Chromatin Modification in Zebrafish Development

CHAPTER 22

Chromatin Modification in Zebrafish Development Jordi Cayuso Mas,* Emily S. No€ el*,y and Elke A. Ober* *

MRC National Institute for Medical Research, The Ridgeway, London, NW7 1AA, UK

y

Present address: Hubrecht Institute and University Medical Centre Utrecht, 3584 CT Utrecht, The Netherlands

Abstract I. Chromatin Modifications A. Histone Acetylation B. DNA and Histone Methylation II. Chromatin Modifications in Development A. Initiation of Gene Transcription During Zygotic Genome Activation B. Histone Modifications Controlling Early Embryo Patterning C. Chromatin Modifications in Neural Development D. Roles of Chromatin Marks in the Development of the Digestive Organs III. Functions of Chromatin Modifications in Tissue Regeneration IV. Outlook Acknowledgments References

Abstract The generation of complex organisms requires that an initial population of cells with identical gene expression profiles can adopt different cell fates during development by progressively diverging transcriptional programs. These programs depend on the binding of transcritional regulators to specific genomic sites, which in turn is controlled by modifications of the chromatin. Chromatin modifications may occur directly upon DNA by methylation of specific nucleotides, or may involve post-translational modification of histones. Local regulation of histone post-translational modifications regionalizes the genome into euchromatic regions, which are more accessible to DNA-binding factors, and condensed heterochromatic regions, inhibiting the binding of such factors. In addition, these modifications may be required in a genome-wide fashion for processes such as DNA replication or METHODS IN CELL BIOLOGY, VOL 104 Copyright 2011, Elsevier Inc. All rights reserved.

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0091-679X/10 $35.00 DOI 10.1016/B978-0-12-374814-0.00022-7

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chromosome condensation. From an embryologist’s point of view chromatin modifications are intensively studied in the context of imprinting and have more recently received increasing attention in understanding the basis of pluripotency and cellular differentiation. Here, we describe recently uncovered roles of chromatin modifications in zebrafish development and regeneration, as well as available resources and commonly used techniques. We provide a general introduction into chromatin modifications and their respective functions with a focus on gene transcription, as well as key aspects of their roles in the early zebrafish embryo, neural development, formation of the digestive system and tissue regeneration.

I. Chromatin Modifications Histones are basic proteins with a net positive charge found in the eukaryotic nucleus and are highly conserved across species. There are two major types of histones: core histones (H2A, H2B, H3, and H4) and linker histones (H1 and H5). Two copies of each core histone form an octamer that interacts with DNA, forming a nucleosome, the basic unit of chromatin. Core histones consist of a globular structure with the N-terminal tails projecting from the main body of the protein. It has been shown that some residues in the tails and in some cases in the globular domains are subject to a multitude of different types of modifications including acetylation, methylation, phosphorylation, ubiquitination, sumoylation, ADP ribosylation, deimination, proline isomerization, and cleavage of the histone tail (Ito, 2007; Kouzarides, 2007; Shahbazian and Grunstein, 2007; Weake and Workman, 2008). Linker histones have been studied to a lesser extent but their involvement in chromatin condensation and the presence of some post-translational modifications have been reported (Wood et al., 2009). Nevertheless, the link between these modifications and their function is largely unknown. Post-translational modifications of histones are dynamic and their roles in regulation of DNA transcription, as well as replication and repair are well established (Groth et al., 2007; Kouzarides, 2007). Three models have been proposed as to how modifications can regulate gene transcription: (i) the charge neutralization model proposes that modifications affecting the basic charge of histones, for example by acetylation of lysines, may interfere with the interactions between adjacent histones, or between histones and DNA, resulting in a relaxed chromatin conformation (Wade et al., 1997). (ii) The histone code model is based on the variety, number, and interdependence of histone modifications. Multiple modifications in the same region, either on the same or neighbouring histone tails, act sequentially or combinatorially to trigger specific cellular downstream events (Strahl and Allis, 2000; Turner, 2000). (iii) The signaling network model establishes parallels between signal transduction mechanisms at the cell membrane and the interpretation of multiple histone modifications in the nucleus. Different chromatin modifications provide redundancy and/or constitute feedback loops conferring bistability, robustness, and adaptability to regulation of transcription (Schreiber and Bernstein, 2002).

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Another layer of complexity arises from the fact that histone modifications can influence each other using different crossregulatory mechanisms (Suganuma and Workman, 2008). Firstly, the presence of an initial histone modification increases the activity of other histone-modifying factors. Secondly, different histone-modifying proteins can be coordinated in the same protein complex, and finally, the presence of certain histone modifications promotes the cleavage of the H3 tail resulting in an irreversible modification of the chromatin. Among the different chromatin modifications, DNA methylation and histone acetylation and methylation are generally the best understood marks and to date represent the most intensively studied modifications in zebrafish.

A. Histone Acetylation All core and linker histones have been found to be acetylated in vivo (de Ruijter et al., 2003; Wood et al., 2009). The levels of histone acetylation depend on the activity of histone acetyltransferases (HATs) and histone deacetylases (HDACs). HATs acetylate lysine residues in histones, resulting in chromatin relaxation and activation of gene expression. HAT activity can be reverted by HDACs, which remove the acetyl groups from histones leading to hypoacetylation, chromatin condensation, and consequently, inhibition of transcription (Fig. 1). Interestingly, in contrast to the classical representation of HATs and HDACs as histone modifiers, not all histone acetylases and deacetylases are localized in the nucleus and non-histone targets have been reported for a subset of these factors, highlighting additional roles and modes of activity for these modifiers (Boyault et al., 2007; Potente et al., 2007). HATs are divided into three different families: GNAT, MYST, and CBP/p300. Members of the GNAT and CBP/p300 families act as transcriptional co-activators, while HATs of the MYST family are associated with diverse functions. The CBP/ p300 group is only found in metazoans (Grozinger and Schreiber, 2002; Yang, 2004). They predominantly acetylate lysines in the N-terminal tail of core histones. On the contrary, acetylation sites in linker histones are located in the globular domain, and have been implicated in mediation of DNA binding (Wood et al., 2009). HDACs are grouped into four major classes based on their similarity to yeast homologs (de Ruijter et al., 2003; Yang and Seto, 2008): Class I – Hdac1, 2, 3, and 8 contain a highly conserved catalytic domain and a more diverse C-terminal tail. Class II – Hdac4-7, 9, and 10; Hdac4, 5, 7, and 9 have a conserved deacetylase domain, while Hdac6 and 10 contain tandem deacetylase domains. This family of deacetylases reportedly has low activity, and it has been suggested that Hdac3 enhances their activity. Notably, Hdac6 localizes to the nucleus and cytoplasm, implicating histone and non-histone targets. Class III – Sir2-like Hdacs; Silent Information Regulators (Sir) or Sirtuins have a NAD+-dependent catalytic domain or sirtuin domain. In mammals, seven homologs have been described, but only SIRT1, SIRT6, and SIRT7 show nuclear localization, suggesting this family also has non-histone targets. Class IV – Hdac11 shares similarities with Class I and

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Class II Hdacs. Despite its high conservation between species little is known about its function. Class I HDACs are widely expressed during the development of all species studied to date. Hdac1 and Hdac2 interact and form the catalytic core of different complexes responsible for transcriptional regulation including Sin3, NuRD, and CoREST, while Hdac3 is found in NcoR/SMRT complexes (Grozinger and Schreiber, 2002). HDACs also interact with more specialized complexes containing CSL (CBF1/Su (H)/Lag2) or Tcf (T-cell factor) transcription factors; these complexes are responsible for regulation of Notch or Wnt signaling (Cunliffe, 2008). Additionally, HDACs can associate with other complexes responsible for coordinating the introduction of different histone modifications (Denis et al., 2009). Thus far, orthologs of the majority of mammalian HDACs have been found in zebrafish, with the exception of Hdac2 and Hdac10. It is important to note that zebrafish Hdac1 is equally similar to mammalian Hdac1 and Hdac2, which is particularly striking as in mammals Hdac2 is a major interacting partner of Hdac1 (Grozinger and Schreiber, 2002). Thus, it is tempting to speculate that in zebrafish Hdac1 might form homodimers in the relevant protein complexes or another protein may carry out the function of a second Hdac protein. Alternatively, the requirement for a second Hdac may have been lost. Functional studies have to date focused predominantly on Hdac1, Hdac3, and Sirt1.

B. DNA and Histone Methylation Regulation of gene transcription during the development of multicellular organisms by chromatin methylation occurs via two mechanisms: either methylation of cytosine nucleotides within DNA or of specific sites of histones (Fig. 1). DNA methylation is associated with a repressive state of gene transcription, either by preventing binding of transcriptional activators or by promoting the recruitment of repressors (Jaenisch and Bird, 2003), while methylation of histones can either promote or repress transcription depending on the modified residue by regulating histone-DNA interaction or via recruitment of transcriptional regulators.

1. DNA Methylation Methylation of CpG sites is found throughout the genome, generally excluding CpG islands, representing DNA stretches of 500 bp with a GC content of 55% that are associated with promoter regions (Takai and Jones, 2002). Methylation is present in most eukaryotic species; however the pattern of methylation with respect to gene bodies is varied (Feng et al., 2010). Bisulfite sequencing was employed to compare DNA methylation in various species, including plants and insects, revealing comparatively higher CpG methylation in vertebrates, with 80% CpG methylation in zebrafish larvae 5 days post fertilization (dpf) compared to 74% in E13.5 mouse embryos and with a slight enrichment in gene bodies in both

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[(Fig._1)TD$IG]

Fig. 1 Schematic representation of DNA and histone modifications and their relation to different transcriptional states. Active transcription is generally associated with unmethylated CpG dinucleotides, as well as H3K4me3, H3K9ac, H3K36me3, H3K56ac, H3K79me3, and polyacetylated H4 in residues 5, 8, 12, and 16. In the active conformation, co-activators recognize the ‘‘activator’’ marks and recruit the RNApol complex, which initiates transcription. Bivalent states have also been suggested in which ‘‘activator’’ and ‘‘repressor’’ modifications co-exist at the same promoter. Extensive CpG methylation of promoters is associated with inactive transcription together with H3K9me3, H3K27me3, and H4K20me3. Methylated residues are highlighted in red, while acetylated residues are indicated in blue. (See color plate.)

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vertebrates (Feng et al., 2010). While in vertebrates CpG methylation equally covers exons as well as intronic and intergenic sequences, in species with overall lower methylation, these sites are enriched in exons, repetitive DNA, and transposons. Methylated promoters are associated with transcriptional repression, whereas unmethylated promoters do not necessarily lead to active gene transcription, thus representing rather a permissive state (Azuara et al., 2006; Sorensen et al., 2010). Concomitantly, promoter regions of developmental regulators remain largely unmethylated during embryogenesis, but are strongly methylated in differentiated cells (Lindeman et al., 2010). Furthermore, changes in methylation of CpG islands associated with specific genes have been observed, for instance, CpG island methylation of no tail is inversely correlated with its expression (Yamakoshi and Shimoda, 2003). Methylation of CpG dinucleotides is mediated by DNA methyltransferases (Dnmt). In vertebrates, Dnmt1, Dnmt2, Dnmt3a, 3b, and 3l have been identified (Goll and Bestor, 2005). There is one zebrafish homolog of Dnmt1 and six of Dnmt3, dnmt3-8, with dnmt3 most similar to mammalian Dnmt3b (Martin et al., 1999; Rai et al., 2007; Shimoda et al., 2005). Vertebrate Dnmt3a and Dnmt3b mediate de novo CpG methylation of unmethylated DNA, whereas Dnmt1 predominantly maintains DNA methylation by recognizing hemi-methylated DNA and modifying the nascent DNA strand. Mutants for ubiquitin-like protein containing PHD and ring finger domains-1 (uhrf1) show a significant reduction of DNA methylation levels at 5 dpf, in agreement with a central role for Uhrf1 in targeting Dnmt1 to the DNA (Feng et al., 2010). Knockdown studies in zebrafish have revealed that individual Dnmt family members perform different methylase activities, for example, Dnmt7 acts more specifically in de novo methylation of the no tail CpG island (Shimoda et al., 2005). Dnmt2 has been implicated in DNA and tRNA methylation; nevertheless its primary activity is still controversial (Goll et al., 2006; Rai et al., 2007). Studies in zebrafish support Dnmt2 activity in the cytosol, as injection of dnmt2 mRNA directed to the cytosol rescued retina and liver differentiation in Dnmt2-depleted embryos, whereas no rescue was observed upon targeting Dnmt2 to the nucleus (Rai et al., 2007). Levels of DNA methylation can be ‘‘passively’’ decreased by failure of CpG methylation on the newly synthesized DNA strand during the replication cycle, whereas ‘‘active’’ demethylation would be replication independent. Experimental evidence supports an active cytosine demethylation activity (Collas, 1998; Niehrs, 2009; Mhanni and McGowan, 2004; Wu and Zhang, 2010) and while no single ‘‘demethylase-enzyme’’ has been identified so far, several multi-step processes have been proposed (Niehrs, 2009; Wu and Zhang, 2010). It is therefore likely that not one single mechanism functions ubiquitously. A recent study combining biochemical techniques examining methylation levels of a 740 bp-fragment of exogenous DNA, with loss-and gain-of-function experiments within zebrafish embryos, proposes an enzymatic process of two consecutive steps involving the cooperation of three proteins (Rai et al., 2008). Demethylation requires the coordinated and combined activity of Activation-induced cytosine deaminase (Aicda)/Apobec leading to a

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replacement of the cytosine with a thymidine intermediate and its subsequent excision by the thymidine DNA glycosylase Mbd4. The efficiency of this process is enhanced by Growth arrest and DNA-damage inducible protein 45a (Gadd45a), which has previously been associated with DNA demethylation (Barreto et al., 2007). It has been suggested that DNA repair subsequently leads to unmethylated cytosines. This represents an attractive model for the demethylation process. Future work will have to determine the mechanistic details and its extent within the embryo and between species.

2. Histone Methylation Methylation of conserved lysines in tails or globular domains of H3 and H4 has been reported in many organisms. Arginine methylation represents an additional histone modification (Di Lorenzo and Bedford, 2011) and several protein arginine methyl transferases have been identified in zebrafish (Hung and Li, 2004). Lysine residues can be mono-, di-, or trimethylated, adding further complexity. Specific histone methylations are associated with euchromatin and active transcription, whereas others are linked with heterochromatic states or silenced transcription. Methylation of histones occurs in nucleosomes spanning upstream regions of genes, core promoters, 50 ends, and 30 ends of ORFs, and the correct positioning of these modifications seems crucial for their function (Li et al., 2007). Furthermore, mono, di-, and trimethylated histones are present in different regions of the same ORF, suggesting that they are regulated independently. Consistently, specific enzymatic requirements have been reported to switch from one state to the other (Li et al., 2007). H3K4, H3K36, and H3K79 methylation are generally connected with active transcription. H3K4me3 accumulates at the 50 end of active genes where it interacts with RNA polymerase II (RNA pol II) and promotes transcription. H3K36me3 is located at the 30 end of active genes and also interacts with RNA pol II, promoting elongation. H3K79me3 has been found in active genes but its function remains unknown. On the other hand, H3K9, H3K27, and H4K20 are associated with transcriptional repression. H3K9 is found in some cases in coding regions of activated genes, raising the possibility that this modification may have different functions depending on whether it is located in promoters or in coding regions (Kouzarides, 2007; Li et al., 2007). Histone lysine methyltransferases (HMT) are often SET (Su(var)3-9, Enhancer of Zeste, Trithorax) domain-containing proteins. In zebrafish, 58 SET-domain genes have been identified, showing different patterns of expression: some of them are deposited as maternal mRNA, while others show either restricted or ubiquitous expression patterns only after zygotic transcription has been initiated (Sun et al., 2008). Functional analysis of these HMTs will be interesting with respect to their residue specificity and their respective functions in embryonic development. In contrast, the first histone demethylases were not reported until recently. They contain either LSD1 or JmjC (Jumonji-containing) domains, and have been shown to be selective for methylation at certain histone positions, as well as for specific states of methylation (mono-, di-, or trimethylated lysines) (Kouzarides, 2007).

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The role of chromatin modifications in zebrafish development has been studied in a variety of processes and tissues, including the early embryo, neural, and digestive system formation, as well as regeneration, which are described in detail below. The growing number of mutations (Table I) identified through various genetic screens together with the development and availability of sophisticated tools and methods (Tables II–IV) have allowed significant progress, providing insights into the regulation of gene transcription.

II. Chromatin Modifications in Development A. Initiation of Gene Transcription During Zygotic Genome Activation The levels of DNA methylation during embryonic development are dynamic and vary between vertebrates. Following fertilization, the mouse genome is globally demethylated at the blastocyst stage and methylation is subsequently restored by the time of implantation (Mayer et al., 2000). In contrast in X. laevis, DNA stays highly methylated after fertilization and only declines progressively during cleavage

Table I Zebrafish lines with mutations in chromatin modifiers Gene

Allele

Type of mutation

Reference (selected)

actl6a (baf53) brpf1

hi1072 t20002 t25114 b943 s872 t24411 t23715 t22925 s436 rw399 b382 hi2628 hi1618 hi2786a s904 hi3020 b719 b999 a8 a50 hi550 hi272

Transgenic insertion Point mutation Point mutation Point mutation Point mutation Point mutation Point mutation Point mutation Point mutation Point mutation Point mutation Transgenic insertion Transgenic insertion Transgenic insertion Point mutation Transgenic insertion Point mutation Point mutation Point mutation Point mutation Transgenic insertion Transgenic insertion

(Gregg et al., 2003) (Laue et al., 2008) (Laue et al., 2008) (Laue et al., 2008) (Anderson et al., 2009) (Stadler et al., 2005) (Stadler et al., 2005) (Stadler et al., 2005) (No€ el et al., 2008) (Yamaguchi et al., 2005) (Nambiar et al., 2007) (Cunliffe, 2004; Golling et al., 2002) (Golling et al., 2002) (Golling et al., 2002) (Anderson et al., 2009) (Golling et al., 2002; Sadler et al., 2007) (Miller et al., 2004) (Miller et al., 2004) (Gregg et al., 2003) (Leung et al., 2008) (Gregg et al., 2003) (Golling et al., 2002; Sadler et al., 2007)

dnmt1 hdac1

hdac3

myst3 smarca4 (brg1) smarca5 (snf2h) uhrf1

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Table II Antibodies recognizing chromatin modifications or modifiers in zebrafish Antigene

Assay

Reference

Dnmt1 Dnmt2 H4ac

IHC WB WB ChIP IHC ChIP ChIP IHC ChIP IHC ChIP IHC IHC ChIP IHC ChIP IHC ChIP

(Anderson et al., 2009; Goll et al., 2009) (Rai et al., 2007) (Lindeman et al., 2009; No€ el et al., 2008; Yamaguchi et al., 2005) (Anelli et al., 2009) (Rai et al., 2010a; Vastenhouw et al., 2010) (Anelli et al., 2009; Lindeman et al., 2010; Vastenhouw et al., 2010; Wardle et al., 2006) (Anelli et al., 2009; Lindeman et al., 2010; Rai et al., 2010b) (Lindeman et al., 2010; Vastenhouw et al., 2010) (Sun et al., 2008) (Sun et al., 2008) (Sun et al., 2008; Vastenhouw et al., 2010)

H4K20me2 Histone H3 H3K4me3 H3K9me3 H3K27me3 H3K36me1 H3K36me2 H3K36me3 H3K9ac Anti-heterochromatin 5meC

(Lindeman et al., 2010) (Anelli et al., 2009) (Rai et al., 2010b)

IHC – immunohistochemistry; WB – western blot; ChIP – chromatin immunoprecipitation.

stages, with the lowest levels at mid blastula transition (MBT) and during gastrulation (Stancheva et al., 2002), suggesting a passive demethylation modus. Studies in zebrafish and other teleosts, using methylation-sensitive restriction enzymes, have arrived at contradictory conclusions regarding the existence of changes in global methylation occurring during early development (Macleod et al., 1999; Martin and McGowan, 1995; Mhanni and McGowan, 2004; Walter et al., 2002). Anti-5-methylcytosine antibody labeling, as well as additional biochemical methods, recently

Table III Bioactive compounds inhibiting chromatin modifiers in zebrafish Compound

Target

Specificity

Reference (selected)

Trichostatin A (TSA) Valproic acid (VPA) 4-(phenylthio)butanoic acid (PBTA) DZnep

Hdac Hdac Hdac

ClassI and II Class I unknown; similar to TSA

(Collas et al., 1999) (Gurvich et al., 2005) (de Groh et al., 2010)

5-azacytidine or 5-aza-20 -deoxycytidine

H3K27me3 and H4K20me3 Dnmt

(Stewart et al., 2009) (Martin et al., 1999)

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410 Table IV Commonly used methods to examine DNA or histone modifications Method

Description

References (selected)

Bisulfite sequencing

Determination of DNA methylation by bisulfite treatment of DNA, which converts unmethylated cytosines into uracil, while 5-methylcytosine residues remain unaffected. Combination of chromatin immunoprecipitation (ChIP) with microarray analysis to study interaction between proteins and DNA Variation of the ChIP protocol to determine different histone modifications co-existing in the same genomic region. Use of restriction enzymes that discriminate methylated and non-methylated targets

(Feng et al., 2010; Lindeman et al., 2010)

ChIP-on-chip

Sequential Chip

Digestion of genomic DNA using modification sensitive enzymes

(Wardle et al., 2006)

(Vastenhouw et al., 2010)

(Macleod et al., 1999; Martin et al., 1999; Rai et al., 2008)

provided novel insights into early methylation state dynamics in the embryo, indicating that global changes occur during early development in zebrafish (MacKay et al., 2007). While sperm DNA is highly methylated, DNA of the early cleavage stage embryo is demethylated and de novo methylation is initiated during late blastula stages, around the onset of MBT. Interfering with DNA methylation using nucleotide analogs 5-azacytidine and 5-aza-2-deoxycytidine prior to MBT causes severe developmental defects in embryos, most notably defective notochord and somite differentiation (Martin et al., 1999). This suggests similar DNA methylation dynamics occur during early zebrafish and mouse development, highlighting the importance of regulation of methylation states during pluripotent embryonic stages and provides insights into mechanisms required for initiation of zygotic genome activation. Fundamental issues in developmental biology include determining the mechanisms by which pluripotency is established and subsequently how lineage specification is initiated. The availability of tools to examine the chromatin landscape has provided insights into these mechanisms with respect to transcriptional control of developmental regulators. A characteristic of early zebrafish development is that maternal gene products support the embryo during the initial rounds of cell divisions, during which the majority of blastomeres are identical and devoid of DNA CpG methylation, H3K4me3, and H3K36me3 (Lindeman et al., 2010; MacKay et al., 2007; Vastenhouw et al., 2010). Zygotic transcription is activated around the 512-cell stage (Kane and Kimmel, 1993), and chromatin immunoprecipitation (ChIP) studies of selected genes revealed that this is accompanied by rapid histone

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modification, including marks associated with active gene transcription such as H3K9ac, H4ac, H3K4me3, and H3K36me3, as well as repressive H3K9me3 and H3K27me3 marks (Lindeman et al., 2010; Vastenhouw et al., 2010). Concomitantly, phosphorylation of RNA pol II and its association with the genome is first detected at the onset of zygotic transcription. ChIP-on-chip (Fig. 2) analysis was used to determine the differential distribution of RNA pol II and trimethylations for about 800 selected genes, revealing that H3K4me3 is detected in promoter regions, H3K36me3 enriched in genes peaking at the 30 end, and H3K27me3 spans promoters as well as entire genes (Fig. 3; Vastenhouw et al., 2010). Following zygotic genome activation, the majority of expressed genes as well as a large fraction of the non-expressed genes exhibit H3K4me3. Slightly later during gastrulation, ChIP analysis of a promoter array for over 11,000 genes showed about 40% H3K4me3 (Wardle et al., 2006), suggesting that H3K4me3 is highly dynamic during early development. A subpopulation of non-expressed genes with H3K4me3 carry in addition the repressive H3K27me3 mark, suggesting a bivalent status (Fig. 1; Vastenhouw et al., 2010). Based on findings in embryonic stem cells, bivalency has been suggested to poise genes for transcription, in that repressive H3K27me3 is dominant over activating H3K4me3 (Azuara et al., 2006; Bernstein et al., 2006). Sequential ChIP analysis (Fig. 2) demonstrated that some of these promoters indeed carried both modifications, instead of a mosaic distribution of one or the other mark in different cells of the embryo. Notably, a group of monovalent H3K4me3-genes was inactive and not associated with RNA pol II. Injection of a Gal4-activatable transgene was used to show that this mark is independent of specific transcriptional activators or association with RNA pol II (Vastenhouw et al., 2010). Thus monovalent H3K4me3 may be important for coordinated and efficient gene activation, as some transcription factors display a higher affinity for trimethylated than for unmethylated H3K4. A study examining additional histone modifications, including H3K9ac, H4ac, H3K4me3, H3K9me3, and H3K27me3, after the onset of zygotic transcription on a small set of expressed genes, revealed modification profiles of similar trends (Lindeman et al., 2010). Either promoters are enriched for activating marks or activating and repressive marks of different combinations, including H3K4me3 and H3K27me3 on the promoter of a gene expressed at low level. Interestingly, one of these genes, a member of the kr€ uppel-like transcription factor family klf4, is expressed in a mosaic pattern in the embryo, indicating that klf4-expressing cells have activating marks while klf4-negative cells carry repressive marks, which could further indicate the origin of lineage specification. Moreover, analysis of the same set of genes in adult muscle tissue or the established zebrafish fibroblast line ZF4 revealed deacetylated promoters with H3K4me3 together with H3K9me3 or H3K27me3 consistent with lack of or low gene expression, while expressed genes maintained acetylation marks (Lindeman et al., 2010). It will be necessary to determine whether these promoters exhibit bivalent marks or whether activating or repressive marks are present in different cells, like for klf4 during early development. Intriguingly, genome-wide experiments in X. tropicalis, including sequential

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[(Fig._2)TD$IG]

Fig. 2 Schematic representation of a ChIP protocol and applications. Protein–DNA complexes are extracted from cells or tissue and cross-linked. DNA is fragmented and incubated with an antibody, for example against a specific chromatin modification and is followed by immunoprecipitation. At this step the collected chromatin can be assayed with another antibody in a second cycle of ChIP (Sequential ChIP). Alternatively, after reversal of the cross-links, the DNA is purified and can be used for: (i) quantitative PCR, using specific primers, (ii) ChIP-on-chip, by respective labeling and hybridization to an array, or (iii) ChIP-Seq, by ligation of generic DNA adaptors followed by sequencing. (See color plate.)

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[(Fig._3)TD$IG]

Fig. 3

Differential distribution of RNA polymerase II, H3K36me3, H3K4me3, and H3K27me3 before and after transition from maternal to zygotic transcription as average density profiles. Enrichment was determined before (256-cells; black line) and after (dome stage/30% epiboly; red line) MBT for 822 genes on the analyzed array and presented as average normalized and smoothed log2ChIP enrichment (MA2C score). Transcription units are represented as metagenes, namely the distance from transcription start site (TSS) to transcription termination site (TTS) is relative, whereas upstream and downstream sequences are shown in absolute distance (bp). Figure derived from Vastenhouw et al. (2010), with permission from the authors. (See color plate.)

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analysis, failed to identify activator and repressor marks occupying the same promoters instead suggesting that these marks solely reflect spatially controlled gene expression (Akkers et al., 2009). Bivalently marked promoters are rare in Drosophila (Schuettengruber et al., 2009), indicating possible species-specific differences in the role of individual chromatin modifications or their combination in governing the activation of the zygotic genome. Unraveling the dynamics of the various chromatin modifications during early developmental stages promises to further our understanding of their roles in the burst of gene transcription at the time of zygotic genome activation. Intriguingly, there is evidence for gene transcription prior to MBT (Londin et al., 2007), pointing toward interesting exceptions to the rule.

B. Histone Modifications Controlling Early Embryo Patterning Complementary to the analysis of chromatin modifications elucidating their role in gene transcription on a broader level, loss-of-function analysis of specific modifying enzymes or their interacting factors have provided insights into their roles for specific patterning processes during early embryogenesis. Forward genetic screens and subsequent positional cloning have identified mutations in the transcriptional corepressor rerea (atrophin-2), reported to interact with nuclear receptors as well as HDAC1 and HDAC2 (Asai et al., 2006; Plaster et al., 2007; Wang et al., 2006; Zoltewicz et al., 2004). rerea embryos display defects in the formation of pharyngeal cartilage, pectoral fins and neural tissues, including the eye and ear, reminiscent of the acerebellar (fgf8) mutant. In rerea-depleted embryos, the expression of several ligands and antagonists of Fgf signaling is altered in a tissue-specific manner. Differences in the reported fgf8 and sef expression in embryos lacking Rerea may be due to the respective alleles examined (Asai et al., 2006; Plaster et al., 2007). Interestingly, rerea and fgf8 act synergistically, as embryos deficient of both factors exhibit severe tail truncations, whereas neither single mutant does (Plaster et al., 2007). Furthermore, treatment of fgf8 mutants with the Hdac inhibitor Trichostatin A (TSA) causes posterior mesoderm defects, phenocopying embryos lacking Rerea and Fgf8 (Plaster et al., 2007). This indicates that transcriptional repression mediated by Hdac1 and Rerea promotes Fgf signaling. Similar interactions underlie the formation of the midbrain-hindbrain boundary and branchial arches. In contrast, Setdb2, a SET domain-containing protein associated with histone H3K9 methyltransferase activity, is required to suppress fgf8 expression in the early embryo (Xu et al., 2010). Analysis of Setdb2-depleted embryos revealed a requirement for H3K9-methylation in controlling fgf8 expression for correct establishment of left – right asymmetry and limiting dorsal organizer tissue. An opposite scenario was recently reported for the regulation of hox gene expression required for the formation of the tail (Lan et al., 2007). Loss-of-function experiments showed that removal of repressive H3K27me3 by Utx, a JmjC-domain

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containing histone demethylase, is central for regulating caudal hox gene expression. Utx is enriched in promoters of actively expressed Hox genes and absent in silenced Hox genes in HeLa cells, with shRNA-mediated inhibition of Utx increasing H3K27me3 levels at HoxD promoters. Morpholino studies showed that Utx1 is required for hox gene regulation in vivo and for development of posterior structures in zebrafish. Intriguingly, posterior trunk and tail tissues failed to maintain their structure resulting in a significantly shortened embryo. In support of this phenotype, expression of ‘‘intermediate’’ hox genes, such as hoxc8a, hoxc11a, and hoxc12b, was mildly reduced in utx1 morphants, whereas expression of more anterior and the most posterior hox genes appeared unaffected (Lan et al., 2007). Overall, these findings suggest that chromatin-modifying enzymes can exert tissue-specific functions, which to some extent is mediated via cofactors such as Rerea imparting specificity to the widely expressed Hdac1.

C. Chromatin Modifications in Neural Development Analysis of the developing nervous system has revealed specific functions for enzymes providing or removing histone marks. Intriguingly, the majority of studies uncovered roles for enzymes controlling histone acetylation and to a significantly lesser extent histone methylation in neural development, encompassing early steps of neurogenesis, hindbrain patterning, as well as differentiation of neuron and glia subtypes. Several reports have identified a requirement for Hdac1 in regulation of CNS and retinal development. hdac1 mutants exhibit a pan-neural phenotype characterized by impaired differentiation of neural tissues and an overall decrease in proliferation (Cunliffe, 2004; Cunliffe and Casaccia-Bonnefil, 2006; Qiu et al., 2009). Conversely, loss of hdac1 in the retina has been associated with increased proliferation, lack of cell differentiation, including lamination defects, and eventually cell death (Stadler et al., 2005; Yamaguchi et al., 2005). These studies have implicated Hdac1 in the positive or negative regulation of several pathways, including Notch, Shh, and Wnt signaling, demonstrating a wide requirement for the intricate control of histone acetylation in neuronal development. Neuroepithelial progenitors give rise to a large array of highly diverse neurons, as well as non-neural glial cells. Analyses of hdac1 mutants show that in the hindbrain, specification and subsequent differentiation of neuronal and glial populations are severely impaired (Cunliffe, 2004; Cunliffe and Casaccia-Bonnefil, 2006). Hdac1 appears to be required for promoting neurogenesis by activating proneural gene expression and to repress the expression of the Notch target and transcriptional repressor her6 (Cunliffe, 2004). Likewise, Hdac1 acts as a Notch antagonist in the eye, with hdac1 mutants displaying increased expression of the Notch effector her4 throughout the retina (Yamaguchi et al., 2005). Similarly, disruption of rhombomere boundaries in the hindbrain, caused by Notch depletion, can be partially rescued by loss of Hdac1 function (Qiu et al., 2009). In the eye, Hdac1 additionally antagonizes Wnt signaling downstream of b-catenin, since (i) expressing a dominant-active form of b-catenin

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can recapitulate the hdac1 loss-of-function phenotype, including increased retinal proliferation, and (ii) excess Hdac1 can suppress b-catenin stimulated proliferation (Yamaguchi et al., 2005). This Hdac1-Wnt antagonism is similar to results from studies of another hdac1 allele, colgate, which exhibit early defects in dorsoventral patterning of the embryo and anteroposterior patterning of the neural plate, reminiscent of Wnt gain-of-function phenotypes (Ignatius et al., 2008). Consistently, colgate phenotypes are enhanced by dominant-negative forms of the Wnt signaling repressors gsk3b and axin, and are compensated by overexpression of Wnt inhibitor dkk1 (Nambiar et al., 2007). Furthermore, treatment of zebrafish embryos with Hdac inhibitors Valproic acid (VPA) and TSA results in teratogenic effects reminiscent of phenotypes caused by increased Wnt signaling (Phiel et al., 2001). In agreement with these studies, cell culture analyses demonstrated that Hdac1 is bound to promoters of Wnt target genes, exerting an inhibitory function, for example, in repressor complexes associated with Groucho and Lef1 (Billin et al., 2000; Brantjes et al., 2001; Chen et al., 1999). In addition, retina differentiation is affected in mutants for the Brahma chromatin remodeling complex, which activates transcription by altering the conformation of the chromatin through interaction with the nucleosomes (Gregg et al., 2003). Analysis of the young mutant, carrying a mutation in SWI/SNF related, matrix associated, actin dependent regulator of chromatin, subfamily a, member 4 (smarca4; previously, brg1), revealed a specific requirement for MAPK-mediated retinal differentiation, while earlier stages of retina development are unaffected. A similar retinal phenotype was observed in actin-like 6a (actl6a) mutants, depleted of another subunit of the same complex, whereas this was not apparent in smarca5 mutants of a different chromatin remodeling complex, highlighting the specificity and importance of each single complex. Levels of histone acetylation are similarly important for CNS patterning. Rhombomere4 (r4) formation requires the activation of a conserved expression cascade of hoxb genes. In tissues actively expressing hoxb genes Choe and colleagues show the presence of Hoxb1, Meis, and Pbx in regions of hoxb promoters (Choe et al., 2009). Importantly, histone acetylation has been implicated in hox gene regulation via interaction with Pbx proteins (Moens and Selleri, 2006), with Pbx and Hoxb1 binding to hox gene regulatory regions executing activator or repressor functions by recruitment of HATs or HDACs. The recruitment of these transcription factors correlates with the acetylation of H4 in Hox gene promoter regions. Overexpression of a dominant-negative form of Meis reduces acetylation of hoxb promoters and Meis, Pbx, and Hoxb1 are no longer detected on the hoxb promoters (Choe et al., 2009). In vitro experiments demonstrate that upon co-transfection of Hoxb1b with Pbx hoxb1a expression was reduced, likely due to the ability of Pbx to recruit Hdac to promoter regions (Saleh et al., 2000), as treatment with TSA can overcome this repression. On the other hand, co-expression of Meis restored hoxb1a expression, suggesting that Meis overcomes Hdac-mediated repression. These and additional interaction studies indicate that Meis competes with Hdacs in binding to Pbx thereby controlling Hdac accessibility at hox promoters, regulating acetylation levels and consequently transcription.

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Patterning of facial motoneurons, which originate in r4, was disrupted in zebrafish embryos containing a mutation in the acetyltransferase monocytic leukemia zinc finger (myst3; previously moz) or hdac1, further highlighting the importance of correct acetylation levels for CNS patterning (Cunliffe, 2004; Miller et al., 2004). Epistasis experiments revealed that hdac1 function is required to maintain the production of branchiomotoneurons in response to Shh signaling, as elevated levels of Shh signaling lead to an excess of these neurons, which is prevented by depletion of Hdac1 in this context (Cunliffe, 2004). The myst3 mutant on the other hand was identified based on a duplication of the first mandibular segment at the expense of the second segment, a homeotic transformation reminiscent of hoxa2b;hoxb2a lossof-function, which is associated with reduced hox gene expression (Miller et al., 2004). The application of TSA partially rescues hox gene expression in the branchial arches and hindbrain, suggesting that restoring histone acetylation rescues cartilage patterning defects. Mosaic experiments elegantly show that while hox genes are expressed widely within the cranium, Myst3-dependent hox expression is cellautonomously required in the cranial neural crest during the post-migratory stages forming the facial skeleton, demonstrating that histone acetylation is required in a specific cell population to control hox gene expression, and henceforth facial skeleton development (Crump et al., 2006). More recent studies have identified a brpf1 mutant, encoding a novel Trithorax group protein that exhibits transformation of anterior structures due to misregulated expression of hox clusters similar to those observed in the myst3 mutant (Laue et al., 2008). Protein interaction between Brpf1 and Myst3 was demonstrated, suggesting that Brpf1 is required for proper chromatin localization and in vivo function of Myst3. Comparatively fewer studies have examined specific requirements for histone methylation in the developing nervous system. The recently identified H3K4 demethylase, SMCX (JARID1C), specifically mediates H3K4me3/2 demethylation in cells, but not H3K4me1 demethylation (Iwase et al., 2007). In zebrafish, kdm5c (smcx) is initially expressed throughout the embryo and later on appears to be restricted mainly to the head. Functional knockdown of Kdm5c during development suggests a possible role in neuronal survival and mosaic analysis points to a cell autonomous requirement (Iwase et al., 2007). These findings are interesting in light of the fact that SMCX has been associated with X-linked mental retardation. Conversely, the zebrafish orthologs of the demethylase KDM7 catalyze the demethylation of mono- and dimethylated H3K9 and H3K27 and while they exhibit a similar expression pattern to kdm5c, they seem to function specifically in the development of the tectum (Tsukada et al., 2010). Morpholino-mediated depletion of kdm7a and kdm7b results in a reduced tectum, accompanied by a cell deathindependent loss of neuronal populations in this area. Importantly, follistatin (fst) expression is reduced in these embryos and Fst depletion results in a phenotype similar to that observed in kdm7a and kdm7b morphants. In mammalian cells, KDM7 associates with the FST promoter close to the transcriptional start site, and KDM7 depletion causes an increase in H3K9me2 and H3K27me2 on the FST promoter and increased FST expression. Overall, these results suggest that

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demethylation of the follistatin promoter is necessary to alleviate repressive methylation marks that control expression levels required for normal tectal development in zebrafish (Tsukada et al., 2010). Given the complexity of the differentiated nervous system, it is conceivable that developmental gene expression is governed by a dynamic and intricate combination of chromatin modifications.

D. Roles of Chromatin Marks in the Development of the Digestive Organs The fact that embryonic development in zebrafish is initially provided for by maternally deposited gene products has proven advantageous for studying the function of chromatin modifications in the development and maintenance of the comparatively late forming digestive system. Hence, a requirement for DNA methylation in the forming digestive system was revealed by the analysis of dandelion (ddn) mutants, encoding for two catalytically inactive dnmt1 alleles, which exhibited severe tissue degeneration of the liver, alimentary canal, and the exocrine pancreas (Anderson et al., 2009). Likely due to maternally provided wild-type Dnmt1, ddn mutants fail to exhibit apparent developmental defects until 84 hpf, a time-point when hepatic and pancreatic differentiation are at an advanced stage. Degeneration of the digestive organs in ddn mutants was shown to be due to p53-dependent and -independent cell death, since loss of p53 in ddn mutants only partially suppresses exocrine pancreas, intestine, and liver degeneration (Anderson et al., 2009). This suggests that loss of cytosine methylation in these organs activates expression of pro-apoptotic genes and/or leads to cellular stress and possibly chromatin changes that are registered as DNA damage. Interestingly, Dnmt1 may execute cell type-specific functions within a single organ, as the endocrine pancreas and pancreatic ducts, which are embedded in exocrine tissue, are unaffected in ddn mutants. Similar observations were made by antisense morpholino oligonucleotide (MO)mediated knockdown of Dnmt1, although with an earlier onset of the phenotype, resulting in embryos with more severe defects (Anderson et al., 2009; Rai et al., 2006). Genome-wide analysis using mass spectrometry revealed a significant reduction of cytosine methylation in Dnmt1 morphants, which was accompanied by a global reduction of H3K9me3 levels. The latter is possibly dependent on cytosine methylation as embryos depleted of the H3K9 methyltransferase Suv39h1 display a similar overall phenotype and reduced H3K9 methylation levels, while global DNA methylation is unaffected (Rai et al., 2006). Altogether, these studies indicate that DNA methylation is essential for organ differentiation and/or maintenance. Concomitantly, DNA hypomethylation has been linked to a failure of intestinal differentiation in embryos with a mutation in the tumor suppressor adenomatous polyposis coli (apc) (Rai et al., 2010b). Genome-wide analysis of DNA methylation by MeDIP (methylated DNA immunoprecipitation; a ChIP variant using an antibody against methylated cytosines, Table II) revealed that promoters of genes implicated

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in early intestinal development are hypomethylated in apchu745 mutants. Rai and colleagues showed that this phenotype is due to increased DNA demethylation activity, as knockdown of DNA demethylation components restored intestinal gene expression in apchu745 mutants, whereas ubiquitous overexpression of dnmt1 failed to rescue. Loss- and gain-of-function strategies combined with quantitative analysis of mRNA levels in whole embryos revealed that Apc suppresses the expression of DNA demethylation enzymes acida, mbd4, and gadd45a indirectly by regulating retinoic acid production. Moreover, Apc regulates retinoic acid production via an array of transcription and chromatin remodeling factors, including Ctbp1, Lsd1, CoRest, Lef1, and Tle3, while retinoic acid signaling in turn negatively regulates Pou5f1 and Cebpb. A similar scenario was observed in human colon adenomas associated with mutations in APC showing an increased expression of DNA demethylation enzymes and factors indicative of an undifferentiated intestine. Altogether, this study demonstrates how the methylation status of genes implicated in early organ fate is controlled and begins to unravel the underlying regulatory cascade and its importance for tissue differentiation and tumor formation. Combining mutant analyses, MO-mediated gene knockdown, and inhibitor treatments produced some of the first insights into the roles of class I Hdacs, in particular hdac1 and hdac3, in vertebrate endodermal organogenesis (Farooq et al., 2008; No€ el et al., 2008). Based on a small liver and pancreas phenotype, a loss-of-function allele of hdac1 was identified in a forward genetic screen for mutants with defects in endodermal organogenesis (No€ el et al., 2008). Its analysis revealed that hdac1 is required for timely specification of hepatic and exocrine pancreatic progenitors at the expense of foregut tissue, suggesting that Hdac1 might serve as a fate switch at the epigenetic level (Fig. 4D, E). Moreover, subsequent hepatic and pancreatic differentiation was defective in hdac1 mutants, increasing in severity over time (Fig. 4F–I; No€ el et al., 2008; No€ el et al., 2010). Importantly, this study also revealed differential cell-autonomous and non-autonomous functions for Hdac1 in liver specification and subsequent differentiation, respectively (No€ el et al., 2008). The use of Hdac inhibitors TSA and VPA confirmed the specific requirements for Hdacs during different stages of organogenesis, with early treatments affecting predominantly specification, and later treatments resulting in liver growth defects (Farooq et al., 2008; No€ el et al., 2008). Consistently, the expression of pescadillo, normally expressed in cycling cells, was initially reduced in the hepatic primordium (Fig. 4J,K). In contrast to the requirement for hdac1 in numerous endodermal tissues and beyond, MO-mediated knockdown indicates a specific function for hdac3 in liver outgrowth and differentiation (Farooq et al., 2008). Interestingly, gdf11 is upregulated in embryos treated with VPA, consistent with the finding that HDAC3 mediated deacetylation of H3 at the Gdf11 promoter regulates Gdf11 expression in mammalian fibroblasts (Zhang et al., 2004). Knockdown of gdf11 partially rescues the hdac3 phenotype, suggesting that in the liver Hdac3 is repressing gdf11 expression (Farooq et al., 2008). In agreement with the finding that Hdac inhibitors cause a more severe hepatic phenotype than individual loss of Hdac1 or Hdac3, double knockdown of both factors shows more severe liver defects (Farooq et al., 2008).

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[(Fig._4)TD$IG]

Fig. 4 Loss of hdac1 leads to tissue-specific defects in zebrafish development. Brightfield pictures of wild-type and hdac1s436 mutants at 48 hpf show several phenotypic defects in hdac1s436 mutants (B), including a slightly shorter axis, fin hypoplasia, and pigmentation defects. Western blot analysis of hyperacetylated histone H4 demonstrates increased acetylation levels in hdac1s436 and hdac1hi1618 mutants, compared to wild-type (C). Hdac activity is required for distinct aspects of endodermal organ development in zebrafish (D-K). Hdac1 is required for allocating the foregut endoderm to its respective fates – hdac1s436 mutants exhibit an expansion of intestinal tissue at the expense of the liver (L) and pancreas (P) (D, E). Subsequently, Hdac1 promotes timely differentiation of hepatic and pancreatic tissues (F–K). Expression of hepatic Tg(fabp10a:dsRed) and pancreatic Tg(elastaseA:GFP), indicative of organ differentiation is delayed in a subset of hdac1s436 mutant embryos (F, G) and not initiated until 4 dpf (H,I). Histone deacetylation promotes tissue-specific expression of cell cycle regulators such as pes, which is reduced in the developing digestive system of hdac1s436 mutants at 48 hpf, consistent with hypoplastic digestive organs at this stage. Conversely, pes expression is increased in the eye, consistent with its hyperproliferation observed of hdac1 mutants, highlighting tissue-specific roles of Hdacs (J, K). (See color plate.)

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Therefore, several Hdacs may act in parallel and/or sequentially in liver formation, as it has been suggested for osteoblast differentiation (Westendorf, 2007). Furthermore, partially redundant functions have, for example, also been observed in tissue-specific deletions of Hdac1 and Hdac2 in the developing myocardium or the B-cell lineage in the mouse, where depletion of both genes causes a more severe developmental defect than loss of either single factor (Montgomery et al., 2007; Yamaguchi et al., 2010). The wider requirement of Hdac1 in endodermal organogenesis and general embryonic development (Burns et al., 2009; Cunliffe, 2004; Nambiar et al., 2007; No€ el et al., 2008; Pillai et al., 2004; Yamaguchi et al., 2005) is in agreement with its described roles in murine development. Mouse embryos lacking Hdac1 die early, before E10.5, due to defects in cell-cycle progression, although initial patterning processes appear to occur undisrupted (Lagger et al., 2002). Hdac3 knock-out mice die due to gastrulation defects before E9.5 (Montgomery et al., 2009). In this case, maternally provided wild-type hdac3 gene products may be sufficient for early embryonic development in zebrafish and allow analysis of later Hdac3 functions in organogenesis of zygotic hdac3 mutants. In light of the classic role of Hdac1 as transcriptional repressor, it is interesting to note that despite the diverse developmental processes examined in hdac1 mutants, surprisingly few genes appeared upregulated upon loss of Hdac1. Intriguingly, there is mounting evidence that Hdacs might in addition ‘‘poise’’ inactive genes for transcription by reducing acetylation levels and preventing RNA pol II from binding by removing acetyl groups added by transiently binding HATs (Wang et al., 2009). Supporting this notion, detailed studies by Harrison and colleagues examining hindbrain neurogenesis show that Hdac1 can act as a positive regulator of gene transcription in addition to its known role as repressor (Harrison et al., 2011).

III. Functions of Chromatin Modifications in Tissue Regeneration Tissue regeneration requires on one hand the maintenance of positional identity and on the other hand the (re-)expression of important developmental regulators, indicating that chromatin modifications are central to this process (Sadler et al., 2007; Stewart et al., 2009; Yakushiji et al., 2009). Following ablation of pancreatic b-cells in larval embryos using the Nitroreductase/Mtz system, de novo formation of b-cells leading to the regeneration of the pancreatic islet appears enhanced in embryos with reduced or absent Dnmt1 function (Anderson et al., 2009). These data suggest the status of DNA methylation is critical for the surrounding pancreatic cells to differentiate into b-cells, in agreement with DNA methylation restricting the capacity of pluripotent cells (Hochedlinger and Plath, 2009). Conversely, liver regeneration following partial hepatectomy in adult zebrafish heterozygous for uhrf1, which recognizes hemi-methylated DNA and is thought to facilitate Dnmt1

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function, was impaired (Sadler et al., 2007). This suggests that the mode of regeneration in the liver versus the endocrine pancreas might differ and/or that Uhrf1 has roles independently from functioning in a complex with Dnmt1. Notably, these findings are consistent with liver outgrowth defects observed in uhrf1 mutants, emphasizing that during regeneration molecular pathways required for the initial formation of a tissue are re-activated (Sadler et al., 2007). Histone methylation was implicated in controlling regenerative potential by a study examining H3K4me3 and H3K27me3 by Sequential ChIP experiments following tail fin amputation (Stewart et al., 2009). Bivalent marks (H3K4me3/ H3K27me3) are enriched on promoters of selected developmental regulators previously implicated in adult fin regeneration, including fgf20a, lef1, and wnt genes, of which a subset loses the repressive H3K27me3 mark upon injury. These findings are in agreement with the concept that the combination of these two modifications poise genes for transcriptional activation (Fig. 1). The loss of repressive H3K27 methylation appears to be at least partially mediated by kdm6bb (previously kdm6b.1), the zebrafish homolog of the H3K27 demethylase JmjD3, which is specifically expressed in the blastema of the adult regenerating fin. Knockdown of Kdm6bb supports its requirement in fin regeneration. Concomitantly, the expression of a single candidate gene, dlx4, is upregulated in these morphants. In regenerating fins, bivalent H3K4me3/H3K27me3 domains change to monovalent H3K4me3 at the promoter of dlx4a (Stewart et al., 2009). The importance of DNA hypomethylation for reactivation of developmental regulators is supported by the finding that the expression of two independent transgenic reporter lines, Tg(h2afv:EFP)nt13 and Tg(XlEef1a1:EGFP)nt12 (previously Tg (H2A.F/Z:EFP)nt and Tg(ef1a:EGFP)nt, respectively), whose activity ceases during larval stages, gets reactivated following caudal fin amputation in the adult (Thummel et al., 2006). Correspondingly, the necessity of DNA hypomethylation for activation of silenced genes was elegantly shown by reactivation of silenced transgenes in a dnmt1 or uhrf1 mutant background (Feng et al., 2010; Goll et al., 2009).

IV. Outlook Generally, it will be necessary to understand the combinatorial activity of the different chromatin modifications, as well as the emerging possibility of RNAs acting as modifiers. Mutant studies have uncovered the requirements for different chromatin remodeling factors in a number of developmental processes ranging from global genome activation to patterning, differentiation and growth of specific tissues. Future work will have to determine whether the observed changes in gene expression are directly governed by the respective factors. Hence, combining detailed phenotypic analysis with characterization of chromatin modifications and occupancy of the respective genomic region promises to further our understanding of the specific functions of chromatin marks in the control of gene expression. This will

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very likely lead to the discovery of ‘‘alternative’’ functions for traditional chromatin modifiers, as it has already been shown for Hdac1 in promoting transcription (Harrison et al., 2011), as well as novel insights into the interactions of the various modifications. Moreover, unraveling the functions and dynamics of chromatin modifications in the context of transcriptional processes during cell fate differentiation (Bai et al., 2010; Keegan et al., 2002; Krishnan et al., 2008) promises to help inelucidating the mechanisms restricting the capacity of pluripotent stem cells. Finally, the suitability of zebrafish for small molecule screens has already been shown to be powerful in uncovering new roles and possible therapeutic targets for compounds against chromatin modifiers in medically relevant fields such as polycystic kidney disease and cancer (Anelli et al., 2009; Cao et al., 2009). With newly emerging technologies and growing understanding of the various functions of chromatin modifications, one might expect exciting progress in this field.

Acknowledgments We would like to thank V. Cunliffe and I. Salecker for critical suggestions on the manuscript. Our work is funded by the Medical Research Council (U117581329).

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