SASP).	The culture medium of senescent cells in enriched in secreted proteins when compared with the culture medium of quiescent i.e. presenescent cells and these secreted proteins constitute the so-called senescence-associated secretory phenotype (SASP), also known as the senescence messaging secretome (SMS). SASP components include inflammatory and immune-modulatory cytokines (e.g. IL6 and IL8), growth factors (e.g. IGFBPs), shed cell surface molecules (e.g. TNF receptors) and survival factors. While the SASP exhibits a wide ranging profile, it is not significantly affected by the type of senescence trigger (oncogenic signalling, oxidative stress or DNA damage) or the cell type (epithelial vs. mesenchymal) (Coppe et al. 2008). However, as both oxidative stress and oncogenic signaling induce DNA damage, the persistent DNA damage may be a deciding SASP initiator (Rodier et al. 2009). SASP components function in an autocrine manner, reinforcing the senescent phenotype (Kuilman et al. 2008, Acosta et al. 2008), and in the paracrine manner, where they may promote epithelial-to-mesenchymal transition (EMT) and malignancy in the nearby premalignant or malignant cells (Coppe et al. 2008). Interleukin-1-alpha (IL1A), a minor SASP component whose transcription is stimulated by the AP-1 (FOS:JUN) complex (Bailly et al. 1996), can cause paracrine senescence through IL1 and inflammasome signaling (Acosta et al. 2013). Here, transcriptional regulatory processes that mediate the SASP are annotated. DNA damage triggers ATM-mediated activation of TP53, resulting in the increased level of CDKN1A (p21). CDKN1A-mediated inhibition of CDK2 prevents phosphorylation and inactivation of the Cdh1:APC/C complex, allowing it to ubiquitinate and target for degradation EHMT1 and EHMT2 histone methyltransferases. As EHMT1 and EHMT2 methylate and silence the promoters of IL6 and IL8 genes, degradation of these methyltransferases relieves the inhibition of IL6 and IL8 transcription (Takahashi et al. 2012). In addition, oncogenic RAS signaling activates the CEBPB (C/EBP-beta) transcription factor (Nakajima et al. 1993, Lee et al. 2010), which binds promoters of IL6 and IL8 genes and stimulates their transcription (Kuilman et al. 2008, Lee et al. 2010). CEBPB also stimulates the transcription of CDKN2B (p15-INK4B), reinforcing the cell cycle arrest (Kuilman et al. 2008). CEBPB transcription factor has three isoforms, due to three alternative translation start sites. The CEBPB-1 isoform (C/EBP-beta-1) seems to be exclusively involved in growth arrest and senescence, while the CEBPB-2 (C/EBP-beta-2) isoform may promote cellular proliferation (Atwood and Sealy 2010 and 2011). IL6 signaling stimulates the transcription of CEBPB (Niehof et al. 2001), creating a positive feedback loop (Kuilman et al. 2009, Lee et al. 2010). NF-kappa-B transcription factor is also activated in senescence (Chien et al. 2011) through IL1 signaling (Jimi et al. 1996, Hartupee et al. 2008, Orjalo et al. 2009). NF-kappa-B binds IL6 and IL8 promoters and cooperates with CEBPB transcription factor in the induction of IL6 and IL8 transcription (Matsusaka et al. 1993, Acosta et al. 2008). Besides IL6 and IL8, their receptors are also upregulated in senescence (Kuilman et al. 2008, Acosta et al. 2008) and IL6 and IL8 may be master regulators of the SASP. IGFBP7 is also an SASP component that is upregulated in response to oncogenic RAS-RAF-MAPK signaling and oxidative stress, as its transcription is directly stimulated by the AP-1 (JUN:FOS) transcription factor. IGFBP7 negatively regulates RAS-RAF (BRAF)-MAPK signaling and is important for the establishment of senescence in melanocytes (Wajapeyee et al. 2008). Please refer to Young and Narita 2009 for a recent review
R-HSA-2559586	DNA Damage/Telomere Stress Induced Senescence.	Reactive oxygen species (ROS), whose concentration increases in senescent cells due to oncogenic RAS-induced mitochondrial dysfunction (Moiseeva et al. 2009) or due to environmental stress, cause DNA damage in the form of double strand breaks (DSBs) (Yu and Anderson 1997). In addition, persistent cell division fueled by oncogenic signaling leads to replicative exhaustion, manifested in critically short telomeres (Harley et al. 1990, Hastie et al. 1990). Shortened telomeres are no longer able to bind the protective shelterin complex (Smogorzewska et al. 2000, de Lange 2005) and are recognized as damaged DNA. The evolutionarily conserved MRN complex, consisting of MRE11A (MRE11), RAD50 and NBN (NBS1) subunits, binds DSBs (Lee and Paull 2005) and shortened telomeres that are no longer protected by shelterin (Wu et al. 2007). Once bound to the DNA, the MRN complex recruits and activates ATM kinase (Lee and Paull 2005, Wu et al. 2007), leading to phosphorylation of ATM targets, including TP53 (p53) (Banin et al. 1998, Canman et al. 1998, Khanna et al. 1998). TP53, phosphorylated on serine S15 by ATM, binds the CDKN1A (also known as p21, CIP1 or WAF1) promoter and induces CDKN1A transcription (El-Deiry et al. 1993, Karlseder et al. 1999). CDKN1A inhibits the activity of CDK2, leading to G1/S cell cycle arrest (Harper et al. 1993, El-Deiry et al. 1993). SMURF2 is upregulated in response to telomere attrition in human fibroblasts and induces senecscent phenotype through RB1 and TP53, independently of its role in TGF-beta-1 signaling (Zhang and Cohen 2004). The exact mechanism of SMURF2 involvement is senescence has not been elucidated
R-HSA-3214815	HDACs deacetylate histones.	Lysine deacetylases (KDACs), historically referred to as histone deacetylases (HDACs), are divided into the Rpd3/Hda1 metal-dependent 'classical HDAC family' (de Ruijter et al. 2003, Verdin et al. 2003) and the unrelated sirtuins (Milne & Denu 2008). Phylogenetic analysis divides human KDACs into four classes (Gregoretti et al. 2004): Class I includes HDAC1, 2, 3 and 8; Class IIa includes HDAC4, 5, 7 and 9; Class IIb includes HDAC6 and 10; Class III are the sirtuins (SIRT1-7); Class IV has one member, HDAC11 (Gao et al. 2002). Class III enzymes use an NAD+ cofactor to perform deacetylation (Milne & Denu 2008, Yang & Seto 2008), the others classes use a metal-dependent mechanism (Gregoretti et al. 2004) to catalyze the hydrolysis of acetyl-L-lysine side chains in histone and non-histone proteins yielding L-lysine and acetate. X-ray crystal structures are available for four human HDACs; these structures have conserved active site residues, suggesting a common catalytic mechanism (Lombardi et al. 2011). They require a single transition metal ion and are typically studied in vitro as Zn2+-containing enzymes, though in vivo HDAC8 exhibits increased activity when substituted with Fe2+ (Gantt et al. 2006). The structurally-related enzyme acetylpolyamine amidohydrolase (APAH) (Leipe & Landsman 1997) exhibits optimal activity with Mn2+, followed closely by Zn2+ (Sakurada et al. 1996).HDACs are often part of multi-protein transcriptional complexes that are recruited to gene promoters, regulating transcription without direct DNA binding. With the exception of HDAC8, all class I members can be catalytic subunits of multiprotein complexes (Yang & Seto 2008). HDAC1 and HDAC2 interact to form the catalytic core of several multisubunit complexes including Sin3, nucleosome remodeling deacetylase (NuRD) and corepressor of REST (CoREST) complexes (Grozinger & Schreiber 2002). HDAC3 is part of the silencing mediator of retinoic acid and thyroid hormone receptor (SMRT) complex or the homologous nuclear receptor corepressor (NCoR) (Li et al. 2000, Wen et al. 2000, Zhang et al. 2002, Yoon et al. 2003, Oberoi et al. 2011) which are involved in a wide range of processes including metabolism, inflammation, and circadian rhythms (Mottis et al. 2013). Class IIa HDACs (HDAC4, -5, -7, and -9) shuttle between the nucleus and cytoplasm (Yang & Seto 2008, Haberland et al. 2009). The nuclear export of class IIa HDACs requires phosphorylation stimulated by calcium or other stimuli. They appear to have been evolutionarily inactivated as enzymes, having acquired a histidine substitution of the tyrosine residue in the active site of the mammalian deacetylase domain (H976 in humans) (Lahm et al. 2007, Schuetz et al. 2008). Instead they function as transcriptional corepressors for the MEF2 family of transcription factors (Yang & Gregoire 2005) .Histones are the primary substrate for most HDACs except HDAC6 which is predominantly cytoplasmic and acts on alpha-tublin (Hubbert et al. 2002, Zhang et al. 2003, Boyault et al. 2007). HDACs also deacetylate proteins such as p53, E2F1, RelA, YY1, TFIIE, BCL6 and TFIIF (Glozak et al. 2005).Histone deacetylases are targeted by structurally diverse compounds known as HDAC inhibitors (HDIs) (Marks et al. 2000). These can induce cytodifferentiation, cell cycle arrest and apoptosis of transformed cells (Marks et al. 2000, Bolden et al. 2006). Some HDIs have significant antitumor activity (Marks and Breslow 2007, Ma et al. 2009) and at least two are approved anti-cancer drugs.The coordinates of post-translational modifications represented and described here follow UniProt standard practice whereby coordinates refer to the translated protein before any further processing. Histone literature typically refers to coordinates of the protein after the initiating methionine has been removed. Therefore the coordinates of post-translated residues in the Reactome database and described here are frequently +1 when compared with the literature
R-HSA-3214847	HATs acetylate histones.	Histone acetyltransferases (HATs) involved in histone modifications are referred to as A-type or nuclear HATs. They can be grouped into at least four families based on sequence conservation within the HAT domain: Gcn5/PCAF, MYST, p300/CBP and Rtt109. The p300/CBP and Rtt109 families are specific to metazoans and fungi respectively (Marmorstein & Trievel 2009). Gcn5/PCAF and MYST family members have no significant sequence homology but share a globular alpha/beta fold with a common structure involved in acetyl-Coenzyme A (ACA) binding. Both use a conserved glutamate residue for the acetyl transfer reaction but may not share a common catalytic mechanism (Trievel et al. 1999, Tanner et al. 1999, Yan et al. 2002, Berndsen et al. 2007). The p300/CBP HAT domain has no homology with the other families but some structural conservation within theACA-binding core (Liu et al. 2008). In addition to histone acetylation, members of all 3 human HAT families have been shown to acetylate non-histones (Glozak et al. 2005). HATs and histone deacetylase (HDAC) enzymes generally act not alone but as part of multiprotein complexes. There are numerous examples in which subunits of HAT or HDAC complexes influence their substrate specificity and lysine preference, which in turn, affect the broader functions of these enzymes (Shahbazian & Grunstein 2007). N.B. The coordinates of post-translational modifications represented and described here follow UniProt standard practice whereby coordinates refer to the translated protein before any further processing. Histone literature typically refers to coordinates of the protein after the initiating methionine has been removed. Therefore the coordinates of post-translated residues in the Reactome database and described here are frequently +1 when compared with the literature
R-HSA-427359	SIRT1 negatively regulates rRNA expression.	Expression of rRNA genes is coupled to the overall metabolism of the cell by the NAD-dependent histone deacetylase SIRT1, a component of the Energy-dependent Nucleolar Silencing Complex (eNoSC) (Murayama et al. 2008, reviewed in Salminen and Kaarniranta 2009, Grummt and Voit 2010). eNoSC comprises Nucleomethylin (NML), SIRT1, and the histone methylase SUV39H1 (Murayama et al. 2008). Deacetylation and methylation of histone H3 in the chromatin of a rRNA gene by eNoSC causes reduced expression of the gene. When glucose is low, NAD is high (NADH is low), activity of SIRT1 is high, and activity of rRNA genes is reduced. It is hypothesized that eNoSC forms on a nucleosome containing dimethylated lysine-9 on histone H3 (H3K9me2) and then eNoSC deacetylates and dimethylates the adjacent nucleosome, thus catalyzing spreading of H3K9me2 throughout the gene
R-HSA-427389	ERCC6 (CSB) and EHMT2 (G9a) positively regulate rRNA expression.	About half of the rRNA genes in the genome are actively expressed, being transcribed by RNA polymerase I (reviewed in Nemeth and Langst 2008, Bartova et al. 2010, Goodfellow and Zomerdijk 2012, Grummt and Langst 2013). As inferred from mouse, those genes that are expressed are activated by ERCC6 (also known as Cockayne Syndrome protein, CSB) which interacts with TTF-I bound to the T0 terminator region (also know as the Sal Box) of rRNA genes (Yuan et al. 2007, reviewed in Birch and Zomerdijk 2008, Grummt and Langst 2013). ERCC6 recruits the histone methyltransferase EHMT2 (also known as G9a) which dimethylates histone H3 at lysine-9 in the coding region of rRNA genes. The dimethylated lysine is bound by CBX3 (also known as Heterochromatic Protein-1gamma, HP1gamma) and increases expression of the rRNA gene. Continuing dimethylation depends on continuing transcription. Mutations in CSB result in dysregulation of RNA polymerase I transcription, which plays a role in the symptoms of Cockayne Syndrome (reviewed in Hannan et al. 2013)
R-HSA-427413	NoRC negatively regulates rRNA expression.	The Nucleolar Remodeling Complex (NoRC) comprising TIP5 (BAZ2A) and the chromatin remodeller SNF2H (SMARCA5) silences rRNA gene (reviewed in Santoro and Grummt 2001, Grummt 2007, Preuss and Pikaard 2007, Birch and Zommerdijk 2008, McStay and Grummt 2008, Grummt and Langst 2013). The TAM domain of TIP5 (BAZ2A) binds promoter-associated RNA (pRNA) transcribed from the intergenic spacer region of rDNA. The pRNA bound by TIP5 is required to direct the complex to the main promoter of the rRNA gene possibly by triple helix formation between pRNA and the rDNA. The PHD domain of TIP5 binds histone H4 acetylated at lysine-16. Transcription Termination Factor-I (TTF-I) binds to a promoter-proximal terminator (T0 site) in the rDNA and interacts with the TIP5 subunit of NoRC. NoRC also interacts with the SIN3-HDAC complex, HDAC1, HDAC2, DNMT1, and DNMT3B. DNMT3B interacts with a triple helix formed by pRNA and the rDNA. HDAC1, DNMT1, and DNMT3B have been shown to be required for proper DNA methylation of silenced rRNA gene copies, although the catalytic activity of DNMT3B was not required
R-HSA-5250924	B-WICH complex positively regulates rRNA expression.	The B-WICH complex is a large 3 Mdalton complex containing SMARCA5 (SNF2H), BAZ1B (WSTF), ERCC6 (CSB), MYO1C (Nuclear myosin 1c), SF3B1, DEK, MYBBP1A, and DDX21 (Cavellan et al. 2006, Percipalle et al. 2006, Vintermist et al. 2001, Sarshad et al. 2013, Shen et al. 2013, reviewed in Percipalle and Farrants 2006). B-WICH is found at active rRNA genes as well as at 5S rRNA and 7SL RNA genes. B-WICH appears to remodel chromatin and recruit histone acetyltransferases that modify histones to transcriptionally active states
R-HSA-5334118	DNA methylation.	Methylation of cytosine is catalyzed by a family of DNA methyltransferases (DNMTs): DNMT1, DNMT3A, and DNMT3B transfer methyl groups from S-adenosylmethionine to cytosine, producing 5-methylcytosine and homocysteine (reviewed in Klose and Bird 2006, Ooi et al. 2009, Jurkowska et al. 2011, Moore et al. 2013). (DNMT2 appears to methylate RNA rather than DNA.) DNMT1, the first enzyme discovered, preferentially methylates hemimethylated CG motifs that are produced by replication (template strand methylated, synthesized strand unmethylated). Thus it maintains existing methylation through cell division. DNMT3A and DNMT3B catalyze de novo methylation at unmethylated sites that include both CG dinucleotides and non-CG motifs.DNA from adult humans contains about 0.76 to 1.00 mole percent 5-methylcytosine (Ehrlich et al. 1982, reviewed in Klose and Bird 2006, Ooi et al. 2009, Moore et al. 2013). Methylation of DNA occurs at cytosines that are mainly located in CG dinucleotides. CG dinucleotides are unevenly distributed in the genome. Promoter regions tend to have a high CG-content, forming so-called CG-islands (CGIs), while the CG-content in the remaining part of the genome is much lower. CGIs tend to be unmethylated, while the majority of CGs outside CGIs are methylated. Methylation in promoters and first exons tends to repress transcription while methylation in gene bodies (regions of genes downstream of the promoter and first exon) correlates with transcription (reviewed in Ehrlich and Lacey 2013, Kulis et al. 2013). Proteins such as MeCP2 and MBDs specifically bind 5-methylcytosine and may recruit other factors.Mammalian development has two major episodes of genome-wide demethylation and remethylation (reviewed in Zhou 2012, Guibert and Weber 2013, Hackett and Surani 2013, Dean 2014). In mice about 1 day after fertilization the paternal genome is actively demethylated by TET proteins together with thymine DNA glycosylase and the maternal genome is demethylated by passive dilution during replication, however methylation at imprinted sites is maintained. The genome has its lowest methylation level about 3.5 days post-fertilization. Remethylation occurs by 6.5 days post-fertilization. The second demethylation-remethylation event occurs in primordial germ cells of the developing embryo about 12.5 days post-fertilization. DNMT3A and DNMT3B, together with the non-catalytic DNMT3L, play major roles in the remethylation events (reviewed in Chen and Chan 2014). How the methyltransferases are directed to particular regions of the genome remains an area of active research. The mechanisms at each locus may differ in detail but a connection between histone modifications and DNA methylation has been observed (reviewed in Rose and Klose 2014)
R-HSA-5578749	Transcriptional regulation by small RNAs.	Recent evidence indicates that small RNAs participate in transcriptional regulation in addition to post-transcriptional silencing. Components of the RNAi machinery (ARGONAUTE1 (AGO1, EIF2C1), AGO2 (EIF2C2), AGO3 (EIF2C3), AGO4 (EIF2C4), TNRC6A, and DICER) are observed associated with microRNAs (miRNAs) in both the cytosol and the nucleus (Robb et al. 2005, Weinmann et al. 2009, Doyle et al. 2013, Nishi et al. 2013, Gagnon et al. 2014). The AGO:miRNA complexes are imported into the nucleus by IMPORTIN-8 (IPO8, IMP8, RANBP8) and also by an unknown importin while associated with the nuclear shuttling protein TNRC6A (reviewed in Schraivogel and Meister 2014).Within the nucleus, AGO2, TNRC6A, and DICER may associate in a complex (Gagnon et al. 2014). Nuclear AGO1 and AGO2 in complexes with small RNAs are observed to activate transcription (RNA activation, RNAa) or repress transcription (Transcriptional Gene Silencing, TGS) of genes that contain sequences matching the small RNAs (reviewed in Malecova and Morris 2010, Huang and Li 2012, Gagnon and Corey 2012, Huang and Li 2014, Salmanidis et al. 2014, Stroynowska-Czerwinska et al. 2014). TGS is associated with methylation of cytosine in DNA and methylation of histone H3 at lysine-9 and lysine-27 (Castanotto et al. 2005, Suzuki et al. 2005, Kim et al. 2006, Weinberg et al. 2006, Kim et al. 2008, reviewed in Malecova and Morris 2010, Li et al. 2014); RNAa is associated with methylation of histone H3 at lysine-4 (Huang et al. 2012, reviewed in Li et al. 2014). Small RNAs in the nucleus have also been shown to play roles in alternative splicing (Liu et al., 2012, Ameyar-Zazoua et al., 2012) and DNA damage repair (Wei et al., 2012; Francia et al., 2012). Nevertheless, elucidation of the detailed mechanisms of small RNA action requires further research
R-HSA-5617472	Activation of anterior HOX genes in hindbrain development during early embryogenesis.	In mammals, anterior Hox genes may be defined as paralog groups 1 to 4 (Natale et al. 2011), which are involved in development of the hindbrain through sequential expression in the rhombomeres, transient segments of the neural tube that form during development of the hindbrain (reviewed in Alexander et al. 2009, Soshnikova and Duboule 2009, Tumpel et al. 2009, Mallo et al. 2010, Andrey and Duboule 2014). Hox gene activation during mammalian development has been most thoroughly studied in mouse embryos and the results have been extended to human development by in vitro experiments with human embryonal carcinoma cells and human embryonic stem cells.Expression of a typical anterior Hox gene has an anterior boundary located at the junction between two rhombomeres and continues caudally to regulate segmentation and segmental fate in ectoderm, mesoderm, and endoderm. Anterior boundaries of expression of successive Hox paralog groups are generally separated from each other by 2 rhombomeres. For example, HOXB2 is expressed in rhombomere 3 (r3) and caudally while HOXB3 is expressed in r5 and caudally. Exceptions exist, however, as HOXA1, HOXA2, and HOXB1 do not follow the rule and HOXD1 and HOXC4 are not expressed in rhombomeres. Hox genes within a Hox cluster are expressed colinearly: the gene at the 3' end of the cluster is expressed earliest, and hence most anteriorly, then genes 5' are activated sequentially in the same order as they occur in the cluster. Activation of expression occurs epigenetically by loss of polycomb repressive complexes and change of bivalent chromatin to active chromatin through, in part, the actions of trithorax family proteins (reviewed in Soshnikova and Duboule 2009). Hox gene expression initiates in the posterior primitive streak that will contribute to extraembryonic mesoderm. Expression then extends anteriorly into the cells that will become the embryo, where expression is first observed in presumptive lateral plate mesoderm and is transmitted to both paraxial mesoderm and neurectoderm formed by gastrulation along the primitive streak (reviewed in Deschamps et al. 1999, Casaca et al. 2014).Prior to establishment of the rhombomeres, expression of HOXA1 and HOXB1 is initiated near the future site of r3 and caudally by a gradient of retinoic acid (RA). (Mechanisms of retinoic acid signaling are reviewed in Cunningham and Duester 2015.) The RA is generated by the ALDH1A2 (RALDH2) enzyme located in somites flanking the caudal hindbrain and degraded by CYP26 enzymes expressed initially in anterior neural ectoderm of the early gastrula and then throughout most of the hindbrain (reviewed in White and Schilling 2008). HOXA1 with PBX1,2 and MEIS2 directly activate transcription of ALDH1A2 to maintain retinoic acid synthesis in the somitic mesoderm (Vitobello et al. 2011). Differentiation of embryonal carcinoma cells and embryonic stem cells in response to retinoic acid is used to model the process of differentiation in vitro (reviewed in Soprano et al. 2007, Gudas et al. 2013).HOXA1 appears to set the anterior limit of HOXB1 expression (Barrow et al. 2000). HOXB1 initiates expression of EGR2 (KROX20) in presumptive r3. EGR2 then activates HOXA2 expression in r3 and r5 while HOXB1, together with PBX1 and MEIS:PKNOX1 (MEIS:PREP), activates expression of HOXA2 in r4 and caudal rhombomeres. AP-2 transcription factors maintain expression of HOXA2 in neural crest cells (Maconochie et al. 1999). HOXB1 also activates expression of HOXB2 in r3 and caudal rhombomeres. EGR2 negatively regulates HOXB1 so that by the time rhombomeres appear, HOXB1 is restricted to r4 and HOXA1 is no longer detectable (Barrow et al. 2000). EGR2 and MAFB (Kreisler) then activate HOXA3 and HOXB3 in r5 and caudal rhombomeres. Retinoic acid activates HOXA4, HOXB4, and HOXD4 in r7, the final rhombomere. HOX proteins, in turn, activate expression of genes in combination with other factors, notably members of the TALE family of transcription factors (PBX, PREP, and MEIS, reviewed in Schulte and Frank 2014, Rezsohazy et al. 2015). HOX proteins also participate in non-transcriptional interactions (reviewed in Rezsohazy 2014). In zebrafish, Xenopus, and chicken factors such as Meis3, Fgf3, Fgf8, and vHNF regulate anterior hox genes (reviewed in Schulte and Frank 2014), however less is known about the roles of homologous factors in mammals. Mutations in HOXA1 in humans have been observed to cause developmental abnormalities located mostly in the head and neck region (Tischfield et al. 2005, Bosley et al. 2008). A missense mutation in HOXA2 causes microtia, hearing impairment, and partially cleft palate (Alasti et al. 2008). A missense mutation in HOXB1 causes a similar phenotype to the Hoxb1 null mutation in mice: bilateral facial palsy, hearing loss, and strabismus (improper alignment of the eyes) (Webb et al. 2012)
R-HSA-5625886	Activated PKN1 stimulates transcription of AR (androgen receptor) regulated genes KLK2 and KLK3.	PKN1, activated by phosphorylation at threonine T774, binds activated AR (androgen receptor) and promotes transcription from AR-regulated promoters. On one hand, phosphorylated PKN1 promotes the formation of a functional complex of AR with the transcriptional coactivator NCOA2 (TIF2) (Metzger et al. 2003). On the other hand, binding of phosphorylated PKN1, in complex with the activated AR, to androgen-reponsive promoters of KLK2 and KLK3 (PSA) genes, leads to PKN1-mediated histone phosphorylation. PKN1-phosphorylated histones recruit histone demethylases KDM4C (JMJD2C) and KDM1A (LSD1), and the ensuing demethylation of histones associated with the promoter regions of KLK2 and KLK3 genes increases their transcription (Metzger et al. 2005, Metzger et al. 2008)
R-HSA-5689880	Ub-specific processing proteases.	Ub-specific processing proteases (USPs) are the largest of the DUB families with more than 50 members in humans. The USP catalytic domain varies considerably in size and consists of six conserved motifs with N- or C-terminal extensions and insertions occurring between the conserved motifs (Ye et al. 2009). Two highly conserved regions comprise the catalytic triad, the Cys-box (Cys) and His-box (His and Asp/Asn) (Nijman et al. 2005, Ye et al. 2009, Reyes-Turcu & Wilkinson 2009). They recognize their substrates by interactions of the variable regions with the substrate protein directly, or via scaffolds or adapters in multiprotein complexes
R-HSA-5693565	Recruitment and ATM-mediated phosphorylation of repair and signaling proteins at DNA double strand breaks.	Activated ATM phosphorylates a number of proteins involved in the DNA damage checkpoint and DNA repair (Thompson and Schild 2002, Ciccia and Elledge 2010), thereby triggering and coordinating accumulation of DNA DSB repair proteins in nuclear foci known as ionizing radiation-induced foci (IRIF). While IRIFs include chromatin regions kilobases away from the actual DSB site, this Reactome pathway represents simplified foci and events that happen proximal to the DNA DSB ends. In general, proteins localizing to the nuclear foci in response to ATM signaling are cooperatively retained at the DNA DSB site, forming a positive feedback loop and amplifying DNA damage response (Soutoglou and Misteli 2008). Activated ATM phosphorylates the NBN (NBS1) subunit of the MRN complex (MRE11A:RAD50:NBN) (Gatei et al. 2000), as well as the nucleosome histone H2AFX (H2AX) on serine residue S139, producing gamma-H2AFX (gamma-H2AX) containing nucleosomes (Rogakou et al. 1998, Burma et al. 2001). H2AFX is phosphorylated on tyrosine 142 (Y142) under basal conditions (Xiao et al. 2009). After ATM-mediated phosphorylation of H2AFX on S139, tyrosine Y142 has to be dephosphorylated by EYA family phosphatases in order for the DNA repair to proceed and to avoid apoptosis induced by DNA DSBs (Cook et al. 2009). Gamma-H2AFX recruits MDC1 to DNA DSBs (Stucki et al. 2005). After ATM phosphorylates MDC1 (Liu et al. 2012), the MRN complex, gamma-H2AFX nucleosomes, and MDC1 serve as a core of the nuclear focus and a platform for the recruitment of other proteins involved in DNA damage signaling and repair (Lukas et al. 2004, Soutoglou and Misteli 2008). RNF8 ubiquitin ligase binds phosphorylated MDC1 (Kolas et al. 2007) and, in cooperation with HERC2 and RNF168 (Bekker-Jensen et al. 2010, Campbell et al. 2012), ubiquitinates H2AFX (Mailand et al. 2007, Huen et al. 2007, Stewart et al. 2009, Doil et al. 2009) and histone demethylases KDM4A and KDM4B (Mallette et al. 2012). Ubiquitinated gamma-H2AFX recruits UIMC1 (RAP80), promoting the assembly of the BRCA1-A complex at DNA DSBs. The BRCA1-A complex consists of RAP80, FAM175A (Abraxas), BRCA1:BARD1 heterodimer, BRCC3 (BRCC36), BRE (BRCC45) and BABAM1 (MERIT40, NBA1) (Wang et al. 2007, Wang and Elledge 2007) Ubiquitin mediated degradation of KDM4A and KDM4B allows TP53BP1 (53BP1) to associate with histone H4 dimethylated on lysine K21 (H4K20Me2 mark) by WHSC1 at DNA DSB sites (Pei et al. 2011). Once recruited to DNA DSBs, both BRCA1:BARD1 heterodimers and TP53BP1 are phosphorylated by ATM (Cortez et al. 1999, Gatei et al. 2000, Kim et al. 2006, Jowsey et al. 2007), which triggers recruitment and activation of CHEK2 (Chk2, Cds1) (Wang et al. 2002, Wilson and Stern 2008, Melchionna et al. 2000). Depending on the cell cycle stage, BRCA1 and TP53BP1 competitively promote either homology directed repair (HDR) or nonhomologous end joining (NHEJ) of DNA DSBs. HDR through homologous recombination repair (HRR) or single strand annealing (SSA) is promoted by BRCA1 in association with RBBP8 (CtIP), while NHEJ is promoted by TP53BP1 in association with RIF1 (Escribano-Diaz et al. 2013)
R-HSA-5693571	Nonhomologous End-Joining (NHEJ).	The nonhomologous end joining (NHEJ) pathway is initiated in response to the formation of DNA double-strand breaks (DSBs) induced by DNA-damaging agents, such as ionizing radiation. DNA DSBs are recognized by the MRN complex (MRE11A:RAD50:NBN), leading to ATM activation and ATM-dependent recruitment of a number of DNA damage checkpoint and repair proteins to DNA DSB sites (Lee and Paull 2005). The ATM phosphorylated MRN complex, MDC1 and H2AFX-containing nucleosomes (gamma-H2AX) serve as scaffolds for the formation of nuclear foci known as ionizing radiation induced foci (IRIF) (Gatei et al. 2000, Paull et al. 2000, Stewart et al. 2003, Stucki et al. 2005). Ultimately, both BRCA1:BARD1 heterodimers and TP53BP1 (53BP1) are recruited to IRIF (Wang et al. 2007, Pei et al. 2011, Mallette et al. 2012), which is necessary for ATM-mediated CHEK2 activation (Wang et al. 2002, Wilson et al. 2008). In G1 cells, TP53BP1 promotes NHEJ by recruiting RIF1 and PAX1IP, which displaces BRCA1:BARD1 and associated proteins from the DNA DSB site and prevents resection of DNA DSBs needed for homologous recombination repair (HRR) (Escribano-Diaz et al. 2013, Zimmermann et al. 2013, Callen et al. 2013). TP53BP1 also plays an important role in ATM-mediated phosphorylation of DCLRE1C (ARTEMIS) (Riballo et al. 2004, Wang et al. 2014). Ku70:Ku80 heterodimer (also known as the Ku complex or XRCC5:XRCC6) binds DNA DSB ends, competing away the MRN complex and preventing MRN-mediated resection of DNA DSB ends (Walker et al. 2001, Sun et al. 2012). The catalytic subunit of the DNA-dependent protein kinase (DNA-PKcs, PRKDC) is then recruited to DNA-bound Ku to form the DNA-PK holoenzyme. Two DNA-PK complexes, one at each side of the break, bring DNA DSB ends together, joining them in a synaptic complex (Gottlieb 1993, Yoo and Dynan 2000). DNA-PK complex recruits DCLRE1C (ARTEMIS) to DNA DSB ends (Ma et al. 2002). PRKDC-mediated phosphorylation of DCLRE1C, as well as PRKDC autophosphorylation, enables DCLRE1C to trim 3'- and 5'-overhangs at DNA DSBs, preparing them for ligation (Ma et al. 2002, Ma et al. 2005, Niewolik et al. 2006). The binding of inositol phosphate may additionally stimulate the catalytic activity of PRKDC (Hanakahi et al. 2000). Other factors, such as polynucleotide kinase (PNK), TDP1 or TDP2 may remove unligatable damaged nucleotides from 5'- and 3'-ends of the DSB, converting them to ligatable substrates (Inamdar et al. 2002, Gomez-Herreros et al. 2013). DNA ligase 4 (LIG4) in complex with XRCC4 (XRCC4:LIG4) is recruited to ligatable DNA DSB ends together with the XLF (NHEJ1) homodimer and DNA polymerases mu (POLM) and/or lambda (POLL) (McElhinny et al. 2000, Hsu et al. 2002, Malu et al. 2002, Ahnesorg et al. 2006, Mahajan et al. 2002, Lee et al. 2004, Fan and Wu 2004). After POLL and/or POLM fill 1- or 2-nucleotide long single strand gaps at aligned DNA DSB ends, XRCC4:LIG4 performs the ligation of broken DNA strands, thus completing NHEJ. The presence of NHEJ1 homodimer facilitates the ligation step, especially at mismatched DSB ends (Tsai et al. 2007). Depending on other types of DNA damage present at DNA DSBs, NHEJ can result in error-free products, produce dsDNA with microdeletions and/or mismatched bases, or result in translocations (reviewed by Povrik et al. 2012)
R-HSA-5693607	Processing of DNA double-strand break ends.	Homology directed repair (HDR) through homologous recombination (HRR) or single strand annealing (SSA) requires extensive resection of DNA double strand break (DSB) ends (Thompson and Limoli 2003, Ciccia and Elledge 2010). The resection is performed in a two-step process, where the MRN complex (MRE11A:RAD50:NBN) and RBBP8 (CtIP) bound to BRCA1 initiate the resection. This step is regulated by the complex of CDK2 and CCNA (cyclin A), ensuring the initiation of HRR during S and G2 phases of the cell cycle, when sister chromatids are available. The initial resection is also regulated by ATM-mediated phosphorylation of RBBP8 and CHEK2-mediated phosphorylation of BRCA1 (Chen et al. 2008, Yun and Hiom 2009, Buis et al. 2012, Wang et al. 2013, Davies et al. 2015, Parameswaran et al. 2015). After the initial resection, DNA nucleases EXO1 and/or DNA2 perform long-range resection, which is facilitated by DNA helicases BLM or WRN, as well as BRIP1 (BACH1) (Chen et al. 2008, Nimonkar et al. 2011, Sturzenegger et al. 2014, Suhasini et al. 2011). The resulting long 3'-ssDNA overhangs are coated by the RPA heterotrimers (RPA1:RPA2:RPA3), which recruit ATR:ATRIP complexes to DNA DSBs and, in collaboration with RAD17:RFC and RAD9:HUS1:RAD1 complexes, and TOPBP1 and RHNO1, activate ATR signaling. Activated ATR phosphorylates RPA2 and activates CHEK1 (Cotta-Ramusino et al. 2011), both of which are necessary prerequisites for the subsequent steps in HRR and SSA
R-HSA-606279	Deposition of new CENPA-containing nucleosomes at the centromere.	Eukaryotic centromeres are marked by a unique form of histone H3, designated CENPA in humans. In human cells newly synthesized CENPA is deposited in nucleosomes at the centromere during late telophase/early G1 phase of the cell cycle. Once deposited, nucleosomes containing CENPA remain stably associated with the centromere and are partitioned equally to daughter centromeres during S phase. A current model proposes that pre-existing CENPA at the centromere drives recruitment of new CENPA, however this has not been proved.The deposition process requires at least 3 complexes: the Mis18 complex, HJURP complex, and the RSF complex. HJURP binds newly synthesized CENPA-H4 tetramers before deposition and brings them to the centromere for deposition in new CENPA-containing nucleosomes. The exact mechanism of deposition remains unknown
R-HSA-69473	G2/M DNA damage checkpoint.	Throughout the cell cycle, the genome is constantly monitored for damage, resulting either from errors of replication, by-products of metabolism or through extrinsic sources such as ultra-violet or ionizing radiation. The different DNA damage checkpoints act to inhibit or maintain the inhibition of the relevant CDK that will control the next cell cycle transition. The G2 DNA damage checkpoint prevents mitotic entry solely through T14Y15 phosphorylation of Cdc2 (Cdk1). Failure of the G2 DNA damage checkpoint leads to catastrophic attempts to segregate unrepaired chromosomes
R-HSA-73728	RNA Polymerase I Promoter Opening.	The activity of the upstream binding factor (UBF-1) plays an important role in the regulation of rRNA synthesis. Studies reveal that phosphorylation of UBF-1 is required for its interaction with the RNA polymerase I complex, suggesting that phosphorylation of UBF-1 bound to the rDNA promoter during promoter opening modulates the assembly of the transcription initiation complex
R-HSA-73777	RNA Polymerase I Chain Elongation.	The assembly of the initiation complex on the promoter and the transition from a closed to an open complex is then followed by promoter clearance and transcription elongation by RNA Pol I. Unlike the RNA polymerase II system, RNA polymerase I transcription does not require a form of energy such as ATP for initiation and elongation. Regulatory mechanisms operating at both the level of transcription initiation and elongation probably concurrently to adjust the level of rRNA synthesis to the need of the cell
R-HSA-8866654	E3 ubiquitin ligases ubiquitinate target proteins.	E3 ubiquitin ligases catalyze the transfer of an ubiquitin from an E2-ubiquitin conjugate to a target protein. Generally, ubiquitin is transferred via formation of an amide bond to a particular lysine residue of the target protein, but ubiquitylation of cysteine, serine and threonine residues in a few targeted proteins has also been demonstrated (reviewed in McDowell and Philpott 2013, Berndsen and Wolberger 2014). Based on protein homologies, families of E3 ubiquitin ligases have been identified that include RING-type ligases (reviewed in Deshaies et al. 2009, Metzger et al. 2012, Metzger et al. 2014), HECT-type ligases (reviewed in Rotin et al. 2009, Metzger et al. 2012), and RBR-type ligases (reviewed in Dove et al. 2016). A subset of the RING-type ligases participate in CULLIN-RING ligase complexes (CRLs which include SCF complexes, reviewed in Lee and Zhou 2007, Genschik et al. 2013, Skaar et al. 2013, Lee et al. 2014).Some E3-E2 combinations catalyze mono-ubiquitination of the target protein (reviewed in Nakagawa and Nakayama 2015). Other E3-E2 combinations catalyze conjugation of further ubiquitin monomers to the initial ubiquitin, forming polyubiquitin chains. (It may also be possible for some E3-E2 combinations to preassemble polyubiquitin and transfer it as a unit to the target protein.) Ubiquitin contains several lysine (K) residues and a free alpha amino group to which further ubiquitin can be conjugated. Thus different types of polyubiquitin are possible: K11 linked polyubiquitin is observed in endoplasmic reticulum-associated degradation (ERAD), K29 linked polyubiquitin is observed in lysosomal degradation, K48 linked polyubiquitin directs target proteins to the proteasome for degradation, whereas K63 linked polyubiquitin generally acts as a scaffold to recruit other proteins in several cellular processes, notably DNA repair (reviewed in Komander et al. 2009)
R-HSA-8936459	RUNX1 regulates genes involved in megakaryocyte differentiation and platelet function.	In human hematopoietic progenitors, RUNX1 and its partner CBFB are up-regulated at the onset of megakaryocytic differentiation and down-regulated at the onset of erythroid differentiation. The complex of RUNX1 and CBFB cooperates with the transcription factor GATA1 in the transactivation of megakaryocyte-specific genes. In addition, RUNX1 and GATA1 physically interact (Elagib et al. 2003), and this interaction involves the zinc finger domain of GATA1 (Xu et al. 2006). Other components of the RUNX1:CBFB activating complex at megakaryocytic promoters are GATA1 heterodimerization partner, ZFPM1 (FOG1), histone acetyltransferases EP300 (p300) and KAT2B (PCAF), the WDR5-containing histone methyltransferase MLL complex and the arginine methyltransferase PRMT1 (Herglotz et al. 2013). In the absence of PRMT1, the transcriptional repressor complex can form at megakaryocytic promoters, as RUNX1 that is not arginine methylated can bind to SIN3A/SIN3B co-repressors (Zhao et al. 2008). Besides SIN3A/SIN3B, the RUNX1:CBFB repressor complex at megakaryocytic promoters also includes histone deacetylase HDAC1 and histone arginine methyltransferase PRMT6 (Herglotz et al. 2013).Megakaryocytic promoters regulated by the described RUNX1:CBFB activating and repressing complexes include ITGA2B, GP1BA, THBS1 and MIR27A (Herglotz et al. 2013). ITGA2B is only expressed in maturing megakaryocytes and platelets and is involved in platelet aggregation (Block and Poncz 1995). GP1BA is expressed at the cell surface membrane of maturing megakaryocytes and platelets and participates in formation of platelet plugs (Cauwenberghs et al. 2000, Jilma-Stohlawetz et al. 2003, Debili et al. 1990). THBS1 homotrimers contribute to stabilization of the platelet aggregate (Bonnefoy and Hoylaerts 2008). MIR27A is a negative regulator of RUNX1 mRNA translation and may be involved in erythroid/megakaryocytic lineage determination (Ben-Ami et al. 2009).The RUNX1:CBFB complex stimulates transcription of the PF4 gene, encoding a component of platelet alpha granules (Aneja et al. 2011), the NR4A3 gene, associated with the familial platelet disorder (FPD) (Bluteau et al. 2011), the PRKCQ gene, associated with inherited thrombocytopenia (Jalagadugula et al. 2011), the MYL9 gene, involved in thrombopoiesis (Jalagadugula et al. 2010), and the NFE2 gene, a regulator of erythroid and megakaryocytic maturation and differentiation (Wang et al. 2010)
R-HSA-8939236	RUNX1 regulates transcription of genes involved in differentiation of HSCs.	The RUNX1:CBFB complex regulates transcription of the SPI1 (PU.1) gene, involved in differentiation of hematopoietic stem cells (HSCs). RUNX1 recruits histone methyltransferase KMT2A (MLL) to the SPI1 gene locus, leading to generation of the activating H3K4Me3 mark on nucleosomes associated with the SPI1 promoter and the upstream regulatory element (Huang et al. 2011). SPI1 transactivation represses self-renewal and proliferation of HSCs (Fukuchi et al. 2008) and is needed for commitment of HSCs to specific hematopoietic lineages (Imperato et al. 2015).As a component of the TAL1 transcription factor complex, involved in acute T cell lymphoblastic leukemia (T-ALL), RUNX1 can promote growth and inhibit apoptosis of hematopoietic stem cells by stimulating transcription of the MYB gene and possibly the TRIB2 gene (Sanda et al. 2012, Mansour et al. 2014)
R-HSA-9018519	Estrogen-dependent gene expression.	Estrogens mediate their transcriptional effects through interaction with the estrogen receptors, ESR1 (also known as ER alpha) and ESR2 (ER beta). ESR1 and ESR2 share overlapping but distinct functions, with ESR1 playing the primary role in transcriptional activation in most cell types (Hah and Krauss, 2014; Haldosén et al, 2014. The receptors function as ligand-dependent dimers and can activate target genes either through direct binding to an estrogen responsive element (ERE) in the target gene promoter, or indirectly through interaction with another DNA-binding protein such as RUNX1, SP1, AP1 or NF-kappa beta (reviewed in Bai and Gust, 2009; Hah and Krause, 2014). Binding of estrogen receptors to the DNA promotes the assembly of higher order transcriptional complexes containing methyltransferases, histone acetyltransferases and other transcriptional activators, which promote transcription by establishing active chromatin marks and by recruiting general transcription factors and RNA polymerase II. ESR1- and estrogen-dependent recruitment of up to hundreds of coregulators has been demonstrated by varied co-immunoprecipitation and proteomic approaches (Kittler et al, 2013; Mohammed et al, 2013; Foulds et al, 2013; Mohammed et al, 2015; Liu et al, 2014; reviewed in Magnani and Lupien, 2014; Arnal, 2017). In some circumstances, ligand-bound receptors can also promote the assembly of a repression complex at a target gene, and in some cases, heterodimers of ESR1 and ESR2 serve as repressors of ESR1-mediated target gene activation (reviewed in Hah and Kraus, 2014; Arnal et al, 2017). Phosphorylation of the estrogen receptor also modulates its activity, and provides cross-talk between nuclear estrogen-dependent signaling and non-genomic estrogen signaling from the plasma membrane (reviewed in Anbalagan and Rowan, 2015; Halodsèn et al, 2014; Schwartz et al, 2016) A number of recent genome wide studies highlight the breadth of the transcriptional response to estrogen. The number of predicted estrogen-dependent target genes ranges from a couple of hundred (based on microarray studies) to upwards of 10000, based on ChIP-chip or ChIP-seq (Cheung and Kraus, 2010; Kinnis and Kraus, 2008; Lin et al, 2004; Welboren et al, 2009; Ikeda et al, 2015; Lin et al, 2007; Carroll et al, 2006). Many of these predicted sites may not represent transcriptionally productive binding events, however. A study examining ESR1 binding by ChIP-seq in 20 primary breast cancers identified a core of 484 ESR-binding events that were conserved in at least 75% of ER+ tumors, which may represent a more realistic estimate (Ross-Innes et al, 2012). These studies also highlight the long-range effect of estrogen receptor-binding, with distal enhancer or promoter elements regulating the expression of many target genes, often through looping or other higher order chromatin structures (Kittler et al, 2013; reviewed in Dietz and Carroll, 2008; Liu and Cheung, 2014; Magnani and Lupien, 2014). Transcription from a number of estrogen-responsive target genes also appears to be primed by the binding of pioneering transcription factors such as FOXA1, GATA3, PBX1 among others
R-HSA-912446	Meiotic recombination.	Meiotic recombination exchanges segments of duplex DNA between chromosomal homologs, generating genetic diversity (reviewed in Handel and Schimenti 2010, Inagaki et al. 2010, Cohen et al. 2006). There are two forms of recombination: non-crossover (NCO) and crossover (CO). In mammals, the former is required for correct pairing and synapsis of homologous chromosomes, while CO intermediates called chiasmata are required for correct segregation of bivalents.Meiotic recombination is initiated by double-strand breaks created by SPO11, which remains covalently attached to the 5' ends after cleavage. SPO11 is removed by cleavage of single DNA strands adjacent to the covalent linkage. The resulting 5' ends are further resected to produce protruding 3' ends. The single-stranded 3' ends are bound by RAD51 and DMC1, homologs of RecA that catalyze a search for homology between the bound single strand and duplex DNA of the chromosomal homolog. RAD51 and DMC1 then catalyze the invasion of the single strand into the homologous duplex and the formation of a D-loop heteroduplex. Approximately 90% of heteroduplexes are resolved without crossovers (NCO), probably by synthesis-dependent strand annealing.The invasive strand is extended along the homolog and ligated back to its original duplex, creating a double Holliday junction. The mismatch repair proteins MSH4, MSH5 participate in this process, possibly by stabilizing the duplexes. The mismatch repair proteins MLH1 and MLH3 are then recruited to the double Holliday structure and an unidentified resolvase (Mus81? Gen1?) cleaves the junctions to yield a crossover. Crossovers are not randomly distributed: The histone methyltransferase PRDM9 recruits the recombination machinery to genetically determined hotspots in the genome and each incipient crossover somehow inhibits formation of crossovers nearby, a phenomenon called crossover interference. Each chromosome bivalent, including the X-Y body in males, has at least one crossover and this is required for meiosis to proceed correctly
R-HSA-977225	Amyloid fiber formation.	Amyloid is a term used to describe deposits of fibrillar proteins, typically extracellular. The abnormal accumulation of amyloid, amyloidosis, is a term associated with tissue damage caused by amyloid deposition, seen in numerous diseases including neurodegenerative diseases such as Alzheimer's, Parkinson's and Huntington's. Amyloid deposits consist predominantly of amyloid fibrils, rigid, non-branching structures that form ordered assemblies, characteristically with a cross beta-sheet structure where the sheets run parallel to the direction of the fibril (Sawaya et al. 2007). Often the fibril has a left-handed twist (Nelson & Eisenberg 2006). At least 27 human proteins form amyloid fibrils (Sipe et al. 2010). Many of these proteins have non-pathological functions; the trigger that leads to abnormal aggregations differs between proteins and is not well understood but in many cases the peptides are abnormal fragments or mutant forms arising from polymorphisms, suggesting that the initial event may be aggregation of misfolded or unfolded peptides. Early studies of Amyloid-beta assembly led to a widely accepted model that assembly was a nucleation-dependent polymerization reaction (Teplow 1998) but it is now understood to be more complex, with multiple 'off-pathway' events leading to a variety of oligomeric structures in addition to fibrils (Roychaudhuri et al. 2008), though it is unclear whether these intermediate steps are required in vivo. An increasing body of evidence suggests that these oligomeric forms are primarily responsible for the neurotoxic effects of Amyloid-beta (Roychaudhuri et al. 2008), alpha-synuclein (Winner et al. 2011) and tau (Dance & Strobel 2009, Meraz-Rios et al. 2010). Amyloid oligomers are believed to have a common structural motif that is independent of the protein involved and not present in fibrils (Kayed et al. 2003). Conformation dependent, aggregation specific antibodies suggest that there are 3 general classes of amyloid oligomer structures (Glabe 2009) including annular structures which may be responsible for the widely reported membrane permeabilization effect of amyloid oligomers. Toxicity of amyloid oligomers preceeds the appearance of plaques in mouse models (Ferretti et al. 2011). Fibrils are often associated with other molecules, notably heparan sulfate proteoglycans and Serum Amyloid P-component, which are universally associated and seem to stabilize fibrils, possibly by protecting them from degradation

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